Progress in the Chemistry of Organic Natural Products 118 9783030920296, 9783030920302, 3030920291

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Progress in the Chemistry of Organic Natural Products

A. Douglas Kinghorn · Heinz Falk · Simon Gibbons · Yoshinori Asakawa · Ji-Kai Liu · Verena M. Dirsch Editors

118 Progress in the Chemistry of Organic Natural Products

Progress in the Chemistry of Organic Natural Products Series 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 , School of Pharmacy, University of East Anglia, Norwich, UK 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 Verena M. Dirsch , Department of Pharmaceutical Sciences, University of Vienna, Vienna, Wien, Austria Advisory Editors Giovanni Appendino , Department of Pharmaceutical Sciences, University of Eastern Piedmont, Novara, Italy Roberto G. S. Berlinck , Instituto de Química de São Carlos, Universidade de São Paulo, São Carlos, Brazil Jun’ichi Kobayashi, Graduate School of Pharmaceutical Sciences, Hokkaido University, Sapporo, Japan Agnieszka Ludwiczuk , Department of Pharmacognosy, Medical University of Lublin, Lublin, Poland C. Benjamin Naman , Department of Marine Pharmacy, Ningbo University, Zhejiang, China Rachel Mata , Facultad de Química, Universidad Nacional Autónoma de México, Mexico City, Distrito Federal, Mexico Nicholas H. Oberlies , Department of Chemistry and Biochemistry, University of North Carolina, Greensboro, NC, USA Deniz Tasdemir , Marine Natural Products Chemistry, GEOMAR Helmholtz Centre for Ocean Research, Kiel, Schleswig-Holstein, Germany Dirk Trauner , Department of Chemistry, New York University, New York, NY, USA

Alvaro Viljoen , Department of Pharmaceutical Sciences, Tshwane University of Technology, Pretoria, South Africa Yang Ye , State Key Laboratory of Drug Research and Natural Products Chemistry Department, Shanghai Institute of Materia Medical, 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. All contributions are listed in PubMed.

More information about this series at https://link.springer.com/bookseries/10169

A. Douglas Kinghorn · Heinz Falk · Simon Gibbons · Yoshinori Asakawa · Ji-Kai Liu · Verena M. Dirsch Editors

Progress in the Chemistry of Organic Natural Products Volume 118

With contributions by Alexander J. E. Novak · Dirk Trauner Nicolas Baldovini Charles T. Hanifin · Yuta Kudo · Mari Yotsu-Yamashita Ninh The Son

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

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

Simon Gibbons School of Pharmacy University of East Anglia Norwich, UK

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

Verena M. Dirsch Department of Pharmaceutical Sciences University of Vienna Vienna, Wien, Austria

ISSN 2191-7043 ISSN 2192-4309 (electronic) Progress in the Chemistry of Organic Natural Products ISBN 978-3-030-92029-6 ISBN 978-3-030-92030-2 (eBook) https://doi.org/10.1007/978-3-030-92030-2 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2022 This work is subject to copyright. All rights are solely and exclusively licensed 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, expressed 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

Complex Natural Products Derived from Pyrogallols . . . . . . . . . . . . . . . . . Alexander J. E. Novak and Dirk Trauner

1

The Chemistry of Agarwood Odorants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nicolas Baldovini

47

Chemical Ecology of the North American Newt Genera Taricha and Notophthalmus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Charles T. Hanifin, Yuta Kudo, and Mari Yotsu-Yamashita The Genus Walsura: A Rich Resource of Bioactive Limonoids, Triterpenoids, and Other Types of Compounds . . . . . . . . . . . . . . . . . . . . . . . 131 Ninh The Son

v

Complex Natural Products Derived from Pyrogallols Alexander J. E. Novak and Dirk Trauner

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Oxidation of Pyrogallols and Their Reactivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 A [5+2] Cycloaddition in the Purpurogallin Cascade . . . . . . . . . . . . . . . . . . . . . . . 2.2 The Formation of the Perkin Dimer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 The Hetero-Diels–Alder Dimerization of Pyrogallols . . . . . . . . . . . . . . . . . . . . . . 2.4 Guidelines to Substrate Dependent-Reactivity Trends . . . . . . . . . . . . . . . . . . . . . . 3 Complex Pyrogallol-Derived Natural Products from Epicoccum spp. . . . . . . . . . . . . . . . 3.1 Biomimetic Synthesis of the Calcineurin Phosphatase Inhibitor Dibefurin, a Perkin-Type Dimer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Biomimetic Synthesis of Epicolactone by a Modified Purpurogallin Cascade . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Beetleane A and Epicoane A—Two Perkin-Type Dimer-Derived Natural Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 [5+2] Cycloadditions in the Synthesis of the Merocytochalasans . . . . . . . . . . . . . . . . . . . 4.1 Overview of and Biosynthesis Hypothesis for the Merocytochalasans . . . . . . . . 4.2 Syntheses of the Asperchalasines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Synthesis of the Aspergilasines, Amichalasines, Asperflavipines, and Epicochalasines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Preuisolactone A—Another Racemic Fungal Natural Product . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 3 8 10 10 13 13 16 20 22 23 27 33 38 42

A. J. E. Novak · D. Trauner (B) Department of Chemistry, New York University, 31 Washington Place, New York, NY 10003, USA e-mail: [email protected] A. J. E. Novak e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2022 A. D. Kinghorn, H. Falk, S. Gibbons, Y. Asakawa, J.-K. Liu, V. M. Dirsch (eds.), Progress in the Chemistry of Organic Natural Products 118, https://doi.org/10.1007/978-3-030-92030-2_1

1

2

A. J. E. Novak and D. Trauner

1 Introduction Pyrogallols (1,2,3-trihydroxybenzenes) abound in Nature and are precursors to several important natural products such as the benzotropolones and merocytochalasans [1–3]. The study of the chemistry of pyrogallols and their derivatives has fascinated the chemical community for more than a century and still inspires ongoing research. Pyrogallol (2) itself, which can be derived from gallic acid (1) by decarboxylation (Fig. 1), is one of the oldest photographic developers and can also be used to determine the oxygen concentration of gas mixtures [4]. In this chapter, we will give an overview of historic synthesis studies on pyrogallols and their chemistry focusing on the studies of the purpurogallin cascade and the Perkin dimer. A special focus will be on the [5+2] cycloadditions of hydroxy-o-quinones derived by oxidation of pyrogallols [2, 5, 6]. After a historical overview, several modern examples of pyrogallols and their application in state-of-the-art total synthesis will be reviewed. A strong emphasis in these examples will be made on biomimetic synthesis, a central topic of this contribution. While the benzotropolones derived from pyrogallols are an important natural product family, these will be discussed only briefly, and the main focus will be on reactions of pyrogallols that generate structurally complex, polycyclic molecules with numerous stereocenters. This focus on molecules that feature a multitude of sp3 -centers also means that the rich phenol coupling chemistry of pyrogallols will be excluded, as this reactivity results in aromatic, and comparatively flat molecules. Fig. 1 Synthesis of pyrogallol (2) from gallic acid (1)

OH HO

HO

OH

OH Δ –CO2

HO

OH

O

1 (gallic acid)

2 (pyrogallol)

2 Oxidation of Pyrogallols and Their Reactivity Pyrogallols rapidly oxidize to the corresponding hydroxy-o-quinones in slightly alkaline solutions upon exposure to air. Several alternative oxidants can be used such as potassium nitrosodisulfonate (Frémy’s salt) or K3 [Fe(CN)6 ]. The resulting hydroxyo-quinones can partake in a number of different reaction modes that are summarized in Fig. 2. Notably, hydroxy-o-quinones feature both electrophilic and nucleophilic sites, as indicated. Oxidized pyrogallols such as 3a can react as the 2π-system in pericyclic reactions or ionic reactions such as Michael addition–enolate functionalization cascades. An example of this reactivity is the purpurogallin cascade, which will be examined in the following section. The reactivity pattern of 3b, a nucleophilic attack of the

Complex Natural Products Derived from Pyrogallols

3

most important reactivity sites: O

OH HO

[O]

OH

HO

2 (pyrogallol)

O

3

reactivity observed in the following examples: O HO

3a

HO

O

3b

purpurogallin formation purpurogallin formation Perkin dimer formation = nucelophilic site

O

O

O O

HO

O

HO

O

3d 3c dieneophile hetero-diene hetero-Diels–Alder reaction

= electrophilic site

Fig. 2 Synthesis of hydroxy-o-quinones from pyrogallol (2) and its main reactivity sites

enol, followed by attack onto the most electrophilic carbonyl is observed in the purpurogallin cascade as well. This reactivity pattern of 3b is also encountered in the formation of the Perkin dimer and related compounds, which will also be presented in the following sections. Additionally, hydroxy-o-quinones, like o-quinones feature a hetero-diene motif that is highlighted in 3c. Like o-quinones, hydroxy-o-quinones can therefore undergo inverse electron demand Diels–Alder reactions. Lastly, the enol double bond highlighted in 3d can also react as a dienophile. This overview shows that pyrogallols can engage in a plethora of different reaction modes and highlights their potential synthetic value if their reactivity profile can be well controlled. In addition, it also shows the potential challenges associated with their use in total synthesis as several different modes of reactivity and dimerization can significantly complicate reaction analysis and optimization.

2.1 A [5+2] Cycloaddition in the Purpurogallin Cascade The formation of purpurogallin from the oxidation of pyrogallol is one of the best studied reaction of pyrogallols and the following investigations provided a foundation for modern applications of pyrogallols, including those to be described in this chapter.

2.1.1

The Purpurogallin Cascade—Historical Overview

In 1869, Girard treated pyrogallol (2) with either AgNO3 or KMnO4 under acidic conditions and obtained a red, crystalline compound that he named purpurogallin (Fig. 3) [7]. He proposed the molecular formula C20 H16 O9 for purpurogallin and his

4

A. J. E. Novak and D. Trauner OH

OH HO HO 2 (pyrogallol)

[O]

OH

+

purpurogallin

OH 2 (pyrogallol) OH

OH OH

X

HO C20H16O9 Girard 1869

C11H8O5 Perkin/Steven 1903

HO

OH O 4 (C11H8O5) Dean/Nierenstein 1913 O

OH

O

X

OH

HO

5 (C11H8O5) Willstätter/Heiss 1923

OH OH OH

6 (purpurogallin; C11H8O5) Barltrop/Nicholson 1948

O OH

O

CO2H

HO HO

O OH

AcHN

HO O

CO2H

O O 7 (purpurogallone) Perkin 1903/1912

8 (stipitatic acid) Dewar 1945

9 (colchicine) Dewar 1945 King 1952

Fig. 3 Formation of purpurogallin (6) from pyrogallol (2), and proposed structures for purpurogallin and related structures of interest

report would spark decades of research directed at the elucidation of the structure of purpurogallin and its mechanism of formation. Several modified molecular formulas were proposed for the initially proposed C20 H16 O9 by Girard, but only in 1903 were Perkin and Steven able to report the correct molecular formula, C11 H8 O5 [8]. Perkin and Steven also speculated that purpurogallin might contain a naphthoquinone unit, based on previous findings that distillation of purpurogallin with zinc dust yielded naphthalene [8]. In 1913 Dean and Nierenstein suggested p-quinone methide (4) as the structure for purpurogallin (Fig. 3) [9]. In 1923, Willstätter and Heiss proposed the antiaromatic structure 5 for purpurogallin [10]. While they were unaware of the instability of such a compound, their proposal laid the groundwork for later studies. They suggested that two molecules of pyrogallol would dimerize after oxidation to the corresponding hydroxy-o-quinone. The quinone part was then hypothesized to undergo a benzilic acid ring contraction, followed by decarboxylation to give 5 [10].

Complex Natural Products Derived from Pyrogallols

5

In 1945, Dewar published his groundbreaking assignments of the structures of stipitatic acid (8) and colchicine (9) (the correct regioisomer of colchicine would later be identified by King et al.), proposing that they both contain a cycloheptatrienone [11–13]. Dewar named these structures “tropolones” and the similarity in their reactivities provided support for the correct benzotropolone structure of purpurogallin (6), which was first proposed by Barltrop and Nicholson and shortly thereafter by Haworth, Moore, and Pauson (Fig. 3) [14, 15]. This assignment was supported by the studies of Caunt, Critchlow, Haworth, and coworkers who studied the reactivity, the degradation, and the synthesis of purpurogallin (6) from different starting materials [15–18]. Finally, in 1952 the structure of purpurogallin (6) was assigned unambiguously by X-ray crystallography by Dunitz, who showed that the benzotropolone structure first proposed by Barltrop and Nicholson was indeed correct [19]. With the structure of purpurogallin (6) established, the focus of study shifted to the mechanism by which purpurogallin (6) is derived from pyrogallol (2). In their studies, Willstätter and Heiss had already proposed the oxidation of pyrogallol (2) to the corresponding hydroxy-o-quinone (3) [10]. Two hydroxy-o-quinones were then proposed to undergo dimerization via Michael addition to give 10 followed by tautomerization to 14 (Fig. 4) [10]. It was, however, unclear how a dimer of this type could yield purpurogallin (6), with cleavage of the C-1–C-6 bond by hydrolysis, Friedel–Crafts-type alkylation, and loss of formic acid proposed by Critchlow, Haworth, and Pauson [18]. In 1957, Salfeld reported the formation of the carboxylic acid ester 15 of purpurogallin that he had obtained from the oxidation of pyrogallol 2 in alcoholic solvents (Fig. 4) [20]. These findings suggested the existence of the carboxylic acid of 12 and led him to reconsider the previously proposed mechanisms. According to Salfield, 10 would not undergo tautomerization to 14, but the most electrophilic carbonyl in 10 would be attacked by the enol resulting from the previous Michael addition, yielding tricycle 11 (Fig. 4) [20]. Compound 11 would then tautomerize to pyrogallol 11a and the carbonyl bridge could be opened by either water (R = H in 12) or in alcoholic solvents by the corresponding alcohol leading to the bicyclic 12. In the case of esters, further oxidation and tautomerization would then give rise to 15, as observed by Salfeld. In the case of free carboxylic acids, a decarboxylation is triggered by oxidation to 13, which then affords purpurogallin (6) [20]. The existence of an ester at C-9 as proposed by Salfeld was proven two years later by Horner and Dürckheimer, who also succeeded in trapping the intermediate hydroxy-o-quinones [21–27]. In 1985, Horner and Dürckheimer managed to obtain X-ray quality crystals of the tricyclic 17 (Fig. 4) from the reaction of o-quinone 16 with pyrogallol (2) [28]. The similarity between 17 and the proposed intermediate 11 was striking and these findings led to the general acceptance of the mechanism initially proposed by Salfeld. Additional evidence for this mechanism emerged in 2005 when Nakatsuka and coworkers were able to fully characterize tricycle 20, which they had obtained from the reaction of 18 with 19 [29]. During their studies, Nakatsuka and coworkers were able to show that tricycle 20 upon addition of water and ensuing decarboxylation is converted to a benzotropolone. This provided further support for the conversion of the tricyclic 11 to the bicyclic 12.

6

A. J. E. Novak and D. Trauner H HO

OH

O

O OH

H O

O O

O

OH HO

O

O

O

O O

[O] OH

O OH

O

OH

OH

HO

O

OH

OH 11

10 HO

OH

OH O

O

3

2x 2 (pyrogallol)

O

=

O

O

OH OH OH

[O]

OH

–H OC

ROH OH

CO2R 12

11a O

O

if R = H

OH

OH OH

O

HO

OH OH

OH O 13

OH

HO

OH 6 (purpurogallin)

O

O 6

H O

–CO2

1

O

OH

O OH

O

O

HO

OH O

OH

O OH OH

COOR

OH 14

15 HO

OH

16 O O

OH

18

O

HO

17 OH OH

O

19

20

Fig. 4 The mechanism of the purpurogallin cascade as proposed by Salfeld

While the later steps of the mechanism of the purpurogallin cascade are supported by solid experimental evidence, it should be noted that the initial steps of the cascade are less clear and that much speculation still remains. After more than a century of research it is still unknown whether two hydroxy-o-quinones dimerize or if one hydroxy-o-quinone is attacked by an unoxidized pyrogallol followed by later oxidation. The existence of the corresponding radical intermediates also cannot be excluded. Additionally, even though the mechanism is shown in Fig. 4 as a stepwise Michael addition–aldol biteback cascade, a pericyclic [5+2] cycloaddition mechanism might also be operative.

Complex Natural Products Derived from Pyrogallols

2.1.2

7

The Purpurogallin Cascade in the Synthesis of Benzotropolone Natural Products

Apart from in the formation of purpurogallin itself, the purpurogallin cascade is found widely in the biosynthesis of benzotropolone natural products [30, 31]. Therefore, it has been applied frequently to the total synthesis of natural products from this class. An illustrative example is the biomimetic synthesis of theaflavin by Takino and coworkers. The theaflavins are natural products that arise during the enzymatic oxidation of green to black tea by dimerization of the naturally occurring catechins. The theaflavins are the components responsible for the characteristic color of black tea and often have been associated with antioxidative effects [32]. For this reason, several research groups have investigated the synthesis of the theaflavins via oxidative dimerization [33]. The first biomimetic synthesis of the theaflavins utilizing the purpurogallin cascade was achieved by Takino et al. in 1964 [32]. In their work, epigallocatechin (22) together with epicatechin (21) was treated with either oxidases or inorganic oxidants, yielding theaflavin (23) (Fig. 5). Since then, many more syntheses of different theaflavins have been published utilizing this approach [34, 35]. The purpurogallin cascade can also be found in the biosynthesis of benzotropolone natural products of fungal origin such as aurantricholone (24) and crocipodin (26) [36, 37]. Crocipodin (26) was synthesized by an oxidative dimerization of gallic acid (1) and catechol 25 (Fig. 6) [37]. A fully biomimetic synthesis involving caffeic acid (27) instead of catechol 25 failed, presumably because the electron-poor acid was not readily oxidized. OH

OH HO

O

OH

OH HO

+

O

OH

OH OH

OH

OH 22 (epigallocatechin)

21 (epicatechin)

HO

O

OH

HO

HO

HO

O OHO

OH

OH

HO

23 (theaflavin)

Fig. 5 Synthesis of theaflavin (23)

polyphenol oxidase or other oxidants

8

A. J. E. Novak and D. Trauner OH O

OH

HO O O

O

O Ph

HO Ph

OH 24 (aurantricholone) OH O

OH HO

OH OH

OH

HO

HO

+ CO2H CO2H

Br CO2H

1 (gallic acid)

26 (crocipodin)

25 OH HO

CO2H 27 (caffeic acid)

Fig. 6 Structure of aurantricholone (24) and synthesis of crocipodin (26)

2.2 The Formation of the Perkin Dimer Another mode of oxidative dimerization of pyrogallols initially was discovered through experiments by Perkin and Steven. In 1906, Perkin and Steven did not generate purpurogallin when they treated pyrogallol (2) with isoamyl nitrite and AcOH, but they obtained a colorless, crystalline solid (Fig. 7) [38]. This compound with the molecular formula (C6 H4 O3 )n is now known as the “Perkin dimer” [3]. The Perkin dimer reverts back to pyrogallol (2) when exposed to reducing conditions but, interestingly, forms purpurogallin (6) upon boiling in water. The structural elucidation of the Perkin dimer therefore is also relevant to the studies of the purpurogallin cascade. Despite its colorless nature, Perkin and Steven first proposed structure 3 as that of the Perkin dimer (Fig. 7) [38]. In 1923, however, Willstätter and Heiss [10] recognized its dimeric nature and two different structures (28 and 29) then were proposed by Horner and Dürckheimer [22, 39] and Salfeld [40] in 1959 and 1960, respectively. None of these structures could explain the colorless nature of the Perkin dimer and therefore Teuber and coworkers were the first to propose a tricyclo[5.3.1.12,6]dodecane skeleton (30), and later were able to prove this structure (Fig. 7) [41, 42]. It should be noted that Teuber and coworkers previously had studied the related dimers that they obtained by the oxidation of 31 and 33 with Frémy’s salt (Fig. 8) [41–43].

Complex Natural Products Derived from Pyrogallols OH HO

O

OH

N

9

O

Perkin dimer

AcOH 2 (pyrogallol) O

O HO

O

O

O

OH

O O 28 (C12H8O6) Horner/Dürckheimer 1959

HO 3 (C6H4O3) Perkin/Stevens 1906

OH

O

O

HO

O 29 (C12H8O6) Salfeld 1960

OH O OC O

CO

OH

30 (Perkin dimer; C12H8O6) Teuber 1966

Fig. 7 The discovery of the Perkin dimer and suggested product structures

Fig. 8 Synthesis of Perkin-type dimers by Teuber and coworkers

OH HO

Fremy’s salt, acetone aq. KH2PO4 buffer (9%)

OH O OC

CO

O HO

OH 31

32

OH

Fremy’s salt aq. KH2PO4 buffer

OH

OH O OC

CO

O HO

33

34

Interestingly, related dimers have been reported from phloroglucinols, such as 35, ˇ celsky to give a compound that he which was oxidatively dimerized in 1899 by Ceˇ named cedrone (36) (Fig. 9) [44]. The correct structure of cedrone (36) remained Fig. 9 Synthesis of cedrone ˇ celsky (36) by Ceˇ

OH

HO FeCl3

HO

OH 35

O OC

H2O O

CO OH

36 (cedrone)

10

A. J. E. Novak and D. Trauner OH

OH OH

OH O

O

O

O HO O

PbO2 OH

38

37

O

O OH t-Bu t-Bu O O

HO

O

39 Critchlow/Haworth 1967 Tkachev/Komissarov 2007

OH

HO O O

O

O

t-Bu

40 Salfeld 1960

41 Critchlow/Haworth 1967

Fig. 10 The hetero-Diels–Alder dimerization of sterically hindered pyrogallols

unknown for decades until it was correctly elucidated by Erdtman and Fales in 1969 and 1971 [45, 46].

2.3 The Hetero-Diels–Alder Dimerization of Pyrogallols A third reactivity mode that is common to both pyrogallols and o-quinones was observed during the investigations of pyrogallol oxidations as well: in 1955, Flaig and coworkers treated pyrogallol 37 with PbO2 , oxidizing it to the corresponding sterically hindered hydroxy-o-quinone 38, followed by heating, and observed dimerization (Fig. 10) [47]. The resulting structure was at first misassigned by Salfeld, who proposed the structure 40 based on analogy to the corresponding o-quinone dimers [40]. In 1967, Critchlow and Haworth deduced by IR and NMR spectroscopy that only two carbonyls had to be present in the dimer and thus assigned it the regioisomeric structures 39 and 41 [48]. In 2007, Tkachev and Komissarov were able to obtain X-ray quality crystals and could thus assign unambiguously the structure of the dimer to hetero-Diels–Alder adduct 39 (Fig. 10) [49]. It should be noted that dimers of this type tend to easily decompose into their monomeric species [3].

2.4 Guidelines to Substrate Dependent-Reactivity Trends A historical analysis of studies concerning the products of the oxidative dimerization of pyrogallols, shows that the structural assignments of the products has been

Complex Natural Products Derived from Pyrogallols

11

challenging. Still, certain guidelines for reactions of hydroxy-o-quinones with themselves or o-quinones can be proposed [3]. Normally, the purpurogallin cascade is undergone by substrates that can aromatize like 47, but several “interrupted” purpurogallin cascade products have also been reported (such as 17 shown in Fig. 4) and more will be introduced below. In general, dimerization via the purpurogallin cascade involves 4-substituted pyrogallols such as 47, but also differently substituted pyrogallols can react if the substituents can be cleaved under the reaction conditions, e.g. by decarboxylation (Fig. 11) [3]. The formation of the Perkin dimer is one of the competing pathways to pyrogallol oxidative dimerization via the purpurogallin cascade. The formation of tricyclic Perkin-type dimer structures like 44 or 45 normally occurs for substrates that cannot aromatize (such as 50 and 51, Fig. 11) [3]. In case the substrates are too sterically demanding and unable to aromatize (52), pyrogallols tend to oxidatively dimerize to structures like 46 via a hetero-Diels–Alder reaction. This reaction places the sterically demanding substituents further away from each other than in the formation of the Perkin dimer. The driving force for this reaction is mainly the rearomatization of one of the quinone partners [3]. Thus, it may be seen that slight modifications to substrates can lead to the desired selectivity. This becomes especially apparent when heterodimerizations are considered. In principle, the purpurogallin cascade only requires one reaction partner to be a hydroxy-o-quinone like 3b with the critical 1,3- or 1,5-relationship between nucleophilic and electrophilic sites. The other coupling partner could be another

HO

O

OH

HO

O

OH

OC

R

R OH

R R

R 42 purpurogallin derivatives

R 43

R

OH OH

R

R R 49

47

OH RL

O

RL

RL

O HO O

CO

RS

RS OH O 45

OH OH

R

OHRS RS OC

RS

OH

O

OH R 48

R

RS

CO

44 Perkin-type dimer derivatives

if partner is an o-quinone

or

O

OH O

R

O OH

R

R

OH

OH

OH

O

OH

OH

OH 50

RS

OH

or

RS

OH RS 51

OH RL

RL

OH

O

46 hetero-Diels–Alder dimer

OH RL 52

Fig. 11 Substitution patterns for different reaction modes in the oxidative dimerization of pyrogallols and some catechols

12

A. J. E. Novak and D. Trauner

hydroxy-o-quinone like 3a or also be any o-quinone like 53 as just a 1,2 relationship between nucleophilic and electrophilic sites is required (Fig. 12) [3]. In contrast to this, Perkin-type dimers can only form between two hydroxy-oquinones like 3b or other benzene triols such as phloroglucinol 35 (Figs. 9 and 12). This is due to the fact that both partners need to contain a 1,3- or 1,5-relationship between the nucleophilic and electrophilic positions [3]. In the following sections, it will be shown that in order to suppress a competing Perkin-type dimer formation in a purpurogallin cascade, a hydroxy-o-quinone should be combined with an oquinone. The hetero-Diels–Alder reaction could, in principle, take place between any o-quinone 53a and any enol such as 3d. In the following sections, several more modern examples of the oxidative dimerization of pyrogallols in total synthesis will be encountered and the different reactivity modes will be discussed. Fig. 12 Minimal reactivity profiles for the oxidative dimerization of pyrogallols and catechols

purpurogallin cascade O

O O

O O

OH

O

OH 3b characteristic of:

3a

hydroxy-oquinone with

O

HO

53 any o-quinone

hydroxy-oquinone or

OH

O

HO

OH OH

OH OH 54

6 (purpurogallin)

Perkin dimer formation O

hetero-Diels–Alder reaction

O

O OH

HO HO

O 3b

O 3b

hydroxyo-quinone

hydroxy-oquinone or

O

OH OC

O O 3d

O 53a any o-quinone

enol

O CO

OH O 30 (Perkin dimer) = nucleophilic site

O HO O 55 = electrophilic site

O

Complex Natural Products Derived from Pyrogallols

13

3 Complex Pyrogallol-Derived Natural Products from Epicoccum spp. 3.1 Biomimetic Synthesis of the Calcineurin Phosphatase Inhibitor Dibefurin, a Perkin-Type Dimer Dibefurin (56) is a natural product that was isolated from a Basidiomycota fungal culture at Abbott Laboratories [50]. Its structure was elucidated using a combination of spectroscopic methods and single-crystal X-ray analysis and is shown in Fig. 13. Dibefurin contains five rings fused into each other and it was suggested by the isolation team that dibefurin (56) is a dimer that might be derived biosynthetically from the pyrogallol flavipin (57) [50]. Structurally, dibefurin features a central cyclohexane ring that has two three-carbon atom bridges in a 1,3-di-axial relationship, forming a tricyclo[5.3.1.12,6 ]dodecane skeleton to which two tetrahydrofuran rings are attached (Fig. 13). A special feature of dibefurin’s structure is that it contains a center of inversion and no other symmetry elements. This makes dibefurin (56) C i -symmetric, which is a very rare property for natural products [51]. In addition to its unusual structural features that make it an interesting target for total synthesis, dibefurin (56) was also shown to possess significant biological activity. Dibefurin was isolated using bioactivity-guided assays to search for novel potential immunosuppressants and was shown to inhibit calcineurin phosphatase (IC 50 = 44 μM), an enzyme involved various immune responses [50]. Inhibition of calcineurin phosphatase can be of importance, since indirect inhibitors of this enzyme, such as cyclosporin A or FK-506, are used widely to prevent organ rejection after transplantation. In contrast to these well-established immunosuppressants, dibefurin does not inhibit calcineurin phosphatase through the formation of ternary immunophilin complexes but was reported to directly inhibit the enzyme [51]. The Trauner group realized that dibefurin (56) bears a resemblance to the Perkin dimer and thus reasoned that dibefurin might be derived from a dimerization of two molecules of epicoccine (61) [51]. At the same time, they studied the biosynthesis Fig. 13 The structure of dibefurin and its structural features

O O

O

HO

O

OH

O

O

O

O

O

HO

tricyclo[5.3.1.1.2,6]dodecane

O O

HO

O

O

56 (dibefurin)

O

OH

O

OH

OH

O

O

HO HO

O monomeric unit

57 (flavipin)

O O

14

A. J. E. Novak and D. Trauner Cl

O O

O OH

formalin, HCl

O O

(94%)

O

Zn, THF/ aq. KH2PO4 (85%)

O O

O 59

58 (eudesmic acid) O O O

1. DIBAL–H 2. Et3SiH, TFA 3. BBr3 (46% over 3 steps)

O

HO O HO

O

OH

60

61 (epicoccine)

Fig. 14 Synthesis of epicoccine (61)

origin of and pursued the biomimetic synthesis of another fungal metabolite epicolactone (64) (see next section). They reasoned that epicolactone and dibefurin might be related biosynthetically, with dibefurin resulting from a homodimerization of two molecules of epicoccine (61) and epicolactone (64) stemming from the heterodimerization of epicoccine (61) with epicoccone B (65) [51]. In order to untangle and study this proposed biosynthetic relationship, they set out to achieve both the biomimetic synthesis of dibefurin (56), and epicolactone (64). Their synthesis efforts began with the development of a reliable and scalable synthesis of epicoccine (61). First, eudesmic acid (58) was converted to isobenzofuranone 59 utilizing chloromethylation conditions (Fig. 14) [51]. Dechlorination using zinc dust yielded 60 that was then converted to epicoccine (61) by exhaustive reduction using DIBAL–H, followed by acid-mediated etherification and global deprotection with BBr3 . Through this sequence, epicoccine (61) was accessed in multi-gram quantities and this synthesis sequence would be utilized in several later syntheses as well [52–54]. The Trauner group then turned their attention to the homodimerization of epicoccine (61) to give dibefurin (56). Even though structurally similar compounds such as the Perkin dimer and cedrone had been accessed from their corresponding pyrogallols and phloroglucinol derivatives, treatment of epicoccine (61) with o-chloranil or 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) failed to yield any dibefurin (56) [51]. Treatment of epicoccine with excess Frémy’s salt gave dibefurin in modest yield and transition metal salts either resulted in decomposition or low yield. Ultimately, it was found that treatment of epicoccine (61) with K3 [Fe(CN)6 ] in slightly alkaline buffer resulted in the formation of dibefurin (56) in good yield (Fig. 15) [51]. Purification of dibefurin, however, was complicated by both its insolubility in most solvents and the formation of an isomer of dibefurin, the C 1 -symmetric compound 63. After repeated trituration with THF, dibefurin (56) was isolated in 49% yield, thereby completing the biomimetic synthesis of 56. In addition, it was also found that molecular oxygen could be used as the oxidant, because treatment of epicoccine

Complex Natural Products Derived from Pyrogallols HO 2x

OH

15

OH

K3[Fe(CN)6] NaHCO3

HO

O

O O

MeCN/H2O

O

[5+5]

O O

HO

O 61 (epicoccine)

62 O O

O O

O

HO

O

OH O

O

+

HO

O 56 (dibefurin) (49%; 62% by NMR)

OH

O

O

O 63 (36% by NMR)

Fig. 15 Biomimetic synthesis of dibefurin (56)

(61) with catalytic amounts of iron salt under an oxygen atmosphere also yielded dibefurin (56). These results show that dibefurin could be formed spontaneously naturally without any enzymatic assistance [51]. With the completion of the biomimetic synthesis of dibefurin (56), Trauner et al. embarked on studying the biological activity of 56 and to also examine its potential existence in Epicoccum spp. (e.g. Epicoccum nigrum, Plate 1) Therefore, an interleukin-2 reporter gene assay was conducted at Novartis Pharma AG in Basel and crystallographic studies were attempted in collaboration with the Peti group Plate 1 Colony of Epicoccum nigrum

16

A. J. E. Novak and D. Trauner

(University of Arizona). These studies showed that synthetic 56 is not an immunosuppressant and was inactive in the assay [3]. The initial results of the isolation team therefore might have been based on non-specific effects rather than actual inhibition of calcineurin phosphatase by dibefurin (56). As had been previously mentioned, it was believed that dibefurin (56) and epicolactone (64) might be biosynthetically related. Dibefurin (56) should therefore also exist in the cellular extract of an Epicoccum sp. Analysis of a cellular extract of Epicoccum sp. revealed the existence of a compound with the same retention time and fragmentation pattern as dibefurin (56) by LCMS analysis [3]. These results suggest that 56 should indeed exist in Epicoccum spp. and provide support for a biosynthetic relationship between dibefurin (56) and epicolactone (64).

3.2 Biomimetic Synthesis of Epicolactone by a Modified Purpurogallin Cascade Epicolactone (64) is a compact and highly complex natural product that has been isolated by two independent groups from Epicoccum spp., endophytic fungi that colonize both the sugarcane and the cocoa tree [55, 56]. Epicolactone displays both antimicrobial and antifungal activity and it has been suggested that epicolactone plays a role in protecting the host plant from pathogens [56]. The structure of epicolactone (64) (Fig. 16) was elucidated by spectroscopic means and by X-ray crystallography. Epicolactone (64) is a pentacyclic natural product with an unprecedented carbon skeleton, featuring five stereogenic centers that are all adjacent to each other, with three of them being quaternary (Fig. 16) [52]. Despite its highly complex structure, epicolactone (64) was isolated as a racemate, raising the question of how it is being formed in Nature [5]. It has been suggested that epicolactone is a polyketide, but no detailed biosynthesis proposal had been put forth by the chemists who isolated it. Fig. 16 The structure of epicolactone (64) and its similarity to purpurogallin (6) when flattened

O

O HO

HO

OH

HO

HO O

O

O

O decalin ring system

OH O OH

O

O

O

O 64 (epicolactone)

HO

OH

O

O

O

≈

HO

O

OH OH OH

O 64

6 (purpurogallin)

Complex Natural Products Derived from Pyrogallols

17

Trauner and coworkers realized that when epicolactone (64) was flattened to highlight the 6/7 ring system, it bears a resemblance to purpurogallin (19), the product of the previously discussed purpurogallin cascade (Fig. 16) [51, 52]. Therefore, they proposed that epicolactone (64) might be formed by an oxidative [5+2] cycloaddition between epicoccine (61) and epicoccone B (65), which bears a resemblance to the purpurogallin cascade [51]. The corresponding hydroxy-o-quinones 62 and 66 could undergo a [5+2] cycloaddition to give 67 (Fig. 17). The lactone ring of 67 was then proposed to undergo hydrolysis, giving β-keto acid 68, which should then be prone to decarboxylation to yield 69. This would give the bridgehead carbonyl intermediate 69 that could be attacked by the free, primary alcohol and then fragment to give the hydroxy dienol 70 in a formal retro-Dieckmann reaction. The corresponding intermediate in the purpurogallin cascade, compound 12 (see Fig. 4), is prone to decarboxylation and aromatization, but hydroxy dienol 70 cannot aromatize due to the presence of a quaternary carbon. Instead, 70 undergoes a vinylogous aldol addition, yielding epicolactone (64) [51]. Unfortunately, when these authors exposed epicoccine (61) together with epicoccone B (65) to oxidative conditions, they did not observe formation of epicolactone (64). Instead, they either observed decomposition or formation of dibefurin (56) the homodimerization product of epicoccine (61) (Fig. 15) [3, 51, 52]. When they exposed epicoccine (61) to o-quinone 71, they did not observe the formation of 64

HO

OH OH

O

OH

O

OH

+

OH

65 (epicoccone B)

COOH O O

O

62

O

O OH

O

OH

O

+

O

O

O

O O

O

O

OAc O

(10%)

O O

O O

O

O

O

62a

O

AcO

then Ac2O

O

OH

O

O

64 (epicolactone)

70

HO

2.0 eq 71

HO O

O

O

61

OH

O

O

69

OH

HO

O

OH

68

O

O

O

OH

OH

O

H HO

O

–CO2

67

66

OH O

O

OH –H

HO

OH

OH

O

H2O

O

61 (epicoccine)

O

O

O

O

OH

O

[5+2]

O

O

O

O

O

O

O

O

[O]

O OH O

OH

72

73

Fig. 17 The initially proposed biosynthesis of epicolactone by Trauner and coworkers and isolation of a hetero-Diels–Alder product

18

A. J. E. Novak and D. Trauner

either, but instead saw the formation of an unstable dimer, which could be isolated as acetate 73 (Fig. 17). The structure of 73 was elucidated using X-ray crystallography that showed it to be a hetero-Diels–Alder adduct [51]. With these results in hand, it was decided to pursue an alternative route. As a refined proposal, the investigators hypothesized that epicolactone (64) could be derived through an oxidative [5+2] cycloaddition cascade from epicoccine (61) and pyrogallol 74 [3, 52]. Epicoccine (61) and pyrogallol 74, itself possibly derived from epicoccone B (65), could be oxidized to the corresponding hydroxy-o-quinones 62 and 76, which could then undergo a [5+2] cycloaddition (Fig. 18). This would give the bridgehead carbonyl intermediate 69, which, as in the above proposal (Fig. 17), could be attacked by the free, primary alcohol and then fragment to give the hydroxy dienol 70. Hydroxy dienol 70 would then undergo a vinylogous aldol addition, yielding epicolactone (64) [3, 52]. When the two starting materials epicoccine (61) and catechol 75 were subjected to a slightly alkaline solution of K3 [Fe(CN)6 ], epicolactone methyl ether 80 and its regioisomer were obtained as the two products in 42% and 16% yield, respectively (Fig. 18) [52]. In contrast to 80, its regioisomer is most likely derived from the [5+2] cycloaddition of 62b, a tautomer of 62 with 77. The methyl ether could then readily be removed by treatment with MgI2 and quinoline, yielding epicolactone (64) in 75% yield, giving support to the refined biosynthesis proposal. The regioisomer could be deprotected using AlCl3 , giving 82 that was named “isoepicolactone” and proposed to also exist in Nature [52]. Indeed, Isoepicolactone (82) was isolated in 2020 from Epicoccum nigrum (Plate 1) providing another example of successful “natural product anticipation” through biomimetic synthesis [57]. Recently, epicolactone A (86) an analog of epicolactone (64) bearing an extra methyl group was isolated from Epicoccum nigrum in 2020 [58]. Presumably, epicolactone A (86) arises from the [5+2] cycloaddition, retro-Dieckmann, vinylogous aldol cascade of 62 with 84 as shown in Fig. 18. In addition to their completion of the synthesis of epicolactone (64), which sheds light on the biosynthetic origin of this fascinating natural product, Trauner and coworkers set out to study the mechanism of their proposed cascade. To this end, they utilized protected benzyl alcohol 87, which they treated together with epicoccine (61) under anhydrous conditions with Ag2 O (Fig. 18). They reasoned that the protection of the primary alcohol in 87 should suppress the fragmentation of the carbonylbridged intermediate. In accordance with their hypothesis, they isolated 89 as a single diastereomer that they were able to characterize via X-ray crystallography [3, 52]. Compound 89 presumably arose from 88, the initial product of the [5+2] cycloaddition. These findings suggest that the [5+2] cycloaddition is highly diastereoselective and provides more support for the intermediacy of carbonyl-bridged intermediates in the purpurogallin and related cascades.

Complex Natural Products Derived from Pyrogallols

HO

OH

O

OH

K3[Fe(CN)6] NaHCO3

OH HO

19 OH O

O O

O

+

OR 62

R = H: 74 R = Me: 75

OH O R = H: 69 R = Me: 78

HO R = H: 76 R = Me: 77

O

O O

HO

O O

O

(42%)

O

(75%)

O

O

O

O

O

O OH 81

HO 77

O

61 +

OH

O

HO

O

O [O]

O

(15%) over 2 steps

62

83

O 84

HO

O OH

+

Ag2O, dioxane

O HO

87

OH

O HO

OH 85

O

HO

O

OH

O O O O

OAc 88

O

O 86 (epicolactone A)

O

OMe AcO

O 82 (isoepicolactone)

O

HO

OH

O O

O

OH

O

HO

O

[5+2]

OH

OH HO

OH

O

O O

O

OH

O

HO

O O

[5+2]

62b

OH

O

O HO

O

MeCN/H2O

HO O 64 (epicolactone)

OH

61 + 75

OH

O

O

O 80

R = H: 70 R = Me: 79

K3[Fe(CN)6] NaHCO3

MgI2, quinoline THF, 60°C

HO

O

O

HO

O

O

OR

61

O O

OR

O

H HO

O

[5+2]

MeCN/H2O

O HO

O 61(epicoccine)

HO

O (50%)

O O O

OAc 89

Fig. 18 The biomimetic synthesis of epicolactone (64), isoepicolactone (82), the possible biogenesis of epicolactone A (86), and isolation of bridgehead intermediate 89

20

A. J. E. Novak and D. Trauner

3.3 Beetleane A and Epicoane A—Two Perkin-Type Dimer-Derived Natural Products In their search for more complex pyrogallol dimers from Epicoccum nigrum (Plate 1), Yang and Kong and coworkers isolated two interesting structures that they named epicoane A and beetleane A [60]. The structures of these natural products were elucidated by spectroscopic studies and by X-ray crystallography (epicoane A), and are shown in Fig. 19. Epicoane A (90) consists of a complex and unique 6/5/5/5/6/6/5 heptacyclic ring system and beetleane A (91) features a [5.5.5.6]trioxafenestrane core that is fused to a cycloheptane [60]. Yang and Kong proposed that beetleane A (91) could be derived biosynthetically from epicoane A (90) and that both natural products would thus be products of a Perkin-type dimer formation [60]. Interestingly, both natural products have been isolated as enantiomerically enriched compounds, raising the question how enantiocontrol over their formation is achieved in Nature. Yang and Kong proposed that the natural products are formed by an initial oxidative dimerization between two units of epicoccine (61) to give compound 92 as a racemate (Fig. 20) [60]. It is important to note here that the proposed dimerization mode of 62 gives the C 2 -symmetric product 92, which has neither been observed by Trauner nor Tang in their studies [3, 51, 53]. Compound 92 was then proposed to undergo functionalization with a C3 -unit. As shown in Fig. 20, this transformation would consume two equivalents of the acetone radical and rely on the persistence of a radical in 92 after the initial Hatom abstraction, which would be a highly unusual transformation. Ketone 93 could then be reduced, giving alcohol 94 that is proposed to form a six-membered ring by attack onto the central ketone, which can then form a hemiacetal to give epicoane A (90) [60]. ent-Epicoane A (ent-90) is then proposed to react further by collapse of the hemiacetal moiety and concomitant cleavage of a C–C bond to give 95 that can also be represented as 95a. The enolate 95a is then proposed to undergo a Michael addition to yield beetleane A (91) [60]. While this is an intriguing proposal for a biosynthesis, several steps raise further questions: as already mentioned, the dimerization of epicoccine (61) has not been found to yield the proposed C 2 -symmetric product 92 by two different research groups, but only the C i - or C 1 -symmetric products have been obtained so far (Fig. 15) [3, 51, 53]. Additionally, the further functionalization of 92 with a C3 unit is highly unlikely to occur as shown. Alternatively, one could imagine the initial

OHO O

O O HO O

O O

O O OH O O OH

proposed monomeric unit

OHO O

OH

O

OH

O OO

O O OH

90 (epicoane A)

Fig. 19 The structures of epicoane A (90) and beetleane A (91)

91 (beetleane A)

Complex Natural Products Derived from Pyrogallols

21

O HO

OH

2 x HO

O

OH

O

OHO

O

H O O

O O OH

O HO O

O O

O O

Michael addition

HO

HO

O O

O

O

O

95a

91 (beetleane A)

91

HO O

O

HO

OH

OH HO

–H

HO

OO

O

O

O

OH

O

O O O

O 95

=

OH O

–HB

O O HO O

O

O

OH O

OH O

B OH O

O O O O H ent-90 (ent-epicoane A)

OH

90 (epicoane A)

(±)-94

HO

+

O O

O O O

O O

O O

(±)-93

O O O OH

O

OH

OHO

–H

(±)-92

62

OHO

O O

O O

O

61 (epicoccine)

“H2”

O

OH

O

HO

O

O

O O

O O

[O]

HO O

O

O O

62b

62c

O

(±)-96

O 97

O

O [O] 61

O

O O

OH O

HO

O O OHO

OH O O

O 62

(±)-93

Fig. 20 The proposed biosynthesis of epicoane A (90) and beetleane A (91) and an alternate biogenesis of 93

functionalization of oxidized epicoccine 62 [51] with a suitable nucleophile via tautomer 62c, followed by reoxidation or the existence of a suitably functionalized flavipin/epicoccine derivative (Fig. 20). The resulting product 96 could then potentially undergo the desired dimerization to give 93.

22

A. J. E. Novak and D. Trauner

While this might seem like a more plausible proposal, the question of stereocontrol still remains. Such a pathway would give racemic 92 if one assumes the absence of any enzymatic catalysis. Given that beetleane A (91) seems to be derived from ent-epicoane A (ent-90), it seems unlikely that Epicoccum nigrum would maintain two different enzymes to give enantiomers of the same product [5]. The existence of racemic 92 would then also imply the (at least transient) existence of racemic epicoane A. The conversion of epicoane A to beetleane A (91) could occur spontaneously, but given that both natural products were isolated in enantioenriched form it seems plausible to suggest the existence of an entity catalyzing the conversion of ent-epicoane A (ent-90) to beetleane A (91). Otherwise, beetleane A (91) should exist as a racemate when one assumes a non-enzymatic, initial dimerization of hydroxy-o-quinones such as 62. In conclusion, the discovery of epicoane A (90) and beetleane A (91) raises interesting questions with regard to pyrogallol chemistry that warrant further studies. A previously unknown dimerization mode of hydroxy-o-quinones was proposed and the formation of enantiomerically enriched compounds from such a cascade in the same organism is certainly unusual [5]. As such, a biomimetic synthesis of epicoane A and beetleane A might help to answer some of the questions posed by these intriguing natural products.

4 [5+2] Cycloadditions in the Synthesis of the Merocytochalasans The cytochalasans are a large (>450 members) and important family of fungal polyketide natural products [61]. The cytochalasans initially were named for their main biological activity, the targeting of the actin skeleton (“cytos” = cell and “chalasis” = relaxation), but with the continuing discovery of additional members they now are known to exhibit a broad range of biological activities, including immunomodulatory, cytotoxic, and nematicidal activities. Owing to their diverse and challenging structures and their interesting bioactivies, members of the cytochalasan group continue to attract significant interest from the chemical synthesis community. Structurally, the cytochalasans are characterized by a perhydro-isoindolone core fused to a macrocyclic ring [61]. The cytochalasans are then further divided into six different groups, based on the different amino acids that the isoindole core includes: the “cytochalasins” (phenylalanine residue), “pyrichalasins” (tyrosine or related residue), “chaetoglobosins” (tryptophan residue), “aspochalasins” (leucine residue), “alachalasins” (alanine residue), and “trichalasin” (valine residue). While these different groups consist of fascinating structures, the recently discovered subclass of the merocytochalasans is arguably the most complex class of cytochalasans and will be the focus of this section.

Complex Natural Products Derived from Pyrogallols

23

4.1 Overview of and Biosynthesis Hypothesis for the Merocytochalasans Merocytochalasans can be defined as cytochalasans that have been fused with one or more pyrogallols. The vast majority of them are derived from aspochalasins (in the following paragraphs “merocytochalasans” is implied to refer to “meroaspochalasins”) [61]. A structural overview of selected aspochalasins and the merocytochalasans is provided in Fig. 21. The merocytochalasans have been isolated between 2015 and 2019 by Zhu, Zhang, and coworkers from fermentation broths of Aspergillus flavipes (Plate 2) and have been shown to potently inhibit cancer cell growth by activation of caspase-3 and the degradation of poly(ADP-ribose) polymerase [62–66]. These highly complex heterodimeric, trimeric, and tetrameric structures have been suggested to arise from the fusion of aspochalasin B (98) or aspochalasin D (99) with the pyrogallol epicoccine (61) and each other (Fig. 21 also shows the suggested parentage) [53, 54, 59, 61–66]. The merocytochalasans can thus be divided into four types: the first type combines a unit of epicoccine (61) with one unit of aspochalasin B (98) to yield asperchalasines B–H (100–102), 113–116) and spicarin B (103). The second type includes epicochalasines A, B (104, 105), asperflavipine B (106), and aspergilasines A–D (107–109, 134), and was suggested to involve two units of epicoccine (61) and one unit of aspochalasin B or D (98, 99 aB, aD, respectively). The third type, represented by asperchalasine A (110) and amichalasines A–E (not shown in Fig. 21), appears to involve one unit of epicoccine (61) and two units of aspochalasin B or D. Type four, represented by asperflavipine A (111), incorporates two units of epicoccine (61) and two of aspochalasin B (98). The exact manner in which these compounds come together is discussed below [54, 61]. The plausible biosynthesis origin of the heterodimers, asperchalasines B–H (100– 102, 113–116), is shown in Fig. 22 [53, 61]. The pyrogallol epicoccine (61) is proposed to undergo oxidation to the corresponding hydroxy-o-quinone 62, which can tautomerize to 62b and also isobenzofuran 62d. Subsequently, 62d is then proposed to undergo an intermolecular isobenzofuran Diels–Alder reaction with aspochalasin B (98) that could lead to four different products (asperchalasines F–H and 112). The different transition states depicting the different cycloaddition modes (endo/exo and regioisomeric additions) leading to the different products are shown as TS1–4 [53]. It should be noted that endo/exo is here defined in regard to the C-19–21 conjugated system. Asperchalasines F–H (113–115 and 112) could then be selectively methylated giving rise to asperchalasines B–E (100–102, 116). Curiously, 112 has never been isolated from natural sources, even though its existence is implicated by the existence of asperchalasine E (116) and the fact that this compound seems to be an intermediate en route to asperchalasine A (110) and asperflavipine B (106). The failure to isolate 112 from natural sources could point toward a tendency of 112 to undergo rapid oxidation and follow-up reactions. A biosynthesis hypothesis for the heterotrimeric and heterotetrameric congeners is shown in Fig. 23, with aspergilasine D (109), epicochalasines A, B (104, 105),

24

A. J. E. Novak and D. Trauner O HO

HN

HO

OO

O

OO

OH

O

O

O

HN OO

OH

O

O

O HO

O

O

O HN

O HO O O

O

O O O

O OH O

O HN

O

O

O

O O HO

HN

O

O

OH

O

O

O O

O

O

O OH 106 (asperflavipine B) (2e + aB)

O

O

O

HN

OH

O

O HO

O HN O

O

O

O OH OH OH 107 (aspergilasine A) (2e + aD)

OAc

OH O HN

O

OH 105 (epicochalasine B) (2e + aD)

O

O

OAc

O HO OH

O HO OH OH 104 (epicochalasine A) (2e + aD)

O

OAc AcO 103 (spicarin B) (e + aB)

HO O 102 (asperchalasine D) (e + aB)

HO O 101 (asperchalasine C) (e + aB)

O OH 100 (asperchalasine B) (e + aB)

OO

OH

OH

OH

HO

O

OH HO 99 (aspochalasin D) (aD)

HN

HN

HN

OO

OH O 98 (aspochalasin B) (aB)

61 (epicoccine) (e)

O

HN OO

OH

O HO

O O OH

OH 108 (aspergilasine C) (2e + aD)

O O OH

OH 109 (aspergilasine D) (2e + aD)

OH HO O

O

H O O

N

O O O H HO

O N

O

O

O

O OH

O HN

NH O

O O

HO 110 (asperchalasine A ) (e + 2aB)

O O HO

O OH OH O OH

111 (asperflavipine A) (2e + 2aB)

Fig. 21 Select overview of the merocytochalasans and their proposed parentage from epicoccine (61) and aspochalasin B (98) or D (99)

Complex Natural Products Derived from Pyrogallols

OH

O

O

HN

OH OH

O 98 (aspochalasin B)

21 O

8'

20

19 18 8'

O

H

HO

OH O

TS-1

TS-2

TS-3

HN O

O

HO

O

O

OH

[Me]

HN O

O

OH

HO O OH 116 (asperchalasine E)

O

OO

OH

HO OH 114 (asperchalasine F) [Me]

HN

O

O

O

OH

OH

OH HO OH 115 (asperchalasine H)

HN OO

O

HO O OH 100 (asperchalasine B)

O

[Me]

HN OO

C20–C8' & C19–C1' linkage

OH

HO OH 113 (asperchalasine G)

[Me]

OO

O

HO

HO 112

endo-(4+2)

HN OO

OH

H

O

TS-4

C20–C8' & exo-(4+2) C19–C1' linkage

HN OO

OH

OH

18 20 19

HO

C19–C8' & C19–C8' & endo-(4+2) exo-(4+2) C20–C1' linkage C20–C1' linkage

OO

OH Me 1' H 8' O O 21

OH

OH HO

HN

HO

20 19 21 OH 18 O O 1' 8' OH

OH 18 20 19

OH O

1'

O

path d

H O

21

OH

path c

H

OH

O

OH

O 62

OH 62b

path b

HO

1'

OH O

O

O

OH 62d

[O]

61 (epicoccine)

path a

HO H

OH

O

+ O OO

25

O

OH

OH HO O 101 (asperchalasine C)

OO

O

O

OH

OH HO O 102 (asperchalasine D)

Fig. 22 The proposed biosynthetic origin of the asperchalasines

asperchalasine A (110), and asperflavipine A (111) as representative examples of the different types. All of these highly complex natural products are proposed to be formed through hydroxy-o-quinone [5+2] cycloadditions between units of epicoccine (61) and aspochalasin B or D (98, 99) [53, 54, 59, 61–66]. These [5+2] cycloadditions are closely related to that one occurring in the Trauner synthesis of

26

A. J. E. Novak and D. Trauner

O O

O HN

O OH HO 62b O

O

O O O OH HN

[5+2] O

O

O O HO

OHO OH

O OOH

118

OH

119

HN OO

O OH OH

[O]

104 (epicochalasine A)

61

105 (epicochalasine B)

OH HO 117

OH O O O HN

O OH HO 62 O

O [5+2]

109 (aspergilasine D)

O OHO OH 118

OH O 112

O

H

N

O O

[O]

[5+2]

O O N O OH H O O HO 98 (aspochalasin B) 120

98

110 (asperchalasine A)

HO O 113 (asperchalasine G) +

O

O O

NH

[O]

[5+2]

O

O OH

O

114 (asperchalasine F) 121

O

111 (asperflavipine A)

HN

HO O

H2O

O O

O OH 122

Fig. 23 The proposed biosynthetic origin of selected complex merocytochalasans

Complex Natural Products Derived from Pyrogallols

27

Plate 2 Colonies of Aspergillus flavipes

epicolactone (64) and isoepicolactone (82) [52]. As an example, for both the aspergilasines and epicochalasines, a unit of epicoccine (61) could dimerize with a unit of aspochalasin D (99) through an intermolecular isobenzofuran Diels–Alder reaction, as shown before in Fig. 22, yielding 117. The pyrogallol 117 then could be oxidized to the corresponding hydroxy-o-quinone 118. This o-quinone may then undergo two regioisomeric [5+2] cycloadditions with another unit of oxidized epicoccine, the tautomeric hydroxy-o-quinones 62 and 62b. These two [5+2] cycloadditions would furnish both the natural product aspergilasine D (109) and its regioisomer 119. Aspergilasine D (109) and 119 are hypothesized to be precursors of epicochalasines A, B (104, 105), which could arise from 109 and 119 via a carbonyl-ene reaction or a Prins-type reaction [54]. In a similar manner, asperchalasine A (110) is proposed to arise from a [5+2] cycloaddition between the hydroxy-o-quinone 120 derived from 112 and aspochalasin B (98) (Fig. 23) [53, 59]. The tetrameric asperflavipine A (111) and most complex member of the natural product class is proposed to arise from a [5+2] cycloaddition between the hydroxy-o-quinones derived from asperchalasine G (113) and asperchalasine F (114), followed by hydration [54, 64].

4.2 Syntheses of the Asperchalasines Following the isolation of the merocytochalasans, three groups independently reported efforts directed toward their synthesis: The groups of Deng, Tang, and Trauner all reported syntheses of aspochalasin B (98), with both the Deng and Trauner

28

A. J. E. Novak and D. Trauner

group reporting the synthesis of aspochalasin D (99) as well [53, 59, 67]. In addition, the Trauner group reported the synthesis of (+)-aspergillin PZ, a pentacyclic aspochalasan, but has so far not disclosed any studies directed toward dimeric, trimeric, or tetrameric merocytochalasans [67]. The three groups achieved independently the synthesis of aspochalasin B (98), all utilizing a Diels–Alder cycloaddition to forge the isoindole core, with both Deng and Trauner relying on a late-stage Horner–Wadsworth–Emmons (HWE) macrocyclization and Tang utilizing a ring-closing metathesis (RCM) [53, 59, 67]. Here, it should be mentioned that Trost and coworkers had previously reported the synthesis of aspochalasin B (98) and that Thomas and Vedejs had synthesized related congeners and that these reports laid the groundwork for the studies following [68, 69]. Nonetheless, more scalable routes toward aspochalasin B (98) were needed, resulting in the three reported novel syntheses of this compound. Having achieved the synthesis of aspochalasin B (98), the group of Tang moved to explore the proposed intermolecular isobenzofuran Diels–Alder reaction between 62d and aspochalasin B (98). Initially, Tang et al. tried to oxidize epicoccine (61) to isobenzofuran 62d with K3 [Fe(CN)6 ] in the presence of aspochalasin B (98). Ultimately, under those conditions, they were unable to detect any formation of the desired asperchalasines, but instead observed only the formation of dibefurin (56), the C i -symmetric homodimer of 62, for which the synthesis had previously been reported by the Trauner group [51, 53]. As an alternate strategy to generate isobenzofuran 62d, the Tang group attempted to dehydrate 135 under acidic conditions at 80°C in the presence of aspochalasin B (98). Under these conditions, they observed the formation of asperchalasines G (113) and H (115) by crude NMR analysis. The purification of these natural products proved to be challenging, but by direct treatment of the crude reaction mixture with NEt3 and Ac2 O they were able to obtain the natural product spicarin B (103, not shown) [53]. In order to access asperchalasines B–E, the Tang group treated a mono-methylated analog of 135 under the same conditions in the presence of aspochalasin B (98). Compound 23 proved to be a superior reaction partner compared to unprotected 135 and from this reaction mixture they were able to isolate all four of asperchalasines B–E (100–102, 116), in a combined yield of 80% in a 10:2:1:1 ratio (Fig. 24) [53]. Interestingly, when the Tang group employed the fully protected hemiacetal 124 as the isobenzofuran source, only endo-products of the Diels–Alder reaction (125 and 126) were detected after complete deprotection in 80% yield and in a 3:2 ratio (Fig. 24) [53]. These results indicate that the steric factors in the isobenzofuran precursors play a crucial role in controlling the endo/exo selectivity of the intermolecular Diels–Alder reaction. The free or partially protected precursors afforded the exo-products as the major products, while the fully protected precursor afforded the endo-products predominantly [53]. As the envisaged biomimetic synthesis of asperchalasine A (110) required endoDiels–Alder product 112 as the precursor, the Tang group was able to proceed with their synthetic plan. Full debenzylation of 125 with Raney Nickel and H2 afforded pyrogallol 112, which was directly exposed to aspochalasin B (98) in the presence of

Complex Natural Products Derived from Pyrogallols

29

HO OH +

O

HN

O OO

OH

OH

O 98 (aspochalasin B)

123 CSA, PhMe, 60°C (80%, 10:2:1:1)

HN

HN OO

O

OH

O

OO

+

O

O

+

OH

HO

HO O

O

OH

OH

116 (asperchalasine E)

100 (asperchalasine B)

HN

HN OO

O

O

OH

OO

+

O

O

OH

OH

OH

HO O 101 (asperchalasine C)

HO O 102 (asperchalasine D)

HO

CSA, PhMe 60°C

OBn + HN

O

(80%, 3:2)

OBn

OO

O

OBn

OH

124

98 (aspochalasin B)

HN

HN OO

O

O

OH

+

OO

O

O

OBn

BnO BnO 125

OH

OBn

BnO

OBn

126

Fig. 24 Synthesis of asperchalasines B–E by Tang and coworkers

30

A. J. E. Novak and D. Trauner

air (Fig. 25). Oxidation of pyrogallol 112 to the corresponding hydroxy-o-quinone 120 presumably effected the desired [5+2] cycloaddition, which furnished asperchalasine A (110) in 57% yield and established an oxidative [5+2] cycloaddition as the key step in the biosynthesis of asperchalasine A (110) [53]. In contrast to the Tang group, who utilized the benzyl-protected epicoccine derivative 124 as the isobenzofuran precursor, Deng and coworkers utilized the allylprotected 127 as the corresponding precursor. Treatment of 127 with AcOH at 120°C generated isobenzofuran 128, which underwent the desired Diels–Alder cycloaddition with aspochalasin B (98), giving the two endo-products 129 and 130 in 78%

Raney Nickel H2, EtOH

HN OO

O

O

air dioxane

HN OO

OH

BnO

O

O

OH

HO

BnO

OBn

HO

125

OH

112

OH O

O

H N

HN

O O

98

OO

O

O

OH

O O N O H

O O

[5+2] (57%)

O

OH O HO

OH

120

120

98 (aspochalasin B)

OH O

O

H O O

N

O O O H HO

O

N

O

HO 110 (asperchalasine A)

Fig. 25 Synthesis of asperchalasine A (110) by Tang and coworkers via an intermolecular [5+2] cycloaddition

Complex Natural Products Derived from Pyrogallols

31

i-Bu

i-Bu HO OAllyl O

AcOH, 120°C

98

O

OO

O

(78%)

OAllyl

OAllyl

HN

HN

OAllyl

O

OH

+

OO

O

OH

O

OAllyl

OAllyl

127

OAllyl

AllylO

128

129

AllylO

i-Bu

i-Bu HN

(49%)

O

HN OO

OH

O

O

O

OO

OH

O

O

OH HO

OH

O

O

O O O H HO

HO

OH

115 (asperchalasine H)

H O O

N

OH

112

OH

OH

O

HO

O

120

130

OAllyl

i-Bu HN

K3[Fe(CN)6]

OO

AllylO

Pd/C, NH4HCO2 (70%)

Pd/C, NH4HCO2 (75%)

98 NaHCO3

1.2:1

OAllyl

N

O

O

HO

110 (asperchalasine A) HO OAllyl O OMe 1. AcOH, 120°C 2. Pd(PPh3)4, Et3SiH

OAllyl

152

+

HN

HN OO

(73%)

O

O

OH

+

OO

O

OH

HO HN

O OO O

OH

HO

1. PTSA 2. Pd/C, NH4HCO2

OH

+

HO

OH

O

116 (asperchalasine E) 1.2:1 102 (asperchalasine D)

98 (aspochalasin B)

O

OH

O

98

HN

HN OO

O

O

OH

+

OO

O

O

OH

OAllyl OH

OH

HO

131 HO

OH

113 (asperchalasine G) (50%)

HO

OH

114 (asperchalasine F) (6%)

Fig. 26 Synthesis of asperchalasines A (110), D (102), E (116), H (115), F (114), and G (113) by Deng and coworkers

32

A. J. E. Novak and D. Trauner

HO

1. AcOH, 65°C 2. TBAF

OTBS

+

O

HN

OTBS

OO

OTBS

+

OO

133 (4%)

O

OH

OH

OH

HO

132

HN

117 (70%)

HO

99 (aspochalasin D)

HO

OH

O

117

O

O

61 K3[Fe(CN)6] buffer, pH = 8

O O

H

O HO

62

O HN

O

+ O

O

O O OH HO

62b

O HO

O HN (51%) O

O

O HO OH

OH

118

118

O

O OH

O

109 (aspergilasine D) + O 2.6:1

HN O

O HO

O

O

O Et3N, air

O

OH

HO

O

(47% from 109)

O O HO

O OH OH

119

HN

O

O OH OH

134 (aspergilasine B)

HO OH O OH 135

117

OH O2, buffer, pH = 8 (42%)

TPAP, NMO

134 (aspergilasine B)

(53%)

108 (aspergilasine C)

Fig. 27 Synthesis of aspergilasines B–D through an intermolecular [5+2] cycloaddition

yield and in a 1.2:1 ratio (Fig. 26) [59]. In agreement with the findings by Tang et al., only the endo-cycloaddition products were observed when the fully protected isobenzofuran precursor 127 was employed. Both cycloaddition products could be deallylated successfully, giving asperchalasine H (115) and 112. The treatment of 112 with K3 [Fe(CN)6 ] resulted in the proposed oxidation of pyrogallol 112 to the corresponding hydroxy-o-quinone 120, which underwent the desired [5+2] cycloaddition with aspochalasin B (98) in the presence of NaHCO3 , thereby yielding asperchalasine A (110) in 49% yield (Fig. 26) [59]. As part of their studies, Deng and coworkers also achieved the synthesis of asperchalasines E and D (116, 102) when 152 was employed as the epicoccine analog. The use of 152 instead of 127 was essential as this research group found that chemical methylation of asperchalasine H (115) and 112 was unselective. In an additional investigation, Deng and coworkers also achieved the synthesis of the exo-Diels– Alder products asperchalasines G and F (113, 114) using monoprotected 131 as the

Complex Natural Products Derived from Pyrogallols

33

isobenzofuran precursor (Fig. 26) [54]. Interestingly, when aspochalasin D (99) was used as the dienophile together with fully TBS-protected 132, only the exo-adducts 117 and 133 where obtained (Fig. 27). Taken together with similar findings by Tang and coworkers, these results indicate that both the steric effects of the diene and electronic effects of the dienophile play crucial roles in controlling the endo/exo selectivity of the isobenzofuran Diels–Alder reaction. In summary, the groups of Tang and Deng achieved (when taken together) the synthesis of the whole class of the asperchalasines and were able to establish isobenzofuran Diels–Alder reactions and [5+2] cycloadditions as the biosynthetic key steps to these fascinating natural products [53, 54, 59].

4.3 Synthesis of the Aspergilasines, Amichalasines, Asperflavipines, and Epicochalasines In a landmark achievement, Deng and coworkers set out to further study the biosynthetic connections between the aspergilasines, amichalasines, asperflavipines, and epicochalasines through biomimetic syntheses [54]. At first, they explored the formation of the aspergilasines via a [5+2] cycloaddition as they reasoned that the epicochalasines could then be derived from the aspergilasines (Fig. 23) [54]. When Deng and coworkers subjected exo-Diels–Alder product 117 and epicoccine (61) to a slightly alkaline buffer and slowly added K3 [Fe(CN)6 ], they observed the formation of aspergilasine D (109) and its regioisomer 119 in a 37% and 14% yield, respectively. They noted that the exact reaction conditions employed were crucial for achieving this regioisomeric outcome, since the use of molecular oxygen as the oxidant afforded only 109 as the sole product in a 61% yield [54]. They also found that aspergilasine D (109) underwent an allylic oxidation when exposed to Et3 N and O2 , yielding aspergilasine B (134) in a 47% yield. Alternatively, aspergilasine B (134) was also obtained from 117 and 135, which already contained the desired hydroxy group. Using a Ley oxidation, aspergilasine B (134) was converted to aspergilasine C (108) in 53% yield. Amichalasines A (138) and B (141) were thought to arise from the union of two units of aspochalasins and one unit of epicoccine, similar to asperchalasine A (110) [54, 66]. Oxidation of 117 under an oxygen atmosphere in the presence of aspochalasin B (98) resulted in a successful intermolecular [5+2] cycloaddition, followed by spontaneous hemiacetal formation, giving 137 in 52% yield (Fig. 28). Amichalasine A (138) was then prepared from 137 by selective oxidation of the keto-hydroxy group to the corresponding diketone using Cu(OAc)2 . In a similar manner, amichalasine B (141) was generated by an intermolecular [5+2] cycloaddition between 139 (prepared by deoxygenation of aspochalasin B (98)) and the hydroxy-o-quinone derived from 117, followed by hemiacetal formation (Fig. 28) [54].

34

A. J. E. Novak and D. Trauner

HN OO

O OH OH +

O2, pH = 8 buffer

HN OO

OH

117

HO

NH O O

O

NH O O

O HN

O HO O

HO O

NH O O

O

OH

OH

Cu(OAc)2 O

(61%)

OH

O HN O OH

138 (amichalasines A)

(45%)

O

O

O O OH HO

O

1. Tf2O pyridine 2. NaI, Δ

HN

OH

O

137

OO

HO O O HO

NH O O

O

O O OH OH HO

(52%)

136

HN

O

O HN O

118

98

O

O

O

OH

OH

O

98 (aspochalasin B)

OH

HN OO

OH

NH O O

118, O2 pH = 8 buffer

O

O

O

O HO O

98 (aspochalasin B)

139

NH O O

O

O

NH O O (54%)

O HO O O HO

140

139

O HN

OH

O HN

O

HO O

OH 118

O HN

O

O O O OH HO

OH 141 (amichalasine B)

Fig. 28 Synthesis of amichalasines A (138) and B (141) through an intermolecular [5+2] cycloaddition

With these results in hand, the Deng group turned their attention to the synthesis of the most complex member of the merocytochalasan family, asperflavipine A (111), a tetramer that is expected to arise from the union of two molecules of aspochalasin B (98) and two molecules of epicoccine (61) (Fig. 23). This constituted a major step regarding the complexity of the desired intermolecular [5+2] cycloaddition.

Complex Natural Products Derived from Pyrogallols

35

While the Deng group had already achieved the heterodimerization of 117 with either aspochalasin B [59] or an epicoccine motif [54], the homodimerization of 117 or the heterodimerization of asperchalasines G (113) and F (114) was without any precedent and several challenges had to be considered. Thus, it was unclear whether the homodimerization of either asperchalasine G (113) or asperchalasine F (114) with themselves would outcompete the desired heterodimerization to asperflavipine A (111). Additionally, either part of asperchalasines G (113) and F (114) could serve as the “5” or “2” part of the [5+2] cycloaddition, with only one possibility leading to asperflavipine A (111). Model studies utilizing asperchalasine G (113) and 117 revealed that the desired heterodimerization product could indeed be formed in synthetically useful yields (not shown), but the resulting product could not be advanced to asperflavipine A (111). Treatment of asperchalasine G (113) in the presence of asperchalasine F (114) with a buffered solution of K3 [Fe(CN)6 ] followed by acidification to facilitate hemiacetal formation, gave asperflavipine A (111) (Fig. 29) [54]. Asperflavipine B (106), in turn, was accessed using pyrogallol 112 and 135 as the [5+2] cycloaddition partners (Fig. 29). Oxidation of 112 in the presence of 135 gave the two regioisomeric products 143 and 144, which were treated under acidic conditions to effect dehydration, giving asperflavipine B (106) in 12% yield. The low yield was attributed to the fact that the major [5+2] cycloaddition product 144 could not undergo dehydration to asperflavipine B (106) [54]. It had been proposed that aspergilasine A (107) might be derived from 119, the regioisomer of aspergilasine D (109) by a formal [3+2] cycloaddition that could be seen also as an aldol–Michael addition cascade [54, 65]. The desired formal [3+2] cycloaddition of 119 to give aspergilasine A (107) could be effected in protic solvent, but the conversion proved to be unsatisfactory. Additionally, the regioisomer aspergilasine D (109) could not be converted to 147 under similar conditions. These findings prompted Deng and coworkers to explore alternative ways to achieve the formal [3+2] cycloaddition of both aspergilasine D (109) and 119. Overall, they envisaged that an intramolecular [2+2] photocycloaddition followed by an acyloin rearrangement of the resultant α-hydroxy ketone (145 or 146) could achieve the desired transformation (Fig. 30) [54]. To this end, neat aspergilasine D (109) and its regioisomer 119 were exposed to sunlight, which gave the desired formal [3+2] cycloaddition products aspergilasine A (107) and 147 in excellent yields (Fig. 30) [54]. The facility of the [2+2] cycloaddition–acyloin rearrangement cascade raises the question of whether this step might be biosynthetically relevant. Additionally, acyloin rearrangements of α-hydroxy ketones are used normally to convert [4.6]bicyclic compounds to [5.5]bicyclic compounds and not to a [3.2.1]bridged bicycle as in this case [54]. The effected cascade could be the first conversion of a [4.6]bicyclic ring system to a [3.2.1]bridged bicyclic system and therefore may find further application in natural product synthesis. The epicochalasines A (104) and B (105) were hypothesized to arise from aspergilasine D (109) and its regioisomer 119 by a carbonyl-ene or Prins-type reaction [54, 63]. However, the transformation of either aspergilasine D (109) or 119 to the epicochalasines proved more challenging than Deng and coworkers had anticipated. In

36

A. J. E. Novak and D. Trauner HO O

O

HO

O

113 (asperchalasine)

O

O

+

NH

K3[Fe(CN)6] buffer, pH = 8

O

1N HCl

O OH

O

O

(43%)

NH

HN

HO

O

O

O

O

O HN O

O HO

O O

O OH

121

114 (asperchalasine)

O

O

O OH OH O OH

111 (asperflavipine A)

122

OH

135 K3[Fe(CN)6] buffer, pH = 8

HN OO

O

OH

O

HO HO

OH

112

O O

O

OH O HN

OH

O

+

O

O

O

O

HO

O O

OH

O

HO

120

O

OH

O

120

O

OH O HN

O O HO

+

OH O HN O

O O HO O

O

O

142a

142

O O HO

O

O

HO

OH O HN

O

HO

O OH

O O

O

O OH

144

143 1N HCl (12%)

O O O HO

OH O HN O

O

O O

O OH

106 (asperflavipine B)

Fig. 29 Synthesis of asperflavipines A (111) and B (106) via intermolecular [5+2] cycloadditions

Complex Natural Products Derived from Pyrogallols

O OH

O

O

O HN

O

neat, sunlight [2+2]

O O HO

O

O

O

O OH

O

HN

neat, sunlight [2+2]

O O HO

O

O

O

TFE

O OH

Me3P

O H Me3P

O

O OH

OH

OH

148

109

TFE Me3P

O OH

O

O

O OH

150

OH

H O Me3P O O

O OH OH

O

O H

HN O

O O O HO

O OH

OH

104 (epicochalasine A) (12%) +107 (aspergilasine A) (73%)

O

O

O

O OH

151

O

O HO

HN O

OH

O

O

HO

OH

OH

HN

O

O HN

O OH

O

147

O

O HN O

O HO

O

OH

149

O O

(81%)

O

O

O O O

OH

OH

OH

OH

O

O OH

O O

HN

O

O HN O

O O HO

PMe3

OH

146

O

HN

O

O OH

O

OH

O

O

107 (aspergilasine A)

H

109 (aspergilasine D)

119

O

O O

OH

O

O OH

PMe3

O

OH

O

O O

(96%)

O

145

O OH

OH

O OH

O

OH

O

O HN

H

119

O

O

OH

37

OH

O HN O

O O HO

O OH

OH

105 (epicochalasine B) (21%) +117 (36%) +109 (24%)

Fig. 30 Synthesis of aspergilasine A (107) via a [2+2] cycloaddition/acyloin ring expansion cascade and epicochalasines A (104) and B (105) via a MBH-type reaction

the case of aspergilasine D (109), thermal conditions (DMSO, 160°C) only yielded the [3+2] cycloaddition product 147, while alkaline conditions in order to effect an aldol-type reaction led to decomposition through a retro-[5+2] cycloaddition to 117 [54]. Additionally, as had been shown before, aspergilasine D (109) converted quite readily to aspergilasine B (134) in the presence of base and air and even thorough removal of oxygen from the reaction mixture did not result in the formation of any carbonyl-ene product.

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Following these setbacks, Deng et al. hypothesized that they might be able to convert aspergilasine D (109) and 119 to the epicochalasines by a Morita–Baylis– Hillman (MBH)-type reaction [54]. A nucleophile, such as a phosphine, could attack 109 or 119 in a Michael addition and generate the corresponding enolate (148 or 150) (Fig. 30). This enolate might then attack the desired carbonyl group in an aldol addition yielding 149 and 151, respectively. Elimination of the nucleophile could then regenerate the olefin, giving 104 and 105. The caged, β,β-disubstituted Michael system in 109 and 119 proved to be quite sensitive to steric factors, as treatment with PPh3 , PCy3 , or PBu3 failed to promote the desired MBH reaction [54]. Ultimately, treatment of 119 with the comparably small nucleophile PMe3 in 2,2,2-trifluoroethanol (TFE) afforded the desired formal carbonyl-ene product, epicochalasine A (104) in a 12% yield together with the [3+2] cycloaddition product aspergilasine A (107) in a 73% yield (Fig. 30) [54]. Under the same reaction conditions epicochalasine B (105) could also be obtained from aspergilasine D (109) in a 21% yield, with the retro-[5+2] cycloaddition product 117 also being isolated in a 36% yield. In conclusion, the work by Tang and Deng shed light on the biosynthesis and the biosynthetic relationships of the merocytochalasans, and has established [5+2] cycloadditions of hydroxy-o-quinones as the elementary processes that generate their complex structures [53, 54, 59, 61]. Through their use of biomimetic [5+2] cycloadditions and Diels–Alder reactions, the Deng group ultimately was able to synthesize 11 different complex merocytochalasans and several such compounds isolated from their studies may also prove to be as of yet undiscovered natural products [54, 59]. These examples present the synthetically most complex cases of [5+2] cycloadditions to date, but also highlight the high selectivity, the mild reaction conditions employed, and the synthetic utility of such cycloadditions in an extremely complex environment.

5 Preuisolactone A—Another Racemic Fungal Natural Product Preuisolactone A (153) is a complex metabolite that has been isolated recently by Yang and Abe from the endophytic fungus Preussia isomera XL-1326, obtained from the stems of Panax notoginseng, (Chinese ginseng) [70] (Plate 3). The structure of preuisolactone A (153) was elucidated by a combination of spectroscopic methods as well as via X-ray crystallography. The complex structure of preuisolactone A (153) consists of an unprecedented, tricyclo[4.4.01,6 .02,8 ]decane skeleton, which features seven stereogenic centers and five rings that are fused to each other (Fig. 31). Other structural features include two butyrolactones, a vinylogous methyl ester, and a tertiary alcohol. In addition to its challenging structure, preuisolactone A (153) possesses antibacterial activity against Micrococcus luteus (MIC = 10.2 μM) and moderate antifungal

Complex Natural Products Derived from Pyrogallols

39

Plate 3 Colony of Preussia isomera

Fig. 31 The structure and crystal structure of preuisolactone A (153) and a ginseng plant (Panax ginseng)

40

A. J. E. Novak and D. Trauner

Fig. 32 The bridgehead ring systems in epicolactone (64) and preuisolactone A (153)

O HO O HO

O

OH

O O OH

O O

O

O

O 64 ((±)-epicolactone)

O 153 ((±)-preuisolactone A)

activity [70]. Preuisolactone A (153) was isolated as a racemate (reported as a 51:49% scalemic mixture) and, unlike the majority of natural products, as an enantiomerically pure compound [5]. The isolation team proposed a terpenoid origin for 153, starting from farnesyl pyrophosphate, which was proposed to undergo an initial cyclization, followed by a complex pathway of rearrangements, further cyclizations, and several oxidative tailoring steps [70]. To account for the racemic nature of 153, the authors proposed that two different terpene cyclases might catalyze the initial cyclization of farnesyl pyrophosphate, thereby affording both enantiomers. This proposal seemed implausible to Trauner and coworkers and they proposed an alternative biosynthesis hypothesis, which was subsequently supported by a biomimetic synthesis of preuisolactone A. The bridged carbon skeleton in preuisolacone A (153) bears a resemblance to epicolactone (64), with an apparent ring contraction having occurred (Fig. 32). Therefore, they reasoned that 153 might be the product of a [5+2] cycloaddition, retro-Dieckmann, vinylogous aldol cascade as occurs in the biomimetic synthesis of epicolactone (64) (see Sect. 3.2). Their biosynthesis hypothesis thus included that preuisolactone A (153) is not a terpenoid but a polyketide [2]. They proposed that 153 is formed through an initial oxidative dimerization of catechol 154 with pyrogallol 155 (Fig. 33). In fact, pyrogallol 155 is a known fungal metabolite that is derived from orsellinic acid [71]. The oxidation of 154 and 155 would yield o-quinone 156 and hydroxy-o-quinone 157, respectively, which should undergo a [5+2] cycloaddition giving the tricyclic intermediate 158. As shown in Sect. 2, such reactions are well-precedented in the synthesis of benzotropolones like purpurogallin (6) [28, 29]. Additionally, these types of cycloadditions also occurred in the synthesis of epicolactone (64) and the syntheses of the merocytochalasans [52–54, 59]. Trauner’s group proposed that water could attack the carbonyl bridge in 158, which should result in a retro-Dieckmann-type fragmentation, yielding hydroxy dienol 159 [2, 5, 6]. In the purpurogallin cascade similar intermediates are prone to further oxidation and tautomerization, but the quaternary carbon atom in 159 prevents aromatization. Instead, 159 should undergo a vinylogous aldol addition furnishing diosphenol 160. They proposed that 160 could then undergo a formal oxidative lactonization, presumably through an oxa-Michael or Prins-type reaction. The resulting ene-diol then may be oxidized to the corresponding diketo lactone 163, which cannot enolize again due to ring strain. For the last steps in the proposed cascade toward preuisolactone A (153), they proposed a ring contraction by a benzilic acid rearrangement from

Complex Natural Products Derived from Pyrogallols OH

41

HO

OH

OH

+

OH

O 154

155

K3[Fe(CN)6] NaHCO3

MeCN/H2O

O O

O

O O

156 O

[5+2] O

OH O

O

O

OH O

H2O O

O

O

OH

O O 161

O 160

O

O

O 162

O O

O

O O

OH

OH

OH O

O

O

O

OH HO

O O

O

O

I OH Ph

1) NaOH then HCl OH 2) I Ph OTs DMF, then phosphate buffer pH = 8

O OH O

(73%)

159

158

157

O

OH

HOOC

OH

O O 163

O O 164

O O

H O (57%)

O

O OH

O O 153 ((±)-preuisolactone A)

Fig. 33 Biomimetic synthesis of preuisolactone A (153)

163 via oxetane 164, which should simultaneously form one of the two butyrolactones and the tertiary alcohol in preuisolactone A (153) [2, 5, 6]. Employing conditions similar to those used in their epicolactone synthesis, Trauner and Novak obtained successfully a mixture of products corresponding to the [5+2] cycloaddition/retro-Dieckmann/vinylogous aldol cascade [2, 5, 6]. The mixture consisted of diosphenol 160, presumably in its keto form, and acetal 161. The ratio of the isomers 160 and 161 could be altered by utilizing an acid–base extraction. Thus, first treating the mixture of isomers with base, followed by acidification, enriched the diosphenol 160. Subsequent treatment with Koser’s reagent effected an oxidative lactonization, presumably via the intermediate iodine(III) species 162, which then underwent the proposed benzilic acid rearrangement upon workup with phosphate buffer (pH = 8) under slightly basic conditions to give (±)preuisolactone A (151) in good yield [2, 5, 6]. Notably, phosphate buffer proved to be essential as a quenching agent, whereas saturated aqueous NaHCO3 led only to decomposition.

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Acknowledgements Alexander J.E. Novak thanks New York University for a MacCracken and a Ted Keusseff fellowship. Dirk Trauner would like to thank the U.S. National Institutes of Health for financial support (grant R01GM126228).

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49. Tkachev VV, Aldoshin SM, Shilov GV, Komissarov VN, Sayapin YA, Korobov MS, Borodkin GS, Minkin VI (2007) Structure of the oxidative dimerization product of 4,6-di(tertbutyl)pyrogallol. Russ Chem Bull 56:276 50. Brill GM, Premachandran U, Karwowski JP, Henry R, Cwik DK, Traphagen LM, Humphrey PE, Jackson M, Clement JJ, Burres NS, Kadam S, Chen RH, McAlpine JB (1996) Dibefurin, a novel fungal metabolite inhibiting calcineurin phosphatase activity. J Antibiot 49:124 51. Ellerbrock P, Armanino N, Trauner D (2014) Biomimetic synthesis of the calcineurin phosphatase inhibitor dibefurin. Angew Chem Int Ed 53:13414 52. Ellerbrock P, Armanino N, Ilg MK, Webster R, Trauner D (2015) An eight-step synthesis of epicolactone reveals its biosynthetic origin. Nat Chem 7:879 53. Bao R, Tian C, Zhang H, Wang Z, Dong Z, Li Y, Gao M, Zhang H, Liu G, Tang Y (2018) Total syntheses of asperchalasines A–E. Angew Chem Int Ed 57:14216 54. Long X, Wu H, Ding Y, Qu C, Deng J (2021) Biosynthetically inspired divergent syntheses of merocytochalasans. Chem 7:212 55. da Silva Araújo FD, de Lima Fávaro LC, Araújo WL, de Oliveira FL, Aparicio R, Marsaioli AJ (2012) Epicolactone—natural product isolated from the sugarcane endophytic fungus Epicoccum nigrum. Eur J Org Chem 2012:5225 56. Talontsi FM, Dittrich B, Schüffler A, Sun H, Laatsch H (2013) Epicoccolides: antimicrobial and antifungal polyketides from an endophytic fungus Epicoccum sp. associated with Theobroma cacao. Eur J Org Chem 2013:3174 57. Yan Z, Li J, Ye G, Chen T, Li M, Liang Y, Long Y (2020) Fused multicyclic polyketides with a two-spiro-carbon skeleton from mangrove-derived endophytic fungus Epicoccum nigrum SCNU-F0002. RSC Adv 10:28560 58. Yan Z, Huang C, Guo H, Zheng S, He J, Lin J, Long Y (2020) Isobenzofuranone monomer and dimer derivatives from the mangrove endophytic fungus Epicoccum nigrum SCNU-F0002 possess α-glucosidase inhibitory and antioxidant activity. Bioorg Chem 94:103407 59. Long X, Ding Y, Deng J (2018) Total synthesis of asperchalasines A, D, E, and H. Angew Chem Int Ed 57:14221 60. Cao P-R, Zheng Y-L, Zhao Y-Q, Wang X-B, Zhang H, Zhang M-H, Yang T, Gu Y-C, Yang M-H, Kong L-Y (2021) Beetleane A and epicoane A: two carbon skeletons produced by Epicoccum nigrum. Org Lett 23:3274 61. Zhu H, Chen C, Tong Q, Zhou Y, Ye Y, Gu L, Zhang Y (2021) Progress in the chemistry of cytochalasans. Prog Chem Org Nat Prod 114:1 62. Zhu H, Chen C, Xue Y, Tong Q, Li X-N, Chen X, Wang J, Yao G, Luo Z, Zhang Y (2015) Asperchalasine A, a cytochalasan dimer with an unprecedented decacyclic ring system, from Aspergillus flavipes. Angew Chem Int Ed 54:13374 63. Zhu H, Chen C, Tong Q, Li X-N, Yang J, Xue Y, Luo Z, Wang J, Yao G, Zhang Y (2016) Epicochalasines A and B: two bioactive merocytochalasans bearing caged epicoccine dimer units from Aspergillus flavipes. Angew Chem Int Ed 55:3486 64. Zhu H, Chen C, Tong Q, Yang J, Wei G, Xue Y, Wang J, Luo Z, Zhang Y (2017) Asperflavipine A: a cytochalasan heterotetramer uniquely defined by a highly complex tetradecacyclic ring system from Aspergillus flavipes QCS12. Angew Chem Int Ed 56:5242 65. Wei G, Chen C, Tong Q, Huang J, Wang W, Wu Z, Yang J, Liu J, Xue Y, Luo Z, Wang J, Zhu H, Zhang Y (2017) Aspergilasines A–D: four merocytochalasans with new carbon skeletons from Aspergillus flavipes QCS12. Org Lett 19:4399 66. Wu Z, Tong Q, Zhang X, Zhou P, Dai C, Wang J, Chen C, Zhu H, Zhang Y (2019) Amichalasines A-C: three cytochalasan heterotrimers from Aspergillus micronesiensis PG-1. Org Lett 21:1026 67. Reyes JR, Winter N, Spessert L, Trauner D (2018) Biomimetic synthesis of (+)-aspergillin PZ. Angew Chem Int Ed 57:15587 68. Trost BM, Ohmori M, Boyd SA, Okawara H, Brickner SJ (1989) Palladium-catalyzed synthesis of macrocycles. A total synthesis of (–)-aspochalasin B. J Am Chem Soc 111:8281 69. Vedejs E, Rodgers JD, Wittenberger SJ (1988) A sulfur-mediated total synthesis of zygosporin E. J Am Chem Soc 110:4822

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A. J. E. Novak and D. Trauner Alexander J. E. Novak received his B.S. and M.S. degrees from Ludwig Maximilian University of Munich in Germany. During his Master’s program, he worked with Prof. Phil S. Baran at The Scripps Research Institute on the development and application of radical reactions in total synthesis. He obtained his Ph.D. degree from New York University in 2021, where he studied the biomimetic synthesis of complex natural products in the laboratory of Prof. Dirk Trauner. In October 2021, he joined Bayer Pharmaceuticals in Wuppertal, Germany, as a head of a laboratory in process research and development.

Dirk Trauner was born and raised in Linz, Austria, studied biology and chemistry at the University of Vienna, and received his Master’s degree in chemistry from the Free University, Berlin. He then pursued a Ph.D. degree in chemistry under the direction of Prof. Johann Mulzer, with whom he moved to the University of Frankfurt and then back to Vienna. Subsequently, he became a postdoctoral fellow with Prof. Samuel J. Danishefsky at the Memorial Sloan-Kettering Cancer Center. After two years in New York City, Dr. Trauner joined the Department of Chemistry at the University of California, Berkeley, where he rose through the ranks to become an Associate Professor of chemistry and a member of the Lawrence Berkeley National Laboratory. In the summer of 2008, he moved to the University of Munich, where he served as a Professor of Chemical Biology and Chemical Genetics for nine years. Subsequently, he returned to the U.S. to become the Janice Cutler Chair of Chemistry at New York University. In the Spring of 2022, he will move to the University of Pennsylvania as a Penn Integrates Knowledge Professor in chemistry and medicine.

The Chemistry of Agarwood Odorants Nicolas Baldovini

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Agarwood Generation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Botanical Aspects of Agarwood-Producing Species . . . . . . . . . . . . . . . . . . . . . . . 2.2 Natural and Artificial Agarwood Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Agarwood Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Agarwood Volatiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Non-volatile Constituents of Agarwood . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Agarwood Odorants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Naturally Occurring Agarwood Odorants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Thermal Generation of Volatiles from Agarwood . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

47 52 52 56 57 58 74 77 79 86 89 91

1 Introduction The name “agarwood” is a general term that refers to the infected heartwood of various trees of the Thymelaeaceae family, belonging mostly to the genera Aquilaria and Gyrinops. These trees are endemic to the tropical forests of Southeast Asia, and their wood is generally soft, light, pale yellow or white, and of little value when the trees are healthy. However, their natural or artificial infection by fungi and microorganisms leads to the pathological secretion of a dark oleoresin in the stem tissues. The resulting resinous dense wood material constitutes agarwood (Plates 1 and 2). In some literature, the term “agarwood” (sometimes misspelled as “agalwood”) is used to designate the oleoresin embedded inside the stem, but to be in line with the most N. Baldovini (B) Institut de Chimie de Nice, Université Côte d’Azur, Parc Valrose, 06108 Nice, France e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2022 A. D. Kinghorn, H. Falk, S. Gibbons, Y. Asakawa, J.-K. Liu, V. M. Dirsch (eds.), Progress in the Chemistry of Organic Natural Products 118, https://doi.org/10.1007/978-3-030-92030-2_2

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N. Baldovini

Plate 1 Agarwood samples, with their alleged geographical origin and botanical species

Plate 2 Highly infected agarwood samples (sinking grades). Major volatile constituents: A epi-γeudesmol, jinkoh-eremol, kusunol, and oxo-agarospirol. B jinkohol

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common definition, in the present chapter, “agarwood” represents the crude material obtained by a simple mechanical carving of the wood collected from infected trees. In many Asian languages, the etymology of agarwood vernacular names illustrates a striking property of this material: the word agarwood itself derives from the Sanskrit name (ag¯aru or aguru) meaning “non-floating wood” [1] and the same sense appears ` hu,o,ng), and Japanese (沈香 in Chinese (沉香 “C’hen siang”), Vietnamese (trâm “Jinkô”), with all terms translated as “sinking incense wood”. Indeed, the density of the wood significantly increases as a result of its infection, and highly infected resinrich samples can even sink in water (Plate 2). In fact, this simple test is still used as a quick way to control agarwood quality, and the “sinking grade” samples are generally considered the most valuable. Nowadays, agarwood is by far the most expensive wood in the world. In today’s market, high-quality samples (sinking grade) reach astonishingly high prices, attaining up to US$150,000 per kilogram [2], and the value of the most beautiful pieces of the prestigious “kynam” type can even be estimated to one million $US per kilogram. The commercial value of agarwood can be explained by its precious status in Asian traditional medicine systems (especially Chinese and Ayurvedic) and by its fragrant properties of high esteem in the Middle East [3] and in northern East Asia (Japan, Korea, China, and Taiwan) [4–6]. Indeed, agarwood is considered the most precious perfumery material in many oriental countries, and it was described as such as early as 1400 B.C.E. in Sanskrit texts [1]. It also occupies a central role in the kôdô (香道), literally, “the way of the incense”, the traditional Japanese art related to the ritual of burning incense (Plate 3) [7]. In the Middle East (Saudi Arabia, UAE, Kuwait, and Qatar), agarwood is named “oud” (meaning simply “wood” in Arabic) and is either burned for religious and scenting purposes or used in the form of its essential oil for the preparation of perfumes (Plate 4). In contrast, the introduction of this material in Western perfumery came rather recently, at first with “M7 oud absolute” (Yves Saint Laurent, 2002), by A. Morillas and J. Cavallier [8]. The launch of this atypical masculine woody perfume was the starting point of a distinct trend for oud in the Western world, as exemplified by many other perfumery creations like “Oud Royal” by E. Boulanger

Plate 3 A The traditional Japanese ritual of incense burning (kôdô). A, B to generate the fragrance of agarwood, a small piece of the wood is heated (ca. 200°C) on a mica plate placed on top of a cone of ashes covering burning charcoal. C A kôdô practitioner is enjoying agarwood odor (“listening to the incense”, according to the Japanese term), by bringing the sample close to her nose. Photograph courtesy of Yamadamatsu Corp., Kyoto, Japan

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Plate 4 Commercial samples of agarwood essential oils. Samples on the left were purchased in Dubai (UAE) in their original Middle Eastern style flask

(Armani privé, 2010), “Oud for love” (B. Duchaufour, The Different Company, 2012), “The Night” (D. Ropion, Frédéric Malle, 2014), “Oud Palao” (F. Pellegrin, Diptyque, 2015), “Oud Satin Mood” (F. Kurkdjian, Maison Francis Kurkdjian, 2017), and “Leather Oud” (F. Demachy, Christian Dior, 2018), to name but a few products [9, 10]. Today, the oud fashion is still very prevalent in fine perfumery, even if behind this trend, the actual use of large amounts of authentic agarwood essential oil in fine perfumes is probably anecdotal due to the extremely high price of this ingredient. Indeed, agarwood essential oil is recognized unanimously as the most prestigious and expensive natural raw material used in modern perfumery [11, 12]. The high prices of agarwood are also a consequence of its scarcity, as the overexploitation of agarwood-producing species has led to a severe depletion of the natural resources in most source countries. Therefore, many of these species have been included in the IUCN (International Union for Conservation of Nature) Red List of Threatened Species and are now protected under CITES regulations (Convention on the International Trade in Endangered Species of Wild Fauna and Flora). To fulfill the demand, the cultivation of agarwood-producing species and their artificial inoculation has been developed since the late 1990s, and a large part of agarwood essential oil on the current market is now produced from cultivated agarwood. However, the complex mechanisms by which agarwood resin is produced and the identity of its most important odorants are still debated. Moreover, the demand for agarwood is still higher than its production level, which was estimated in 2008 to embrace only 40% of the demand [13]. However, a number of adulteration issues complicate the access to authentic samples and impede rigorous phytochemical investigations on this material. This is due to the fact that access to samples of wild agarwood of certified geographical and botanical origin is very difficult: the rarity of the resource and the extremely high income received by successful agarwood hunters result in strong tensions in the trade chain. For safety reasons, the exact location of infected trees is generally kept confidential by local harvesters [14]. This situation was already common in the nineteenth century [15], which illustrates the length of time this material has been highly valued. The substantial rarefaction of authentic wild agarwood

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during the last few decades has further increased even more the pressure on collectors, who would have to leave their village for weeks after a successful agarwood harvesting, to protect themselves from racketeering [16]. The generation, the pharmacological properties, and the chemical composition of agarwood and all have been investigated extensively, and several reviews have been published on these various topics, especially on the phytochemical aspects [2, 17– 24]. It appears that the composition of agarwood is extremely variable and complex and is dominated by two main groups of secondary metabolites: the sesquiterpenoids and the chromones. To date (July 2021), 203 sesquiterpenoids and 266 chromones have been described as agarwood constituents, a significant increase since the 2012 review of Chen et al. [18], who reported 69 and 53 representatives of these two groups, respectively. Indeed, analytical chemistry on this material has enriched considerably the knowledge of its composition, particularly as a result of the contribution of Chinese research groups who have investigated the phytochemistry of agarwood (especially from Aquilaria sinensis) over the last five years. These advances are also reflected in the pace of publication of the reviews on agarwood composition, as 10 out of the 13 reviews cited above were published after 2016. Nevertheless, a critical evaluation of the literature is crucial whenever one wants to present a reliable overview of phytochemical knowledge on such a chemically complex and difficult to authenticate natural material. Among the high-quality reviews published recently on the composition of agarwood, the reader is invited to refer to the very recent publication of Li et al. [17], for an exhaustive overview of its constituents and their biosynthesis. In view of the importance of agarwood in systems of Asian traditional medicine, the pharmacological properties of its bioactive components are obviously a relevant aspect, which was also covered by several authors [17, 22, 25–27] and hence will not be surveyed in the present contribution. In addition, other interesting topics were also reviewed recently, such as the methods for the artificial production of agarwood [28, 29] and historical summaries on the use and trade of this material, both worldwide [1] and specifically in Japan [30]. The present contribution aims at bringing a new perspective on the chemistry of this fascinating natural material, by focusing specifically on the agarwood components of relevance for its olfactory properties. Indeed, in spite of the major use of agarwood as a fragrance material, this issue is rarely addressed in research articles and reviews. The masterful 2011 review of R. Näf [2] was an exception, which emphasized the utilization of agarwood in perfumery, but, given the advances made since this publication appeared, it is considered timely to provide an updated treatment of the chemistry of agarwood odorants. Therefore, the present chapter gives an overview of the literature on the odorant constituents of agarwood, either genuinely present in the material or generated during heating. More than 300 references (articles, reviews, patents, congress proceedings, Ph.D. theses, website pages, and reports, including sources available only in the Japanese language) from 1935 until the present have been evaluated critically, to build an updated and comprehensive perspective of the current knowledge on the molecular identity of the odor of the “wood of the gods”, to use one of the names given to this fascinating material. As mentioned prudently by Li et al. [17], in view of

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the tremendous increase in knowledge of the science of agarwood, the dawn has been reached of a golden age of the research on agarwood properties and its exploitation. The same is certainly true for the quest for its key odorants since the complexity and the variability of the agarwood fragrance will still require many further investigations to identify the specific nature of all of its mysterious olfactory aspects, and maybe to eventually help perfumers to use agarwood at an affordable price in their product compositions.

2 Agarwood Generation In contrast to most fragrant materials derived from aromatic plants, agarwood is the product of specific pathogenesis involving both the host tree and a complex community of fungi and microorganisms introduced inside the stem after its natural or artificial wounding. Consequently, the phenomenon of agarwood formation and the biosynthesis of its constituents are more difficult to characterize than in the case of secondary metabolites produced only by a single organism. The considerable financial interest behind the sustainable supply of high-quality agarwood samples still motivates very active investigations in many fields (agronomy, microbiology, and chemistry), in order to discover faster and more efficient methods to induce agarwood production. Different factors influence the quality of the final product, including the age of the tree, the type of induction (e.g., natural or artificial), the pedoclimatic conditions, and the nature of the microbial community introduced into the stem, etc. However, one of the key parameters is the genotype of the host species [31], and agarwood phytochemistry strongly depends on the botanical identity of the producing trees.

2.1 Botanical Aspects of Agarwood-Producing Species To date, the recognized agarwood producing trees belong to various species included in seven genera within the Thymelaeaceae family. These genera are the following: Aquilaria, Gyrinops, Gonystylus, Aetoxylon, Enkleia, Wikstroemia, and Phaleria [13] (Plate 5). Aquilaria is the most important in terms of agarwood production, followed to a lesser extent by Gyrinops [32]. Aquilaria and Gyrinops are closely related taxonomically and often confused, and the two genera are in fact paraphyletic (sharing the last common ancestor) [33]. In comparison, the other genera mentioned above each have much lesser importance. For example, the wood of some Gonystylus species is sometimes sold in Malaysia under the same name as agarwood from Aquilaria [34] and Aetoxylon sympetalum is the source of “white oud”, a low-cost substitute and adulterant of agarwood essential oil (see Sect. 3.1.2). The Aquilaria species are distributed mainly in the Indomalesian region, in an area that can be roughly defined as an ovoid region spanning from Bangladesh and

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Plate 5 A–E Aquilaria crassna trees in an experimental plantation of French Guyana. A Five years old trees reach about 10 m in height. B Mature leaves are dark green with a shiny, elliptical cuticle. C The flowers are small, yellow, and clustered in hairy inflorescences. D, E The fruits are two-valved capsules measuring 3–5 × 2–4 cm, with one to two seeds. Photographs courtesy of C. Zaremski, Université de Guyane/CIRAD

Bhutan to Papua New Guinea for the northwestern and southeastern limits, Taiwan, and the south of mainland China (mostly Hainan) for the northern edge, and Sumatra, Java, and the Sunda islands in the south [35]. Up to now, 21 Aquilaria species are recorded, and among these, 13 are renowned resin-producing species. Concerning the genus Gyrinops, nine species are accepted, with three of these being recognized agarwood producers [32]. The potential to generate agarwood has yet to be investigated for the remaining Aquilaria and Gyrinops species [32]. The botanical names and geographical locations of the members of these two genera are summarized in Table 1. Today, A. malaccensis is retained as a type specimen for the Aquilaria genus [32]. Also, A. malaccensis and A. agallocha are considered the same species by most authors [32], even if this point is still a matter of debate [25]. Many old sources used the name A. agallocha; in the current chapter, both terms will be employed to be in line with the nomenclature used in the cited reference. In general, several discrepancies still remain concerning the taxonomy of the Aquilaria and Gyrinops genera, as the botanical delineation of their species is rather challenging. In the wild, the trees rarely possess fruits and flowers, and the morphological differences based on other criteria (like leaf morphology) are generally very limited [32]. The

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Table 1 Distribution of Aquilaria and Gyrinops species Species

Natural distribution

Cultivation placesb

Aquilaria apiculata Merr.

Bukidnon (Philippines)

–

Aquilaria baillonii Pierre ex. Lecomte

Cambodia

–

Aquilaria banaensis P. H. Hô

Bana Hills, Vietnam

–

East Malaysia, Brunei, and Kalimantan of Indonesia

–

Aquilaria brachyantha (Merr.) Hallier f.

Cagayan (Philippines)

–

Aquilaria citrinicarpa (Elmer) Hallier f.

Mount Urdaneta (Philippines)

–

Aquilaria crassna Pierre ex Lecomte

Cambodia, south of Laos, north of Thailand, and Cochinchina, Vietnam

Widely spread

Aquilaria cumingiana (Decne.) Ridl.

Indonesia, Mindanao (Philippines)

–

Aquilaria decemcostata Hallier f.

Laguna (Philippines)

–

Aquilaria filaria (Oken) Merr.a

Indonesia, Philippines, West Papua of Indonesia, and Papua New Guinea

Home gardens of East Indonesia

Aquilaria hirta Ridl.a

South Thailand, Northeast and South of Peninsular Malaysia, Singapore

Home gardens of Peninsular Malaysia and West Indonesia

Aquilaria beccariana

Tiegh.a

Aquilaria khasiana Hallier f.a Khasi and Meghalaya (Northeast India)

–

Aquilaria malaccensis Lam. a (syn. Aquilaria agallocha)

Bangladesh, Bhutan, Assam (Northeast India), and Sumatra and Kalimantan (Indonesia), Malaysia, Myanmar, southern regions of the Philippines, Singapore, South Thailand

Peninsular Malaysia, West Indonesia, Bangladesh, East India

Aquilaria microcarpa Baill.a

Johor (Peninsular Malaysia) Singapore, Borneo Island

Borneo

Aquilaria parvifolia (Quisumb.) Ding Hou

Camarines (Philippines)

–

Aquilaria rostrata Ridl.a

Peninsular Malaysia

–

Aquilaria rugosa K. Le-Cong Kontum (Vietnam), North & Kessler Thailand

–

Aquilaria sinensis (Lour.) Spreng.a

China, (mainly South, Hainan Island, Hong Kong), Taiwan

China

Aquilaria subintegra Ding Houa

South Thailand

South Thailand, Peninsular Malaysia (continued)

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Table 1 (continued) Species

Natural distribution

Cultivation placesb

Aquilaria urdanetensis (Elmer) Hallier f.

Mount Urdaneta (Philippines)

–

Aquilaria yunnanensis S.C. Huanga

Yunnan (China)

–

Gyrinops caudata (Gilg) Domke

Papua New Guinea

–

Gyrinops decipiens Ding Hou Indonesia

–

Gyrinops ledermanii Domkea Indonesia, Papua New Guinea

–

Gyrinops mollucana (Miq.) Baill.

Indonesia

–

Gyrinops podocarpa (Gilg) Domke

Indonesia

–

Gyrinops salicifolia Ridl.

Indonesia, Papua New Guinea

–

Gyrinops versteegii (Gilg) Domkea

Indonesia, Papua New Guinea

–

Gyrinops vidalii P. H. Hô

Laos

–

Sri Lanka

–

Gyrinops walla a

Gaertn.a

Acknowledged agarwood-producing species. b — Not cultivated so far. Data taken from [32]

botanical identification based on molecular biology techniques [33, 36–39] can be useful for living specimens, but is often difficult, if not impossible, for old samples of wood [40]. Therefore, the determination of the actual species of the parent tree of a given agarwood sample is still an extremely complex task. The development of metabolomic approaches based on the study of samples of secured origin would be of great help in this field. A consequence of the almost impossible traceability of the botanical species is seen in the behavior of the consumers and the traders for whom the botanical identity is not considered any longer as a key criterion. For instance, even if the market in the Middle East is supplied mainly by agarwood from Malaysia and Indonesia, the so-called “Cambodian” agarwood is the most popular type, although it generally has no link to this geographical origin. So the attributed provenance works currently rather like a “brand name” to refer to specific agarwood samples [3]. This situation blurs information on the actual origin of agarwood and consequently may affect the reliability of some bona fide phytochemical studies based on commercial samples. The most important species exploited by mass cultivation for agarwood production are Aquilaria malaccensis and A. crassna [32] and their traditional centers of production are the northeast of India and Bangladesh, Indochina (Laos, Vietnam, Cambodia, and Thailand), Malaysia, and Indonesia [41]. It has been estimated that currently more than 10,000 distillation units are functioning in Assam (India), supporting up to 200,000 people [41]. Besides these two classical species, the cultivation of A. sinensis in the south of China is a more recent trend that has been developed in order

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to meet the booming Chinese agarwood market [42]. In addition, the high incomes generated by the agarwood trade have stimulated the emergence of new production regions like Sri Lanka, Papua New Guinea, and even Australia and French Guyana [41, 43, 44]. Useful information on the trade and utilization of agarwood in various regions of the world is available through a number of TRAFFIC reports [3–5, 45–47].

2.2 Natural and Artificial Agarwood Formation It is common knowledge that the natural infection of Aquilaria trees is induced by physical wounding due to various reasons (lightning strikes, wind, and damage by animals (elephants) or wood-boring insects (ants)). The external fungi and microorganisms can then enter inside the stem through the wounds, and develop a parasitic infection that spreads gradually in the heartwood [48] (Plate 6). Few statistical data are available concerning the rate of naturally infected trees among the wild Aquilaria populations, which is certainly highly variable and dependent on the region. According to Paoli et al., some forests host only healthy trees while others are characterized by a high rate of infection (25–100%) [49]. There are no obvious external signs that a tree may contain agarwood, and if it does, the amount can only be fully determined after the tree has been cut down and opened. Therefore, many healthy trees have been destroyed by agarwood hunters, and such indiscriminate fellings are a major cause of the depletion of wild populations of agarwood-producing trees. To overcome this problem, non-destructive detection techniques based on acoustic methods have been proposed [50].

Plate 6 A Aquilaria agallocha Roxb. (syn. A. malaccensis) trees from a plantation in Moulvibazar district, Bangladesh. B To induce fungal infection, the trees are nailed with iron pegs. Printed with permission from Rahman M, Nath NM, Sarker S, Adnan MD, Islam M (2015) Management and Economic Aspects of Growing Aquilaria agallocha Roxb. in Bangladesh. Small-scale Forestry 14:459. C Wild Aquilaria crassna tree from Khao Yai National Park (Thailand). The tree was illegally damaged by poachers to induce agarwood production (dark area at the bottom). Photograph Blaise Droz, Wikimedia Creative Commons

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The exact identity of the microorganisms and fungi participating in the production of agarwood remains to be clarified. Various authors have reported a large number of fungal species as being present in the stems of agarwood-producing trees. These belong to the following genera: Acremonium, Aspergillus, Botryodyplodia, Botryosphaeria, Chaetomium, Cladosporium, Cochliobolus, Cunninghamella, Curvularia, Cylindrocladium, Diplodia, Epicoccum, Fusarium, Hypocrea, Lasiodiplodia, Massarina, Megacapitula, Menanotus, Nigrospora, Paecilomyces, Paraphaeosphaeria, Penicillium, Periconia, Phaeoacremonium, Philalophora, Phomopsis, Preussia, Pythium, Rigidoporus, and Trichoderma, to name just some of these taxa [31, 43, 51–53]. Apart from fungi, bacterial endophytes were also identified in seven Malaysian Aquilaria species [54]. This long list of potential (and established) agarwood inducers is certainly far from exhaustive, and many efforts are currently ongoing to better characterize these populations. Whatever their identity, it seems that the role of agarwood secretion is mainly defensive, as Mohamed et al. have demonstrated that the production of agarwood oleoresin induces a quick decline of the fungal abundance over time [55]. The high financial interests behind agarwood production have stimulated many investigations aimed at finding a cost-effective process for the artificial induction of agarwood secretion in healthy trees. Hence, as early as 1929, such studies were conducted in Assam, India [48], and were followed by numerous other investigations leading to many publications and patents. The techniques for the artificial induction of agarwood are based on the physical wounding and/or the introduction of chemical agents or microorganisms into the tree and have been addressed in several reviews [28, 29, 31, 56, 57]. This active topic is outside of the scope of the present contribution, but it is clear that the future of agarwood production lies in the selection of a proper induction technique or techniques. Obviously, the determination of the optimum methods must be guided by careful control of the agarwood quality, based on rigorous chemical analysis and on a good knowledge of the key markers of quality, namely, the bioactive constituents and key odorant principles.

3 Agarwood Composition When considering the two main families of agarwood constituents, i.e., sesquiterpenoids and chromones, the former are obviously important contributors to its fragrance since their volatility means that they often play a key role in the odor of many natural raw materials. However, chromones also participate in the generation of the typical odor of heated agarwood, because even if they have much lower volatility than the sesquiterpenoids, they are thermally unstable and produce volatile aromatic substances on heating. It is, therefore, necessary to review both these groups of metabolites in order to understand the chemistry of agarwood odorants. Since their characterization requires different analytical techniques, agarwood sesquiterpenoids and chromones have been investigated by different research groups, so the

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phytochemical advances on these two compound classes have been made somewhat independently of one another. In this contribution, an initial section on agarwood volatiles will describe the chemistry of specific sesquiterpenoids and other volatile constituents, followed by a presentation of the global composition of agarwood volatile fraction determined by GC-based studies. Then, the structures and properties of the main chromones identified in agarwood will be discussed, to better understand their potential role as precursors of agarwood odorants.

3.1 Agarwood Volatiles As for all aromatic plants, the fact that agarwood is an odorant is an obvious indication of the presence of volatile constituents. Resin-rich agarwood samples are often odoriferous, but old pieces generally require scratching and rubbing, or light heating, to produce a noticeable smell. The oldest practice to fully benefit from agarwood odor is to burn it as an incense, but an alternative way is to produce agarwood essential oil by hydrodistillation of the wood. The “Ain-i-Akbari”, an administrative account of the sixteenth-century Mughal empire, is one of the first written records mentioning the preparation and use of an agarwood essential oil for perfume formulation [1]. Nowadays, an important part of agarwood production is hydrodistilled to furnish agarwood essential oil for the fragrance industry. According to the first account of the technical aspects of its preparation [48], the essential oil yield is proportional to the resin content and rate of infection, with values ranging from 0.1% for slightly infected wood to 3.5% for black sinking wild agarwood. Non-infected wood from healthy trees older than 25 years can nevertheless produce an essential oil in yields of around 0.05–0.1% [48], but its composition is radically different from that of agarwood essential oil. Hence, Jain et al. showed that an essential oil prepared from non-infected wood contained mainly hydrocarbons as well as elemental sulfur [58] and later, Tamuli et al. reported that wood samples of A. agallocha healthy trees produce an essential oil composed mostly of fatty acids (tetradecanoic, pentadecanoic, and palmitic acids (1–3)) [59] (Fig. 1). Even if chemically and olfactorily, the essential oil prepared from the wood of healthy Aquilaria trees is totally different from agarwood essential oil; it is nevertheless a commercial byproduct of the agarwood industry and is called “Indian boya” [60]. If we compare the distillation yields and the compositions of agarwood essential oil and Indian boya, it is apparent that most of the sesquiterpenoids are actual constituents of agarwood oleoresin, and result from the infection, while healthy wood contains only traces of these metabolites. With very few exceptions [43, 58, 59], all of the phytochemical investigations on agarwood were made exclusively on samples distilled from the wood of infected Aquilaria trees. However, the occurrence of large amounts of 1–3 and smaller fatty acids in agarwood essential oil may be indicative of oils prepared from poorly infected woods. Nowadays, this practice is common among agarwood essential oil producers, because of the rarity and the increasing prices of good quality agarwood pieces. In fact, the most

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Fig. 1 GC–MS chromatograms. A Standard agarwood oil (A. malaccensis from India) B Indian boya

resinous agarwood samples and the largest pieces are processed into wood chips to be used as incense, while the lowest quality and the sawdust are generally distilled for agarwood essential oil production [60].

3.1.1

Structural Analysis of Agarwood Volatiles

The initial NMR-based studies on agarwood sesquiterpenoids were conducted in the late 1950s by the group of Bhattacharyya in Pune, India. They first described the isolation of “agarol”, an optically active sesquiterpene alcohol, for which the structure proposed was probably erroneous since it was never later confirmed [61]. In subsequent publications, they reported on the presence of tetrahydrofuranoid eudesmanes, the so-called agarofuranes (4 and 5), and their dihydro-, hydroxy-, and nor-keto derivatives (3–6) [62, 63] (Fig. 2). At that time, the availability of wild agarwood allowed the possibility of working on impressive quantities, and in these investigations, a total of 600 kg of wood material was extracted, to provide the purified samples of 4–9 after distillation and fractionation “50 to 100 times” by chromatography on alumina [64]. On considering an average 2021 price of 3000 e per kg for a medium quality agarwood [28], such an experiment would now require an astonishing investment of more than 2 millions euros. The structures proposed initially for 4–6 were actually wrong, and were corrected later by synthesis studies [64–66]. Bhattacharyya et al. also described the first

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O

O

4 (α-agarofuran)

O

O

O

8

7

6

5 (β-agarofuran)

HO HO

O

HO

O 9

O

O O

O

HO

O

O

O

OH

12 (agarol)

10 (agarospirol)

14 ((+)-dihydrokaranone)

13 (gmelofuran)

15 ((+)-karanone)

O OH

OH OH 16 (kusunol)

OH 17 (jinkoh-eremol)

OH 20 (epi-γ-eudesmol)

19 (iso-agarospirol)

18 (oxo-agarospirol) OH

OH

21 (jinkohol)

22 (jinkohol II)

Fig. 2 Agarwood sesquiterpenoids identified in the period 1960–1990 [62, 63, 67, 74–79]

O

O

O

O

O

O O

a)

O

b)

O

c)

O O

O

O d)

R

e) O

R = CH3 R = OH R = OCH3

O k) l) m)

j)

OH 11 ((–)-vetivone)

i)

O

f) g)

h)

O O

O

O

O

10

Scheme 1 Syntheses of (−)-agarospirol (10) and (−)-β-vetivone (11) [72, 73]. Reagents and conditions: a) O3 , Me2 S; b) KOH 5%, 80°C; c) H2 , Pd/C; d) Br2 , NaOH; e) CH2 N2 ; f) 3N HCl, DME; g) chromatographic separation of epimers (alumina); h) Ph3 P = CH2 ; i) pTsOH, benzene, reflux; j) MeLi; k) Ac2 O, AcONa; l) Na2 CrO4 , Ac2 O, AcOH; m) BF3 , Et2 O

example of a sesquiterpenoid bearing an “agarospirane” skeleton, through the isolation of (−)-agarospirol 10 [67]. The name of this peculiar skeleton later evolved as “spirovetivane” [68], then “vetispirane” [69], when other sesquiterpenoids of this family were discovered in vetiver [70, 71]. As mentioned by Näf [2], the relative and absolute configuration of 10 were often represented incorrectly in the literature, but

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were established unambiguously by the synthesis of (−)-β-vetivone 11 via 10 [72, 73] (Scheme 1). After a 15-year-latent period, phytochemical investigations on agarwood resumed in the 1980s, with the isolation of the furanoid cadinane sesquiterpenoids agarol (12) and gmelofuran (13) [74]. It should be mentioned here that despite their same trivial name, 12 is totally unrelated to the agarol of Bhattacharyya [61]. From then on, major advances were made by Japanese workers through the identification of additional eremophilane sesquiterpenoids such as (+)-karanone (15) and (+)-dihydrokaranone (14) [75], as well as jinkoh-eremol (17), and its isomer, kusunol (valerianol) (16) [76]. The same Japanese group also identified (−)-oxoagarospirol (18) and (−)iso-agarospirol (19) and two vetispiranes related to 10 [75]. Compound 18 was simultaneously discovered by Yang and Chen in A. sinensis and named baimuxinal [77], but both names appear in the literature (with two different CAS numbers) as they were initially considered as two different compounds. This confusing situation is due mainly to the difficulty in representing the stereochemistry of vetispiranes in two dimensions, and Nakanishi and Näf mentioned that oxoagarospirol and baimuxinal are probably identical [2, 78]. A plausible biogenetic link between the eremophilane, eudesmane, and vetispirane sesquiterpenoids of agarwood was proposed when epi-γ-eudesmol (20) was identified in a sample of Indonesian agarwood [78]. According to its structure, 20 could be a logical precursor of 4–10 and 14–19 [76]. Investigations on this Indonesian agarwood sample furnished even two rare prezizaane sesquiterpenoids, (−)-jinkohol (21) and (+)-jinkohol II (22) [76, 79], and their complex tricyclic skeleton is also suspected to be related biogenetically to the vetispirane structure [80], but this link has not been demonstrated in agarwood so far. The early 1990s saw significant advances in agarwood chemistry, initially through the contributions of the Ishihara group. In a series of four articles, they reported on the presence of (nor)guaiane (18) [81–84], eremophilane [84], and eudesmane [84] sesquiterpenoids (23–38) in a sample of A. agallocha from Vietnam, together with compounds already described previously in agarwood (Fig. 3). Many of these constituents were indeed identified for the first time as natural products, and the characterization of several of them was secured by total synthesis or semisynthesis, a valuable way also to confirm their olfactory properties. Hence, in addition to the important odorant 23 (see Sect. 4.1), (−)-34a/34b and (+)-35a/35b were prepared in three and four steps, respectively, by semisynthesis from (+)-βselinene 39 isolated from natural celery seed oil (Scheme 2). Moreover, as (−)-37 was already described as a synthetic product [85], Ishihara repeated the reported procedure for its preparation from (+)-carvone (40) and reduced it to its corresponding alcohol (+)-21 [86, 87]. In addition, (−)-17 and (−)-32 were semi-synthesized from (+)-nootkatone (41) via a mixture of (−)-aristolochene (42) and (−)-nootkatene (43), and 42 was used to prepare neopetasane epimer (44), which eventually gave (−)-14 after isomerization (Scheme 3). All these syntheses helped to ascertain the absolute configuration of the levorotatory enantiomers naturally present in their agarwood sample of origin. In the same

62

N. Baldovini O

HO

O

COOH

25

24

23

O

HO HO

O

O

O

O

27

26

28

O O

O OH 30

29

31 OH

R

HO

R

32

33

34a R = CHO 34b R = COOH

OH

O

O 36

35a R = CHO 35b R = COOH

38

37

Fig. 3 Agarwood sesquiterpenoids identified by Ishihara et al. [81–84]

a)

b)

+

+ HO

O OH

39 ((+)-ß-selinene) O

OH

OH

c)

40 e)

d)

f)

37

R

R

OH

O

34a R = CHO 34b R = COOH

d)

35a R = CHO 35b R = COOH

38

Scheme 2 Ishihara syntheses of eudesmane agarwood odorants 34a/b–35a/b 37, 38 [84, 85]. Reagents and conditions: a) mCPBA; b) 2% H2 SO4 , MeCN/H2 O 1/1; c) PCC, Celite, CH2 Cl2 , then separation by flash chromatography (SiO2 ); d) 10% aq. KOH, AgNO3 , EtOH; e) see Ref. [85]; f) NaBH4 , CeCl3

period, Näf et al. characterized many additional constituents after a detailed analysis of a commercial agarwood essential oil produced in India: (nor)eudesmanes (45–52) [88, 89], (nor)eremophilanes (54–58) [89, 90], spirovetivanes (59–62) [90], and a linear compound (53) [89] (Fig. 4). Here again, the identification of several compounds (46–44) was confirmed by synthesis.

The Chemistry of Agarwood Odorants

63 O

O

g)

ent-14

44 e) f) O

RO

AcO a)

b)

c)

+ 43

42

R=H R = Ts

41 ((–)-nootkatone)

d)

+ OH

OH 32

17

Scheme 3 Ishihara syntheses of eremophilanes ent-14, 17, 32 [84]. Reagents and conditions: a) isopropenyl acetate, TsOH; b) NaBH4 ; c) NaBH4 , DMF; d) HgO, pivalic acid, HClO4 , ultrasound, then NaBH4 , 10% aq. NaOH; e) O2 , hν, then NaBH4 ; f) PCC, Celite, CH2 Cl2 ; g) TsOH, THF

O

OH 46

45

47

49

48

O O O OH 50

O

O

O O

51

53

52

54 O

O

O

OH

OH

55

56

58

57 O

O

HO

60

61

Fig. 4 Agarwood sesquiterpenoids identified by Näf et al. [88–90]

62

59

64

N. Baldovini OH OH

O

OH

COOCH3 HO

OH

HO 63

OH O

64

HO

OH O

O

O

O O

69

O

O

O

O

70

71

O

HO

67

O

O

OH

OH

66

O

OH OH 68

65

O

HO

OH

OH

OH

72

HO

OH O

OH

OH HO

OH 73

75

74

76

77

OH O

O

O

OH

COOCH3 OH

OH 80

81

OH

O

HO COOCH3 O

O O

83

82 O

O

OH O

OH

OH 79

78

O O OH

OH

84

85

86

87

OH O HO O HO

O

O

HO 88

O

OH

89

O 90

91

92

Fig. 5 Agarwood sesquiterpenoids identified since 2006 [17]

From this period to the present, the number of new sesquiterpenoids reported from agarwood has increased significantly, due mainly to the contributions of Chinese groups working on A. sinensis and other species. Most of these new compounds, isolated by fractionation of solvent extracts by liquid chromatography, are polyfunctional sesquiterpenoids (di- and triols, hydroxylated esters, aldehydes, and ketones). Therefore, they are probably too hydrophilic and not volatile enough to be efficiently hydrodistilled, which would explain why they were not reported previously as agarwood essential oil constituents. The exhaustive listing of these compounds is out of the scope of the present chapter, but can be found in the review of Li et al. [17], which, in turn, could be updated with 21 additional new molecules described in very recent

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publications [91–94]. To illustrate the structural diversity of all the new agarwood sesquiterpenoids discovered during the last 15 years, selected examples are provided in Fig. 5. As expected, many belong to well-known structural classes, i.e., eudesmanes (63–65) [94–96], eremophilanes (66–67) [94, 95], guaianes (68–72) [92, 97–101], prezizaanes (73–74) [102, 103], and cadinanes (75) [104]. However, more surprisingly, other types are represented: zizaanes (76–77) [102], acoranes (78) [97], humulanes (79–81) [93, 99], an elemane (82) [94], and a rotundane (83) [91], and some of these compounds display degraded, rearranged, and other uncommon skeletons (84–92) [17, 93, 94, 97, 99, 100, 105]. The possibly non-terpenoid biogenetic origin of 91 [17] deserves attention, as does the puzzling structure of the new sesquiterpene 92 with a C 3 symmetry axis [97]. The fact that many new compounds are being discovered continuously is consistent with the results of a study by Wong et al., in which GC × GC-TOFMS analysis of A. malaccensis agarwood essential oil samples revealed the presence of at least 550 constituents, among which only a small fraction could be identified definitively [106]. Therefore, it can be concluded that many minor constituents are still unknown among the agarwood volatiles.

3.1.2

General Composition of Agarwood Volatiles

In parallel with work devoted to the structural analysis of isolated compounds, many studies have afforded a more general picture of the global composition of the agarwood volatile part, by describing the GC profiles of agarwood essential oil or solvent extract. This phytochemical information, which includes quantitative aspects (relative amounts of the constituents), is highly useful to compare the different agarwood varieties and establish their specificities. It is also a prerequisite to investigate the relationship between the chemical components and the olfactory properties of these samples. However, such GC analytical procedures need to rely on a rigorous methodology being conducted, both for the qualitative and quantitative characterization of the constituents present. The development of easy-to-use software with computerized commercial mass spectra (MS) databases has brought significant progress in the interpretation of GC–MS analytical data. However, it also led to a high number of careless analytical practices, and eventually to publications biased with compound misidentifications. Such a situation has brought a good deal of confusion to the field of agarwood chemistry and has misrepresented the actual compositions of agarwood samples. Consequently, the general background phytochemical knowledge required to control their quality and authenticity has become somewhat compromised. Hence, in the long term, there is a risk of lowering the quality of commercially available agarwood products, resulting in lower demand from consumers. On the basis of the International Organization of the Flavor Industry (IOFI) requirements, several criteria have been proposed [107] to define the correct analytical practices for the identification of volatile constituents of natural extracts and essential oils. A discussion of these criteria is given below, with a focus on those specifically concerning agarwood.

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Nowadays, the authenticity of the analyzed material is one of the main issues in agarwood analysis, since the scarcity of wild samples has led to a strong increase in their price, and consequently to a higher risk of fraud and then to the agarwood market becoming flooded with adulterated and fake samples. The situation was less problematic at the time of the first detailed investigations on agarwood volatiles during the 1980s because the raw plant material was more generally available. Nevertheless, even then, many analyses were performed on samples purchased from secondary (at best) suppliers in the traditional agarwood trading hubs (e.g., in Singapore, Hong Kong, and Bangkok). The reliability of their claimed geographical origin consequently may be questionable, and the same is true for the botanical identity of the trees from which the samples were taken since the distinction of the species by a simple visual examination is extremely difficult. Nevertheless, the generally high quality of the analyses reported in the period 1980–1995 period affords a good picture of the volatile composition of agarwood available on the market, even if the earliest investigations were not able to identify many components discovered later by Firmenich and the group of Ishihara [81–84, 88–90]. In the publications dealing with agarwood chemistry, many constituents common to other well-known aromatic plants have been identified. The question of their actual origin is crucial, since their presence may also result from sample mixing, a classical issue with expensive commercial products like powdered agarwood and agarwood essential oil. For instance, the presence of patchoulol (93) and α-gurjunene (94), occasionally reported in agarwood, may actually originate from contamination or adulteration with patchouli and gurjun balsam (Dipterocarpus turbinatus [108]), respectively (Fig. 6), since both species are endemic in some of the agarwoodproducing areas. Similarly, Aetoxylon sympetalum is another member of the family Thymelaeaceae, growing in Malaysia and Indonesia, where its vernacular name is “Gaharu buaya”. Like Aquilaria species, A. sympetalum trees produce a fragrant resin when their trunk is damaged and infected. The essential oil obtained by the distillation of infected A. sympetalum wood has a typical smoky and sweet woody aroma, albeit totally different from the agarwood essential oil odor, but is sometimes

HO OH 93

94

OH

OH

96

95

97

O

OH O

COOR

O

O O

O

COOR 98

99a R = Et 99b R = n-Oct

100

O

O

OH 101

OH O O

O OH 102

Fig. 6 Potential and obvious agarwood adulterants or misidentified products

103

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named “white oud”. It is believed that A. sympetalum volatile oil samples are used occasionally to adulterate agarwood essential oils [109] since they are distilled in the traditional agarwood-producing countries and their price is generally much lower than authentic agarwood essential oil. The composition of A. sympetalum volatiles has never been published so far, but an analysis of several commercial samples has revealed that they are dominated by the presence of γ-eudesmol (95) [110]. Therefore, the presence of an abnormally high amount of this compound in an agarwood essential oil sample should be considered with caution, since it may be indicative of adulteration with A. sympetalum oil. The occurrence of small quantities of α- (96) and/or β-santalol (97) in A. sinensis and A. crassna has been also reported independently by several research groups [111–114]. Compound 96 was also identified in a sample of commercial jinkoh powder for use in kôdô purchased in Japan [110]. Among agarwood components, the patchoulane and aromadendrane skeletons of 93 and 94 are very rare, and, similarly, the occurrence of the santalanes 96 and 97 is puzzling, as no other sesquiterpenoids of this group are present. Contamination with sandalwood powder (another key kôdô ingredient [7]) in commercial jinkoh is plausible, but the identification of 96 and 97 in agarwood essential oil and in cultivated and grafted agarwood deserves to be confirmed by further phytochemical investigations based not solely on GC/MS experiments. The actual occurrence of 93, 96, and 97 in agarwood would not only provide a new perspective on the biogenesis of agarwood volatiles but also would be very relevant for a better understanding of the key odorants of this material. Indeed, 93, 96, and 97 are the main odorant principles of patchouli and sandalwood, and it would be interesting to determine if they also are included as components of the complex fragrance of agarwood. The olfactory properties of the constituents identified in agarwood should also be used as a clue to confirm or refute their actual presence in a given sample. For instance, an unusually high percentage of 4.5% of the very potent pepper odorant, rotundone (29) [115], was reported in a Malaysian agarwood essential oil [60], which should therefore display a totally distorted olfactory profile. The same is true for the potent musky odorant cyclohexadecanolide (98), claimed to be a constituent at a content of 3% in an agarwood essential oil from Laos [114], and at an inconsistent retention index [116]. In comparison with wild agarwood, it is generally easier to secure the authenticity and the traceability of cultivated agarwood when the sampling is done in close collaboration with the producers. However, detailed analyses of cultivated samples often reveal the presence of non-natural constituents that may come from the inoculation protocol or the processing of the wood and/or its adulteration. Indeed, some non-natural pollutants are frequently encountered even in “good quality” samples of agarwood and agarwood essential oil [110]. For instance, cosmetic products, plasticizers, solvents (phthalates (99), isoamyl laurate (100), triethylcitrate (101), dipropylene glycol (102)), and antioxidants such as di-tert-butylphenol (103) are sometimes reported in agarwood compositions. The presence of 99a, 99b, 101, and 102 in commercial agarwood essential oil samples was pointed out as a possible sign of multiple adulteration steps, as the product passes from one retailer to another and is “diluted” with another solvent to increase profit [117]. Diethyl phthalate (99a)

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is a very common non-natural compound present not only in agarwood essential oil and occurs also sometimes in commercial wood samples [110]. It can only be wondered whether or not this cheap, non-volatile, odorless, and rather dense (d = 1.12 g/cm3 ) adulterant is used to impregnate the wood in order to increase its density, and then eventually its retail price. Unfortunately, the high cost of authentic agarwood stimulates the appearance of low-quality “agarwood” and its substitutes, like poorly resinous agarwood coated with agarwood powder mixed with wax or varnish, or even substitute wood impregnated with a tincture of low-quality agarwood essential oil. The latter product is popular in the Middle East under the name “black magic wood”, and is sourced from Indonesia and Malaysia [3, 118]. Many other additives are certainly used to adulterate and imitate agarwood and agarwood essential oil, and a monitoring based on chemical analyses would be helpful to promote a healthier marketplace in the field of agarwood products. A report of uncommon synthetic compounds in the phytochemical literature on agarwood may also be the result of a wrong interpretation of the GC–MS data, based only on the MS information. Indeed, it is now common to consider that a valid identification requires two orthogonal criteria, generally retention index (RI) and MS [107]. The proper use of reliable MS and RI databases is crucial for the correct determination of a list of components, and the co-injection of authentic reference compounds is highly advisable. Most agarwood constituents are specific to this material and are often not present in general commercial databases. Therefore, misidentifications and errors frequently occur if the analyst neglects the critical examination of the “best” hits proposed by the MS interpretation software. The EI-MS data reported in the literature on agarwood [78, 79, 81–84, 88–90, 119–121] are highly valuable to build databases for an accurate interpretation of the GC–MS results. However, as mentioned by Näf [2], the direct use of these data requires some attention since many of the MS spectra were recorded at 20 eV, and accordingly are not totally analogous to those acquired at 70 eV. Nevertheless, the 70 eV MS spectra and the RI of the major agarwood alcohols (10, 16–17) are fully accessible in the literature [122, 123] and can then be taken into account for their identification. In addition, it is also the duty of the analyst to consider the plausibility of the constituents proposed by the interpretation software used. Some claimed agarwood constituents are clearly non-volatile, synthetic, or of dubious natural origin (sugars and derivatives, nitro compounds and other nitrogen-containing molecules, iodinated and halogenated compounds). Among other major pitfalls that have been made, several papers on agarwood mention the absolute configuration of certain chiral constituents when determined by GC analysis on a classical column, but such characterizations are impossible unless a column with a chiral stationary phase is used. The quantification of constituents is another issue that has to be considered carefully, especially when the results are used for classification purposes on multiple sample analyses. Quantifications should be determined by GC-FID, rather than TIC MS [124]. Preferentially, they should be conducted by internal standardization, and the method developed by Cicchetti et al. [125, 126] is reliable, affordable, and easy to implement in simple GC-based analytical procedures. Moreover, in the context of agarwood essential oil analyses, such a quantification approach is highly useful since

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69

adulterations with non-volatilizable constituents are frequent, and solvent extracts naturally contain non-volatile substances or semi-volatile ones (chromones) that do not appear systematically in GC profiles. Sampling by solid-phase microextraction (SPME) sometimes has been used for the analysis of agarwood smoke or of the headspace above heated agarwood. Recommendations for the proper practice of this technique also exist [127], and it should be recalled that it is not adapted for the quantification in complex mixtures, since for a given constituent, the extraction yield depends on its affinity for the SPME fiber. Therefore, there will be strong variation from one compound to another, and consequently, the SPME sampling procedure is not representative of actual headspace composition [127]. All these considerations were taken into account in the critical evaluation of the literature proposed in the present chapter, and a global overview of the composition of the volatile part of agarwood was produced from carefully selected articles, and is developed hereafter and displayed in a graphical manner in Fig. 7. Many publications dealing with agarwood compositions were excluded voluntarily from this overview when it appeared that the analytical procedures used did not meet the criteria cited above. The collection of information eventually included for this overview, however, should be interpreted with caution, as it is difficult to compare analyses achieved with

Fig. 7 Agarwood constituents reported among the main volatiles. Upper line: composition number, Botanical species (when available), (geographical origin of the sample), year of publication. Lower line: type of sample (solvent extract or essential oil (EO); (*commercial EO) (%: total amount of volatiles reported in the chart)

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Fig. 7 (continued)

different sampling procedures, equipment, and analytical practices. The pie charts of Fig. 7 are provided to give at a single glance a rough overview of the compositions of the volatiles, together with the relative amounts of the different constituents. For the sake of clarity, only the compounds appearing at a percentage higher than at least 1% in at least one study were taken into account. For better visualization, the contribution of the unknown compounds is not included in the pie charts, but the total percentage of the identified compounds appearing in the charts is reported in brackets in the legend. This value was obtained by summing the percentage reported in the source article and should be considered as much lower than the actual amount in the case of solvent extracts, which also contain non-volatile constituents not eluted by GC. Even if they are sometimes eluted using GC, the constituents of poor volatility (chromones) were excluded as their direct participation in the resultant odor is considered as not significant. For this reason, the proportion of identified compounds in agarwood essential oil samples (Charts C5, C8–C15) was generally higher than in their solvent extracts (C1–C4, C6–C7). Of course, unknown compounds also end up in this non-identified portion, and the early studies of the group of Yoneda et al. [128, 129, 133, 134] were very informative in understanding the variability of agarwood volatiles but were obviously unable to report several important compounds identified later in the 1990s. Therefore, this issue should also be kept in mind when comparing the different compositions on the basis of the pie charts, as displayed in chronological order in

The Chemistry of Agarwood Odorants

71 O

O

O 105

104

106

107

HO

OH 108

109

110

111

OH 112

O OH

OH

113

115

114

116

117

COOH

COOH 118

119

Fig. 8 Additional agarwood constituents

the figures. The relative proportion of the different sesquiterpenoid skeletons is also a highly useful piece of information to classify the samples on biosynthetic criteria. To display this aspect in a visual manner, a color code has been adopted, and this is represented at the bottom of Fig. 7, where the numbering of the main components is recalled. The structures of additional compounds 104–119 (not cited previously) are displayed in Fig. 8. The most striking aspect of agarwood chemistry to appear at first glance when considering the charts of Fig. 7 is the high variability of its volatile constituents. No clear pattern arises, where a given chemotype would be associated with a specific geographical origin or a single botanical species. This issue was observed very early on and was emphasized by the Japanese chemists who performed the first studies reported herein. As fine connoisseurs and traditional users of agarwood, Japanese sourcing experts were aware of the different agarwood varieties and the difficulty to link these with the botanical origin of the samples. Hence, Yoneda et al. indicated that two kinds of wood were present on the market, attributed as A. agallocha and A. malaccensis, two species considered later as identical [133]. They analyzed the composition of the benzene extracts of both varieties, purportedly collected in Vietnam and Indonesia, respectively, but purchased in Singapore [133]. This comparison was refined later by a more detailed examination of the compositions of 72 samples from various localities of Southeast Asia [128]. This type of study, assisted by principal component analysis and factor analysis [134, 135], led to the definition of three chemical types, corresponding to the respective representative compositions C1–C3 of Fig. 7. The first one (C1, 30% of the samples) contained prezizaanes

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21–22 as main compounds, the second type (C2, 49% of the samples) contained no prezizaanes at all, but instead vetispiranes and eremophilanes: often high amounts of 14 and 18, variable amounts of 10/16/17, and often (but not always) high amounts of eudesmane 20. The third type (C3, 21% of the samples) was similar to C2, except that it was devoid of 14. Here again, there was no certainty concerning the botanical identity or the actual collection place of the wild samples used in this study, since most were traded via Singapore or Hong Kong. However, taking into account their alleged origins, the trends were the following: samples from Java, Sumatra, and Malaysia were mostly of types C1 and C3, while samples from Thailand, Vietnam, Cambodia, and Borneo were of type C2. Nevertheless, a few mixed compositions were also described in this work, maybe due to sample mixing. To help the on-site agarwood analysis, the authors also developed a detailed TLC analytical procedure to distinguish these different types [136]. In addition, they also described the composition of two A. sinensis extracts from China, dominated by the vetispirane 18 (C4) [129]. These investigations were based on the GC–MS analyses of solvent extracts, and the first detailed study on the global composition of an agarwood essential oil stricto sensu (i.e. obtained by hydrodistillation of the wood from a single tree), appeared only in 2000 [108], when Yaacob and Joulain reported a composition C5, similar to C3 (but devoid of the vetispirane 10), in an agarwood essential oil prepared from A. malaccensis agarwood from Malaysia. The sample was analyzed by a combination of GC–MS and GC–FTIR, to pinpoint the ketones and aldehydes present. They observed the presence of large amounts of 104, which was due probably to the thermal degradation of chromones during the hydrodistillation process (vide infra), a phenomenon that is not able to occur by solvent extraction at a lower temperature. Ishihara et al. also contributed to the better understanding of the general composition of volatiles in agarwood [130], by analyzing acetone extracts of four Vietnamese agarwood samples graded according to the traditional Japanese classification: three samples of kanankoh (the highest grade, also named “kyara”), and one of jinkoh, of lower quality. Indeed, the extraction yields reflected well the proportion of resin and the differences in quality, since the kanankoh extracts were obtained with a 45% yield while jinkoh gave only 2.7% of the extract. Significant qualitative differences were also observed between the four samples, with two of these being represented in Fig. 7. The jinkoh sample (C6), tentatively identified as A. sinensis, contained mostly vetispiranes (especially 18) and eremophilanes (especially neopetasane 33, which is suspected to produce 14 by isomerization [84]). In contrast, eudesmanes 34a, 34b, 35a, and 38 and the guaianes 23, 24, 25, and 30 reported for the first time in this study were typical of one of the kyara samples (C7), attributed to A. agallocha. The detailed investigation by Näf et al. in 1995 on a commercial Indian A. agallocha oil represented another milestone in the understanding of the global composition of agarwood [90]. They reported, together with previously described compounds, several new additional eremophilanes (15, 55, 57) and vetispiranes (115, 116, 59, 60– 62), after careful fractionation of the oil by chromatography on silica gel and preparative GC. In addition to these new constituents, this agarwood essential oil (C8) contained the typical molecules observed in the other types (C2, C3, C5) as well as

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the aromatic compounds 104–105, with the latter also being plausible degradation products of chromones (see Sect. 4.2). From this date on, agarwood essential oil compositions reported afterward were in general characterized by the presence of the three classical groups of sesquiterpenoids (eremophilanes, eudesmanes, and vetispiranes) with variations among the nature and proportion of the main components within these compound classes. The aromatic compounds 104 and/or 105 typically were present in all samples, sometimes in high amounts (C11). Fatty acids (especially palmitic acid (3)) were also observed occasionally in high concentration levels (C15). Studies performed in the same laboratory on different samples are useful for comparison purposes, because they are in principle realized under identical analytical conditions and sometimes use the same procedure for sample preparation (for laboratory-made essential oils). An investigation by Nor Azah et al. [131] showed the variability of four A. malaccensis agarwood essential oil samples prepared from woods collected in different parts of the Malaysian peninsula. Three of these compositions are represented in Fig. 7 (C10–C12) and show high variability, both qualitative (eremophilane and eudesmanes) and quantitative (vetispiranes). On the other hand, the comparative investigation of six agarwood essential oil samples recently published by Tran et al. [114] showed a reverse tendency: one sample was distilled from Vietnamese A. crassna agarwood while the five others were commercial oils from India, Indonesia, Thailand, Laos, and Taiwan. Surprisingly, most compositions showed a roughly similar pattern, with only little quantitative variations among the three main groups of compounds (eremophilanes, eudesmanes, and vetispiranes). The differences among these six samples were mostly in the amounts of the fatty acids present (here again dominated by 3), either not detected in the laboratory-made agarwood essential oil, or amounting to 15% in the commercial Laotian sample (C14 and C15, respectively). A broadly similar composition was reported by Tissandié [132] after a detailed 2D GC–MS study of a commercial sample of agarwood essential oil of non-specified geographical origin (C13). In conclusion, there is a puzzling contradiction between the homogeneity of the compositions of commercial agarwood essential oil present in the current market and the variety of profiles reported in the literature from the initial studies. Hence, neither a composition rich in guaianes (C7) nor of the prezizaane type (C1) appeared again in the literature. However, small amounts of prezizaanes have been reported from time to time (for instance in C10), and recent studies on high-quality “Qi-Nan” samples of wild A. sinensis seem to indicate that they sometimes contain significant amounts of guaianes [137]. It appears that there is still a need for an in-depth analysis of the compositions of agarwood essential oils prepared from samples of certified botanical and geographical origin.

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3.2 Non-volatile Constituents of Agarwood If the history of agarwood chemistry is put in perspective, it may be noted that the early period of study was devoted mainly to the characterization of the components of its volatile fraction, mostly monofunctional sesquiterpenoids. However, over the last 15 years, the LC-based analysis of agarwood solvent extracts has helped to discover many new polyfunctional sesquiterpenoids that are poorly extractable by hydrodistillation. Another group of compounds, the chromones, is now becoming the source of an even larger number of molecules. The identification of the first representative of this family by Yoshii et al., agarotetrol (120a), is nonetheless from over 40 years ago (1978) [138] (Fig. 9). These authors were intrigued by the unprecedented structure of this compound, as the linkage of the chromone moiety with a phenylethyl group at position C-2 was radically different from those of the flavones, the largest family of natural chromones, widespread in most plant species. Nevertheless, this discovery apparently received very little attention, except from the group of Kiyosawa in Kyoto, who identified nearly 30 additional 2-phenylethylchromones in the decade 1982–1992 [139–150]. R7

R4 R3

O A

R8

R2 R1

R3

R6

B

R

R

2

R1

R3

R6

B

8

R

OH

R1

O

122

O O

O

OH HO

HO O

O

O

120a (agarotetrol)

O 120c

120b O

OH

O

O

O O

HO

HO

R6 R5

R2

Cl O

OH

O

5

121 (FTPEC) 121a: R7, R2 = H 121b: R7 = OCH3, R2 = H

120 (THPEC)

R7

R4

C

O A

R5

O

HO

R7

R4

C

O

O

O

O

O

OH

O

O

O

OH

O

120d HO

OH OH

O O

O

O 121c

120e

O

O

OH

O

O

O

O

HO O O

O 121d

O 121e

Fig. 9 Agarwood chromones (general formulas and monomers)

O 122a

The Chemistry of Agarwood Odorants

75

Shortly after the discovery of 120a, the prototypical compounds 121a and 121b were identified in agarwood [119, 151], and the characterization of other chromones in this material has accelerated over the last 10 years, to reach a total of 266 chromones reported in various Aquilaria and Gyrinops species so far. An exhaustive review of agarwood chromones was published by Li et al. in 2021 [17], and this may be complemented by 26 additional chromones reported in 2020 and 2021 [93, 152–156]. All these compounds can be classified roughly in two main types: Flindersia-type 2-(2-phenylethyl)chromones (FTPECs, 121), after the name “flindersiachromone” given to 121a isolated first from Flindersia laevicarpa [157], and their saturated analogs 5,6,7,8-tetrahydro-2-(2-phenylethyl)chromone (THPECs 120), with 120a being the first isolated representative. All positions (R1 –R7 ) on rings A and C, as well as one on the chain (R8 ), can be substituted, mostly by hydroxy and methoxy groups and sometimes by a chloro substituent. THPECs are often highly oxygenated in ring A, mainly by hydroxy groups with α- or β-stereochemistry, or alternatively with epoxy functions. In this last case, THPECs can be split into two subclasses: monoepoxy- or diepoxy-derivatives of THPEC (EPECs and DEPECs, respectively, identified only in agarwood so far). Finally, a few examples of glycosylated or acetoxylated FTPECs are known, as well as six examples of 2-(2phenylethenyl)chromones (122). Accordingly, statistically, a high number of possible variations in the positions and the stereochemistry of the substituents exist, and this explains the multitude of chromones reported so far. Selected examples (120b–120e, 121c–121d, 122a) of such compounds are represented in Fig. 9, by way of illustration. Another classical structural variation that increases considerably the potential number of chromones in agarwood is the possibility of having dimers and trimers, in which monomeric units are linked by one or two C–C or C–O bonds on ring A or C (123–127). Moreover, the group of Dai recently reported the existence of derivatives in which a sesquiterpenoid moiety is linked to the chromone with a covalent linkage (ester or ether), as in 128–129, and benzylacetone derivatives linked on ring A by C–C and C–O bonds (130) (Fig. 10) [158–160]. Given the high diversity of agarwood chromones and sesquiterpenoids, and the variety of possible linkages between these structures, one may wonder if the nearly 500 members characterized to date of these two groups represent only just the visible part of the iceberg. In the near future, new agarwood constituents will probably be identified, including maybe tetramers and higher polymers, covalently bound with various moieties (sesquiterpenoid and others). Moreover, from their number and their structural diversity, chromones are ideal substrates for the development of metabolomic approaches to propose analytical tools for the authentication and profiling of agarwood samples [161–166].

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O O HO O

O

O

O O

OH O

O

OH

OH

OH 123

124 O O

OH O

O

O HO

O

O

O

O

OH O

O O

O

O

O

O 126

125

OH O

OH O

O

O OH

OH O

O O

O OH

O

O

O

HO O

HO OH

OH OH

128

127

O O

O

OH O

O O

OH HO

O O

HO O

O

O 129

130

Fig. 10 Agarwood chromones (dimers, trimers, and combined compounds)

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4 Agarwood Odorants The specific odor of burnt agarwood has been described by Roman Kaiser, a renowned fragrance specialist from the Givaudan Company, as “one of the most fascinating scent sources of this world (…) a pervasive, mysterious, wonderful scent, bringing to mind the perfumes of all imaginable precious woods, balsams, and resins, as well as those of amber, musk, and castoreum and, somewhat hidden, even tender floral notes” [11]. Such a vibrant personal account of the olfactory character of this mythical material resonates with its prestigious status in Asia and the Middle East. As a result, when phytochemists started to investigate the chemical composition of this unique material, many of their studies were motivated by an interest in its odorant fraction, and the olfactory properties of its constituents were sometimes mentioned in their publications. However, as in many studies on the analysis of fragrant natural aromatic raw materials, they simply reported a description of the odor of some individual constituents isolated from the material, without clearly demonstrating if they are key contributors to the fragrance of the entire product. Moreover, the compounds evaluated were often obtained as partially pure samples by the fractionation of agarwood extracts. Thus, the olfactory character of a given constituent isolated from such a complex mixture should be considered with caution, because residual traces of unidentified potent odorants can significantly alter the odor of the sample and lead to the wrong conclusions. The parallel evaluation of synthesized samples is an excellent way to confirm the sensorial properties of an isolated compound, but it is seldom performed because of the significant additional workload it requires. Nevertheless, the characterization of the most important olfactory contributors to the odor of a natural mixture is a very complex task [167], which can be achieved definitively only by recombination experiments following a comprehensive analytical characterization of the sample. In the case of agarwood, reconstitutions are extremely challenging, because of the complexity of its composition, and the difficulty to obtain pure samples of its odorant constituents. This situation explains why the identity of the main key odorants of this prestigious fragrant raw material is still controversial, as no real consensus exists about typical agarwood odor markers. Of course, the chemical variability of agarwood considerably complicates this task, and indeed, a similar diversity exists in the olfactory character of agarwood samples, depending on their origin and their manufacturing method. First of all, a distinction should be made between the odors of agarwood solvent extracts, agarwood essential oils, and the particular smells of burned agarwood, which are all radically different despite the presence of constituents common to all such samples. The odor of agarwood solvent extracts is certainly the closest to the smell of agarwood resin, but the latter are seldom used and their commercial significance is so far very limited. Hydrodistillation of agarwood is the standard technique for the preparation of agarwood essential oil, and this process varies significantly from one producer to another. However, in all cases, chemical reactions necessarily occur when agarwood constituents are placed in boiling water for extended periods (sometimes

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for more than 48 h). Many hydrophilic components also simply dissolve in water, where they may eventually undergo thermal degradations such as 120a [168]. Microbial transformations in water are also possible, because a common practice among agarwood essential oil producers is to allow the powdered agarwood to macerate in water for some time (up to 3 months [41]). Depending on the duration of this step and the microbial population in the macerate, various fermentation reactions will occur and eventually modify the composition of the oil, but a rigorous analytical study to explore the effects of the fermentation is still required. In contrast, the traditional habit of burning agarwood produces other compounds, as the sample is brought in the presence of oxygen to a temperature higher than in classical hydrodistillation. However, here again, strong differences exist among burning procedures, in which the samples are either heated to a moderate temperature (ca. 120–150°C in kôdô) or placed directly on glowing coal, far above 500°C, as is the custom in the Middle East. To bring some order, an exhaustive overview of the odorant constituents isolated from agarwood so far should be useful to provide the basis for future analytical work, where these compounds can be traced in all forms of samples obtained from extracted, hydrodistilled, and burned agarwood. Surprisingly, none of the pioneering studies of the Bhattacharyya group devoted to the characterization of agarwood sesquiterpenoids mention any olfactory descriptions for the isolated compounds. However, the importance of agarwood as a perfuming material in the traditional Japanese culture may explain why the first studies mentioning clearly the odorant properties of agarwood constituents were conducted in Japan. The simultaneous communications of two independent research groups at the 9th International Congress on Essential Oils in Singapore in 1983 [75, 119] illustrate the two different approaches following studies on agarwood odorants. These examined either its sesquiterpenoids or the pyrolysis products of its heavier constituents (mostly chromones), for the origin of its specific odor. Indeed, as pointed out later by Ishihara et al. [169], it seems that the odor of agarwood smoke is due to a harmonious combination of volatile compounds naturally occurring in the wood and various pyrolysis products. The same is true for the essential oil, which contains significant amounts of some constituents like 104 and 105, produced when chromones are heated, and not obtained from cold solvent extraction. Therefore, the following subsections will consider both types of constituents, and an exhaustive list of agarwood odorants with their olfactory descriptors is presented in Figs. 11 and 12. Of course, many other agarwood constituents are also odorants, and the description of their smell can be found elsewhere in the literature. The overview proposed in Figs. 11 and 12 is intended to focus only on those constituents described in studies on agarwood to help discern the specific odorants of this material.

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79

O

O

OH OH 10 intensive odor [76]

16

15

14 strong woody and slightly camphoraceous note [75]

woody and amber-like characteristic Jinkoh odor [75]

intensive odor [76]

CHO OH

O

COOH

OH

OH

18

O

woody, slightly camphoraceous, typical agarwood, when heated [170]

O

25

23

22

21

woody and slightly balsamic [75]

intensive odor [76]

pleasant, ß-damascenone-like woody note with a touch of camphor [81]

weak woody, harmonize other compounds on heating [84]

O O

OH 28

32

33

woody, slightly balsamic, bitter [84]

like 14, but rather sweet, woody [85], eucalyptol-like [171]

29

powerful, long lasting, woody, sweet note [82]

pepper [115]

OH

O

O

OH 34a

floral and woody note, with a nuance of smoky sandal [84]

34c floral, herbaceous, minty [137]

O

35a pennyroyal-like minty [84]

37

38

fresh, sweet, reminiscent of blooming flowers [81]

woody, sweet, rather weak [81]

O

O O

O

O

O 53

56

green, galbanum [89]

pleasantly woody, vetiver [90]

61 typically agarwood-like, sweet, woody, smoky, phenolic, like oak-moss, but weak [90]

104 sweet, floral, herbal, balsamic

105 very soft fruity floral [11]

Fig. 11 Agarwood odorants

4.1 Naturally Occurring Agarwood Odorants As mentioned above, a good overview of the global composition of the volatile part of agarwood can be provided by GC-based analysis of its extracts or essential oils. However, the characterization of the key odorants of these mixtures also requires rigorous sensory evaluations, because their olfactory properties are due only to a small part of their volatile constituents. The first mention of the odor of individual agarwood constituents was published by Nagashima et al., who isolated (+)-karanone

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OH

HO 146 extremely strong sweet [137]

O

HO

O

149

OH delicate sweet smell [96]

HO

O

HO

O

HO

OH 155

extremely strong spicy, slightly smoky [100]

OH O

157

HO

fir flavour, slightly refreshing cool, bitter, smoky [100]

OH O 159

158

floral, elegant [172]

cool minty, sweet note [172]

OH 154

powerful long-lasting pennyroyal-like minty, fumigating [171]

O

156

OH

153

powerful long-lasting pennyroyal-like minty, fumigating [171]

O

O

O

152

151

honey [96]

O

OH O

150

delicate sweet smell [96]

balsamic with a slightly cooling, camphor-like [171]

sweet woody [96]

OH

OH

HO

O 148

147

fresh floral, refreshing cool, slightly sweet note [172]

slightly spicy, cool minty, rather weak [172]

O

HO

HO

OH O

O

160

HO

long lasting refreshing cool, strong balsamic [172]

O

O

HO

HO 165

162 sweet, fruity, floral, anise, raspberry

163

164

leather [132]

smoky, phenolic, castoreum

O

SH

O 166

clove, spicy [132]

HO

HO

161 strong camphoraceous, sweet note [172]

clove, spicy [132]

SH

O

167

168

grilled meat, nut [132]

169

grilled, coffee [132]

O

fruity, sweet, plum [132]

HO CH2OOH

HO 170 smoky, phenolic, burnt [132]

171 frankincense [132]

172 leather, animalic, phenolic

Fig. 12 Agarwood odorants

(14) and (+)-dihydrokaranone (15) from the hexane-soluble portion of the acetone extract of a Cambodian agarwood: 15 was reported to give a “woody and amberlike characteristic jinkoh odor” and 14, a “strong woody and slightly camphoraceous note” [75]. These assertions were supported by the olfactory evaluations of both compounds, in the racemic and enantiopure forms, prepared, respectively, by total synthesis [173], and by semisynthesis from (+)-fukinone (131) isolated from

The Chemistry of Agarwood Odorants Scheme 4 Nagashima syntheses of (+)-14, (+)-15, (±)-14 (one enantiomer drawn), (±)-15 (one enantiomer drawn) [75]. Reagents and conditions: a) DDQ, dioxane; b) see Ref. [173]

81 O

O

a)

O

a)

(+)-14

131 (+)-fukinone

O

OH b)

(+)-14

15 O

a)

(+)-15

Petasites japonicus [174] (Scheme 4). Thus, (+)-14 was reported to have “more fumigating” and (+)-15 “more elegant” odors than their racemic counterparts. In the same study, they also isolated 18, described as possessing a “woody and slightly balsamic note”. Nevertheless, the authors concluded at the end of their communication that even if (+)-14 and (+)-15 “were the most important components which gave an oriental and elegant fumigating character to kanankoh”, they also recognized that they were not sufficient to reproduce the odor of this material, implying that some other constituents were certainly involved. They noted indeed that (+)-14 and (+)-15 possess odors “similar but stronger” than the other constituents 10 and 16 having “intensive odors”, without giving any greater detail on their individual olfactory properties [76]. The same group also isolated 10 and 16 in another agarwood sample from Indonesia [76], together with the prezizaanes 21 and 22, and mentioned laconically that these four constituents “appear to be a source of the fragrance of agarwood”. However, the respective epimeric structures 132 and 133, proposed in their original papers for 21 and 22 [76, 79], were actually erroneous (Fig. 13). These authors apparently overlooked that 132 had been isolated previously from vetiver oil by Bhattacharyya and named “allokhusiol” [177]. At the time, few prezizaanes were known: (+)-prezizaene 134 was identified also in vetiver [178] and its levo-antipode, ent-134, isolated along with its corresponding alcohol 135 by Ghisalberti from the Australian shrub Eremophila georgei [179]. In addition, the structures and absolute configurations of ent-134 and 135 were confirmed later by total synthesis from pulegone [180]. The wrong structure 132 proposed for jinkohol came mostly from the fact that the POCl3 /pyridine dehydration of this alcohol produced a dextrorotatory alkene, 136, a C-2 epimer of 134. The 1 H NMR and MS spectra of 136 obtained were closely comparable to those published for 134 and ent-134 [178, 179], hence the confusion. On the basis of the positive optical rotation, Nakanishi et al. assumed wrongly that 136 is identical to 134, and proposed the structure 132 for jinkohol after a careful examination of its 1 H NMR spectrum completed by NOE experiments to support the configuration of C-7. However, when Sakurai et al. performed the total synthesis of ent-132, ent-134, 135, as well as their epimers 137 and 138 [181], they noticed that the spectroscopic data of their synthetic ent132 and ent-134 were in good agreement with those published by Ghisalberti and Bhattacharyya, but not with those obtained by Nakanishi. The 13 C NMR chemical shift of 20.0 ppm of the C-12 methyl group of ent-134 was indeed significantly more

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N. Baldovini 14 13 4

6 12

3

11

1

2

OH

OH

OH

15

OH

7 8 9

10 12

21 ((–)-jinkohol)

132 ((+)-allokhusiol)

22 ((+)-jinkohol II)

133 (prezizan-15-ol)

OH

OH

ent-134 ((–)-prezizaene)

134 ((+)-prezizaene) OH

135 ((–)-7β -hydroxyprezizaene)

137 ((–)-epiprezizaene)

138 ((–)-epiprezizanol)

1) MCPBA 2) silica gel, n-hexane, reflux

POCl3/pyridine [79, 175]

OH

3) LiAlH4, Et2O [175] 21

22

136

O

OH NaBH4 [176]

139

133

1) MCPBA 2) silica gel, n-hexane, reflux

OH

ent-22

3) LiAlH4, Et2O [175]

OH

ent-134

ent-133

Fig. 13 Prezizaane and zizaane sesquiterpenoid compounds

deshielded than the C-12 of the dehydration product of jinkohol obtained from agarwood (14.3 ppm). Moreover, the spectroscopic data of 135 and 138 did not match either with those of jinkohol. Later, Weyerstahl eventually proposed the structure 21 for jinkohol, on the basis of the comparison of the 13 C NMR data of its diastereomers 132, 135, and 138 [176]. He noted the same discrepancy for the chemical shift of C-12 in jinkohol II, for which he attributed the corrected structure 22 instead of 133. Indeed, 133 could be prepared by NaBH4 reduction of aldehyde 139 isolated from vetiver oil and displayed a 13 C NMR spectrum different from that reported by Nakanishi for jinkohol II [176]. Ironically, Selvakumar and Rao had prepared (±)-133 and claimed to have synthesized jinkohol II, but did not notice any problem since they also based their confirmation only on 1 H NMR values [182]. The odor of 22 raised the interest of the fragrance industry since the Takasago Company in 1982 filed a patent on its use as a perfumery ingredient [170]. However, at that time they were unaware of the future controversies concerning the actual stereochemistry of 21 and 22, and they synthesized 22 starting from 21 extracted from

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83

agarwood, and ent-22 from ent-134 obtained either by the total synthesis reported by Coates [180] or from E. georgei essential oil [179]. If 22 can indeed be obtained from 21 via the dehydration product 136, in contrast, ent-134 must produce ent-133 and not ent-22. Surprisingly, virtually identical melting points and almost exactly opposite optical rotations were reported in the patent for the two epimers 22 and ent-133. According to the authors, 22 is one of the important agarwood odorants, and both 22 and ent-133 possess the “woody note characteristic of sesquiterpenes in combination with a somewhat camphorous odor” [170]. Considering the structure–odor relationships in the zizaane and prezizaane series, it seems that many of their members display interesting olfactory properties, whatever the variations in the nature, the position, and the stereochemistry of the oxygen function (Fig. 14) [176, 183, 184]. Research on agarwood chemistry conducted by Ishihara et al. was partially guided by interests in perfumery, and the olfactory properties of many of the compounds they isolated and synthesized were reported in their publications. Hence, the eudesmanes 32, 34a, 34b, 35a, 37, and 38, guaianes 23 and 28, and eremophilanes 14 and 33 were all characterized as important agarwood odorants [81, 84]. Among them, 23 was highlighted as “the most important component which contributes to the gorgeous OH

22

21 woody, slightly camphoraceous, typical agarwood, when heated [170]

OH

OH

camphorous, woody [170]

OH

O

ent-133

132

camphorous, woody [170]

strong, woody, ambra-like, camphoraceous, bitter, patchouli-like [176]

warm, woody, vetiver-like, slightly ambra-like [176]

O

O O

khusimone

slightly fruity, grape- fruit-like [176] vetiver, green, woodyambery th. 4.7 ng/dm3 air [183]

O

AcO

AcO vetiver, tenacious [184]

vetiver [184]

O

woody-ambery, cedar th. 0.13 ng/dm3 air [183]

O

HO

woody, vetiver, rhubarb th. 1.3 ng/dm3 air [183]

O vetiver [184]

vetiver, fruity [176] clean, fresh, transparent, woody-ambery, vetiver th. 0.13 ng/dm3 air [183]

aqueous, vetiver [184]

vetiver, cedarwood [185] woodyambery, typical vetiver [183]

vetiver, fruity, sour [176] strong vetiver, woody-ambery, transparent th. 29 pg/dm3 air [184]

HO

weak vetiver [184]

O strong, woody, grapefruit-like, typical note of vetiver [176]

Fig. 14 Structure–odor relationships in prezizaane and zizaane series with olfactory thresholds

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N. Baldovini O

R

HO

a)

b)

d)

74% 141

140 (-bulnesene)

142 e) f)

23 R = CHO 24 R = CH2OH 25 R = COOH

OH

HO

c)

HO

O

d)

+

92% 144

143

HO

R

THPO HO b)

h) i)

O

THPO

145

g)

O

c)

R = CHO 30 R = COOH R = COOCH3 d)

O

O

O

O O e) j)

k)

30

28

Scheme 5 Ishihara syntheses of guaiane agarwood odorants [81–83]. Reagents and conditions: a) MCPBA; b) Lithium dicyclohexylamide, cyclohexylmagnesium bromide, DME; c) LDA, EtMgBr, toluene; d) PCC, CH2 Cl2 ; e) NaBH4 , CeCl3 , MeOH; f) 10% aq. KOH, AgNO3 , EtOH; g) DHP, PPTS; h) NaClO2 , 2-methylbutene, aq. t-BuOH; i) CH2 N2 ; j) 5% HCl; k) stand in CDCl3 for two weeks

and elegant character of Kanankoh” [130]. Its olfactory contribution was confirmed unambiguously by semisynthesis from α-bulnesene (140) isolated from guaiac wood oil [81] (Scheme 5). A comprehensive exploration of the behavior of its epoxide 141 toward various strong bases helped to design two alternative routes leading selectively to either 142 or 143 [82]. Oxidation of 142 with PCC led to 23, which could be transformed further either to its corresponding alcohol or acid (24, 25). Alternatively, 142 was also obtained by photooxidation of a mixture of patchouli sesquiterpenes (mainly 140 and α-guaiene). This second approach was less efficient but provided an isomer of 142 that could be used for the semisynthesis of rotundone (29) [83]. Surprisingly, even if 29 is a well-known potent odorant [115, 130], the authors did not describe its contribution to the smell of agarwood. Compound 143 was also a useful intermediate since its PCC oxidation furnished the epoxyalcohols 144 and 145. This mixture was used directly to eventually provide the agarwood odorant 28.

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85

The same group also achieved the semisynthesis of both enantiomers of 14 and confirmed that, as mentioned previously by Nagashima [73], the natural (+)enantiomer is a “very important contributor which gives an oriental and fumigating character to agarwood”. In contrast, its antipode ent-14 has a “woody and slightly citrus note with a nuance of (+)-nootkatone” [84]. Neopetasane (33), an isomer of 14, was also isolated and showed the same olfactory properties as 14. As expected, 33 is thermally unstable, and easily isomerizes to 14 when heated. Nevertheless, some authors have claimed recently its identification in significant amounts in agarwood essential oil of various different origins [114, 185]. The contributions of Näf et al. to the characterization of agarwood volatiles brought a few additional odorants among the newly identified constituents: the eremophilane (56) (vetiver-like), vetispirane (61) (typically agarwood-like, sweet, woody, smoky, phenolic, like oak-moss, but weak) [90] as well as an unexpected linear ketone 53 with a green, galbanum-like odor [89]. It should be mentioned here that a typographical error in the review by Näf [2] led to some confusion in following the literature concerning the actual olfactory properties of 59 and 61. Since 2013, the group of H. F. Dai in Hainan, China has made a major contribution to the phytochemistry of agarwood and has reported a series of odorant constituents isolated from various agarwood samples of A. sinensis. Hence, the cadinane 146, eudesmanes 147, 148, and 149–151, eremophilanes 152 and 153, acoranes 154 and 155, and guaianes 156–161 were all isolated and reported to possess odorant properties [96, 100, 137, 171]. Most of these constituents are di- or trioxygenated, and the volatility of some is probably limited, but it may be interesting to explore their olfactory participation in the long-lasting base note of agarwood. Even if a constituent isolated from a mixture is an odorant (which is indeed the case of almost all organic compounds whenever they are sufficiently volatile), it does not guarantee that it will participate in the global odor of the mixture. Indeed, it may be present at amounts lower than its detection threshold in the headspace above the sample evaluated. However, synergistic effects may complicate this situation and here again, only recombination experiments (including omission tests) may bring a definitive answer [167]. Apart from the description of the odor of isolated constituents, the use of gas chromatography–olfactometry (GC–O) is extremely useful to afford more global information about the key odorants of mixtures. Despite the high value of agarwood, only two GC–O investigations of this material have been reported in the literature so far. In 2011, Pripdeevech et al. [186] published the results of GC–O analyses on three Aquilaria species, based on SPME sampling of their essential oils. The odors perceived in these GC–O experiments were attributed to their parent odorant compounds solely on the basis of their retention index, without confirmation by MS or the co-injection of reference samples. Moreover, for all agarwood essential oil samples examined, the major constituent identified by GC–MS was the pollutant 100. Therefore, this work will not be discussed further here. More recently, Tissandié [132] conducted a GC–O/MS study of a commercial agarwood essential oil, completed by GC × GC–MS analysis. The identifications were helped by fractionation of the sample by chromatography on silica gel and preparative

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GC followed by NMR analysis (see Chart C13, Fig. 7). The GC–O experiments involved three panelists and led to the detection of 53 odor zones, with the attribution of 13 of them to their corresponding odorant constituent. The characterization of the most potent odorants was proposed on the basis of aroma extract dilution analysis (AEDA) [187, 188] and revealed significant differences in sensitivity between the panelists. The odorants with the highest flavor dilution factor were 14, 104, phydroxybenzylacetone (162), 4-propylphenol (163), and some unknown compounds. Additional odorants of lower flavor dilution value were 15, 4-ethylphenol (164), eugenol (165), (E)-isoeugenol (166), 2-methylfuran-3-thiol (167), 2-furfurylthiol (168), β-damascenone (169), guaiacol (170), and olibanic acid (171). The characteristic animalic/leather odor of many agarwood essential oils is then probably due to compounds like 170, 163, and 164. 3-Propylphenol (172), an isomer of 163, was patented as a perfumery ingredient that boosts the animal aspect of artificial oud compositions, thus “approaching the reconstitution fragrance to the natural resin” [189]. Finally, the only sesquiterpenoids clearly identified as odorants in this study were 14 and 15, and contrary to previous reports [84, 90], 32 and 61 were reported here as odorless. The results of this work illustrate how difficult the determination of the key odorants of a complex material like agarwood essential oil can be, even using such a potent combination of up-to-date analytical tools.

4.2 Thermal Generation of Volatiles from Agarwood When used as an incense, a precious property of agarwood is its longevity: even very old samples can still be burned for perfuming purposes, in a similar manner to frankincense, another important natural incense [12]. Hence, these materials can be stored and transported over long distances, as a high-value commodity [1]. The reason for the durability of agarwood fragrance is probably linked to the fact that volatile compounds remain embedded inside the stem, and also that a significant part of the odor is liberated on heating, because of the thermal degradation of some oleoresin constituents. This heat sensitivity is indeed one of the properties of the chromones, which are then potential precursors of the smell of burned agarwood.

4.2.1

Chromone Pyrolysis

The pioneering (1935–1939) chemical investigations on agarwood led by Ichikawa [190, 191] gave an insight that some heat-sensitive constituents are responsible for the thermal generation of agarwood odorants. Indeed, to make up for the limited technical means of that time, olfactory evaluation of their fractions guided their chemical analyses. These early experiments on ethanol extracts of agarwood involved several steps of partition between different solvent mixtures, followed by simple sensory tests to help tracking the fractions that become odorant on heating. When one such fraction in a solution of potassium hydroxide was boiled and a fractional distillation

The Chemistry of Agarwood Odorants

87 O O

O

O O

O

O

O

O

173

OH 175

174

176

O 178

177

O O OH 179

OH

HO

HO

O

O

O

HO

O

181

180

O

183

182

184 OH

O

O O

OH 185

O 187

186

O

O O

O

HO

O

HO

189

188

O

O

OH

191

190 COOH

O

HO O

HO

194

193

192

195

196

197

O

HO 198

199

200

201

O

O

O

202

203

204

205

O O

HO O 206

HO

O 207

208

Fig. 15 Pyrolysis products of chromones and agarwood

performed on the result, it was possible to isolate and characterize 104. To do so, simple methods available at that time were utilized, including elemental analyses, measurements of boiling point and of the melting point of the semicarbazide, and, last but not least, evaluation of the odor. They also identified 105 in the distillation cuts as “possessing an odor of heated agarwood”. The characterization of the precursor of these compounds was obviously out of reach at that time, but when Yoshii et al. identified 120a 40 years later, they suggested that it was the “benzylacetone producing substance” evoked by Ichikawa, as 120a that produced 104 when heated in 5% methanolic soda. Shortly after the discovery of 120a, Yamamoto et al. identified two other chromones, 121a and 121b, in the steam distillation residue of an acetone extract of Vietnamese agarwood [119]. Both of these compounds were sufficiently volatile to be detected by GC and were synthesized to confirm their identifications. They also hypothesized that 121a and 121b were thermally unstable in the same way as 120a. Both synthetic and natural samples of 121a and 121b were pyrolyzed for 6 h at 150°C under an air flow and the emitted volatiles were trapped on tenax,

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to identify benzaldehyde (173) and p-anisaldehyde (174), as well as traces of 104 and 4-methoxyacetophenone (175) (Fig. 15) [151]. The participation of these simple compounds in the typical odor of agarwood was not clearly stated, but 173 and 174 are well-known odorants, although their individual odor is not particularly reminiscent of agarwood. Recently, Takamatsu and Ito studied the decomposition of pure 120a on heating (190–200°C) by SPME/GC–MS. They could thus confirm the formation of 104 and 173 and identified also methyl phenylpropionate (176) [168]. In addition, they showed that 120a is fairly hydrophilic and can be detected by HPLC in significant amounts in agarwood decoctions [168].

4.2.2

Smoke Analysis

In parallel with the studies on the pyrolysis of pure agarwood constituents, a number of authors have investigated the composition of the smoke produced when agarwood is burned. In 1984, Nagashima reported the identification of several small aromatic substances by GC–MS analysis of the agarwood smoke emitted in the traditional conditions of the kôdô. Several genuine agarwood (nor)-sesquiterpenoids (4, 9, 10) were detected, in addition to 104, 105, 173, 174, and other compounds rather common for pyrolysis products of wood material: guaiacol (170), acetic acid (177), furfural (178, furfuryl alcohol (179), phenol (180), vanillin (181), 4-ethylguaiacol (182), dihydrocoumarin (183), and phenylpropionic acid (184) [192]. Later, Ishihara et al. gave a more detailed account of a similar experiment in which they trapped the smoke of Kanankoh and Jinkô (180–200°C) on tenax and analyzed the result by GC–MS after solvent desorption. The samples were similar to those studied in his work on solvent extracts [130] discussed previously (see Sect. 3.1.2) and represented in charts C6 and C7 of Fig. 7. For the same sample, the comparison of the GC profiles of the agarwood extract and of the smoke extract displayed a surprising similarity for the sesquiterpenoid part, which did not show any sign of degradation. The two chromones 121a and 121b were also detected, albeit in lower amounts than in the solvent extract. The smoke extract contained the compounds reported in other experiments on smoke: 104, 105, 170, 173, 174, 177–181, and additional ones: 185–195. Ishihara concluded that the odor of the smoke was due to a harmonious combination of the smell of the odorant sesquiterpenoids and the aromatic compounds resulting from the pyrolysis [169]. A detailed analysis of agarwood smoke components was performed using Curiepoint pyrolysis-GC/MS [193]. This technique helped to explore the behavior of agarwood heated at higher temperatures (up to 500°C), a good model of the harsher conditions used in the Middle East when it is placed directly on glowing charcoal. Many of the aromatic constituents mentioned above were identified as well as 196– 208. In addition, many C15 compounds suspected to be sesquiterpenes also appeared on the chromatograms. Interestingly, when pure 120a was treated under the same conditions, it yielded a surprisingly high number of constituents: 104, 105, 184, 186– 188, and 198–204. Therefore, if so many substances are produced by the pyrolysis of a single compound, the whole set of chromones present in agarwood should deliver

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a significant number of volatile components on burning. Here again, these highly degraded products certainly participate in the smell of agarwood smoke, as some of them are well-known odorants. Impressive work was conducted by the Takasago Company on the odorants of agarwood smoke and was presented in 2002 by Yaguchi at the 46th TEAC Congress in Tokushima (Japan) [194]. They reported on the analyses of the “Ichiboku shimei”, a set of four titled pieces of an agarwood sample of the highest historical value, presumed to have been imported to Japan in 1624. To cope with the inestimable value of these samples, only a very limited amount (ca. 1 mg) of each was collected and submitted to thermodesorption-GC–MS/O. Despite the age and the scarcity of the samples, the analyses indicated the presence of several sesquiterpenoids (14 and others), a chromone (121a), and aromatic compounds (181, 184, 197). In addition, the GC–O olfactograms revealed many sweet, animalic, amber-like, musky, and agarwood-like odor zones reminiscent of high-value kyara. Lastly, the elegant account of R. Kaiser on his kôdô style experiment should be mentioned, where he trapped the smoke of a 50 mg kyara sample to analyze it by GC–MS [11]. His conclusion was that several compounds are important contributors to the smoke odor, among which 14 and 15 (oriental woody), 105 (soft fruity floral), “raspberry ketone” 162 and 29 (an often neglected potent peppery odorant), 174 (sweet, powdery), vanillin (181), and 192, which bring floral, spicy, and vanilla notes, while 4-ethylphenol (164) imparts animal-like, castoreum, and leathery tonalities. Taken alone, none of these substances is totally characteristic of agarwood odor, but the subtle smell of this noble material is a balanced association of all of their odiferous notes.

5 Conclusion The use of agarwood as an incense emerged more than three millennia ago in the Indo-Gangetic plains and then spread throughout almost all parts of Asia where it has become a most precious perfumery material. Occasionally, agarwood also reached Europe via the Silk Road but never received much attention from Western perfumers despite the crucial role of agarwood and agarwood essential oil in the oriental olfactory culture. It was only in 2002 when agarwood essential oil was introduced in a French perfume for the first time, that the “oud fashion” boomed in the Western perfumery industry. In parallel, the overexploitation of wild agarwood to fulfill the growing Asian market has created a strong depletion of the natural resources and has stimulated the development of the cultivation of agarwood-producing trees and artificial induction techniques. Many research groups all over the world have worked to explore its chemical composition, its medicinal properties, and the complex mechanisms by which agarwood resin is generated in parasitized trees. This burgeoning scientific interest for agarwood observed in the last few decades is not only due to mere curiosity for a seemingly vanishing plant. It is also motivated with the aim of participating in the rebirth of this fascinating material since hopefully, its

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sustainable production is now realistic. Thanks to the creative involvement of forestry researchers, agronomists, microbiologists, and chemists, many advances have been made in the exploitation and production of agarwood. However, there is still a long way to go to completely unravel its complex properties and fully benefit from its rich potential in pharmacology and perfumery. Many challenges still await researchers of all specialties participating in the study of agarwood. The establishment of ecological and sustainable production processes is required and should be conducted through fair and ethical business models involving local stakeholders. The techniques for the botanical characterization of agarwood-producing species still need to be improved. Ideally, they should be made possible directly on agarwood samples. Several interesting advances have been made in the detection of agarwood adulteration and authenticity using various techniques. Conventional chemical analyses (LC and GC–MS) can also still be extremely helpful to characterize agarwood and agarwood essential oil samples, with the proviso that they follow recommended analytical practices. Finally, the question of the identity of the key odorants of agarwood is still unresolved, because the chemical complexity and the variability of this material together prevent any “simple” answer. An extensive survey on commercial agarwood essential oil samples of different origins reveals an impressive diversity in their olfactory character, even if their chemical composition is often roughly similar [110]. This situation is possibly due to the participation of poorly characterized trace constituents in the subtle harmony of the agarwood odor. In addition, many odorant components described in early studies are not detected in conventional agarwood essential oils but occasionally reappear in some atypical samples. A consideration of the odor of burned agarwood adds another layer of complexity to this question, as a similar variability exists also in the smell of the smoke obtained from different agarwood samples. Many professional perfumers seduced by the “oud trend” often eventually turn away from this material because of its high price and its inconsistent quality. Better characterization of agarwood odorants should enable defining the best markers of quality, eventually to help develop a sustainable agarwood industry able to provide perfumers with affordable good quality samples, to finally restore the “wood of the gods” to its former glory. Acknowledgments It is my pleasure to thank many people with whom I had the occasion to talk during the writing of this review. In particular, I am extremely grateful to Dr. Daniel Joulain (SCBZ Consulting, Grasse, France) for sharing with me his deep knowledge and his bibliography, for many discussions on various aspects of fragrance chemistry, and for the precious samples of agarwood oil he kindly offered me. I would like to thank also Elise Carenini (Albert Vieille-Givaudan, Vallauris) for a 2020 Christmas gift of many samples of agarwood oils. Kazutoshi Sakurai and Brian Lawrence are also acknowledged for the exchange of bibliographical information. I wish to express my thanks to Alan Mahaffey (Agarwood Consulting, Grasse) for providing me with many agarwood oil and wood samples, and much useful information about agarwood. Finally, Juliette Pattinson is warmly thanked for proofreading this manuscript.

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N. Baldovini Nicolas Baldovini studied organic chemistry at the University of Aix-Marseille. He obtained a Ph.D. degree in analytical and organic chemistry from the University of Corsica in 2000, working on the chemistry of natural volatile compounds. After two postdoctoral stays in the groups of Prof. Solladié at the University Louis Pasteur in Strasbourg (2000–2001) and at the University of Tokyo with Prof. Koichi Narasaka (2002–2003), he was appointed Assistant Professor in 2003 at the University of Nice-Sophia Antipolis. His current research interests are focused on fragrance chemistry, analytical and synthetic chemistry of odorant compounds, analysis of natural raw materials, and structure-odor relationships.

Chemical Ecology of the North American Newt Genera Taricha and Notophthalmus Charles T. Hanifin, Yuta Kudo, and Mari Yotsu-Yamashita

Contents 1 2 3 4 5 6 7

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Taxonomy, Systematics, and Distribution of North American Newts . . . . . . . . . . . . . . . . Tetrodotoxin Structure and Stereoisomers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pharmacology of Tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phylogeny and Evolutionary History of Tetrodotoxin in Taricha and Notophthalmus . . . . . Origin of Tetrodotoxin: Biosynthesis or Sequestration? . . . . . . . . . . . . . . . . . . . . . . . . . . . Tetrodotoxin: Levels and Variation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Geographic and Species Level Variation in Tetrodotoxin Isomers . . . . . . . . . . . . 8 Tetrodotoxin as a Chemical Defense: Protection Across Multiple Life History Stages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 General Protection Against Vertebrate Predators in Adult Newts . . . . . . . . . . . . . 8.2 Coevolution Between Thamnophis Snakes and Newts . . . . . . . . . . . . . . . . . . . . . 8.3 Predation and Tetrodotoxin Sequestration in Newt Eggs . . . . . . . . . . . . . . . . . . . . 8.4 Protection from Parasites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Broader Ecological Impacts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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C. T. Hanifin (B) Department of Biology, Utah State University, 320 N. Aggie Blvd, Vernal, UT 84078, USA e-mail: [email protected] Y. Kudo Graduate School of Agricultural Science, Tohoku University, 468-1 Aramaki-Aza-Aoba, Aoba-ku, Sendai, Miyagi 980-8572, Japan e-mail: [email protected] M. Yotsu-Yamashita Graduate School of Agricultural Science & Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, 468-1 Aramaki-Aza-Aoba, Aoba-ku, Sendai, Miyagi 980-8572, Japan e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2022 A. D. Kinghorn, H. Falk, S. Gibbons, Y. Asakawa, J.-K. Liu, V. M. Dirsch (eds.), Progress in the Chemistry of Organic Natural Products 118, https://doi.org/10.1007/978-3-030-92030-2_3

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1 Introduction Chemical defenses play critical roles in the ecology of amphibians [1]. This group is known for the toxins and secondary metabolites that serve to protect them across a range of antagonistic interactions [2–8]. Many amphibians rely on a combination of aposematic coloration coupled with chemical defenses (as well as crypsis) to ward off potential predators across multiple life history stages (e.g., eggs and larval stage as well as adulthood) [2, 9–12], but chemical defenses also protect amphibians from parasites, fungal infections, and microbial pathogens [6, 13–15]. Well-studied examples of chemical defenses in amphibians include the alkaloids present in poison dart frogs [7, 8, 16], toads (e.g., Bufo; [3, 16, 17]), as well as numerous species of salamanders (Caudata) including the clade that contains the North American newt genera Taricha and Notophthalmus: the Salamandridae [5, 10]. The term chemical defense incorporates multiple strategies and compounds [18]. In some cases, amphibians produce sticky or slimy secretions that have little pharmacological effect but interfere physically with predation or attack [2, 19]. In other cases, amphibians produce noxious or emetic compounds that deter predators from attack [2, 20, 21]. These types of secretions are frequently coupled with aposematic (warning) coloration and/or distinctive antipredator displays to further enhance the efficacy of the antipredator strategy [2, 9, 12, 22–25]. Last, some amphibians (e.g., poison dart frogs and newts) possess extremely toxic compounds that cause death or severe physiological effects in predators [8, 16, 26–28]. These toxins frequently are neurotoxins that can have rapid physiological effects and are often coupled with striking aposematic coloration to deter predators [10, 28–33]. Chemical defenses are common in the Salamandridae [2, 5, 10, 18, 34, 35]. These salamanders are known for the variety of highly toxic alkaloids and/or pseudoalkaloids that have typically significant physiological effects [5, 10, 16, 33, 34, 36–39]. Fire salamanders (genus Salamandra) secrete and spray a mix of protective alkaloids including samandarine and samandarone that can cause pain and blindness [10, 16, 18, 34, 36, 39–42]. European primitive newts such as Pleurodeles watl possess toxic skin secretions as well as a mechanism to inject these secretions during predation events [43, 44], and Asian species of the same clade (e.g., Tylototriton) are also chemically defended [2, 45–48]. The defensive chemical ecology of Taricha and Notophthalmus as well as other closely related genera in the modern newt clade has been investigated and studied (reviewed in [5]; see also [35, 49]). This group uses tetrodotoxin (1), a low-molecular-weight toxin, as a defense across multiple life history stages [10, 26–29, 37, 50–54]. Tetrodotoxin (1) has been identified in all genera of Asian and European modern newts and appears to be a shared derived character (synapomorphy) that is one of the defining characteristics of the group [5, 27, 33, 35, 55, 56]. Examinations of genes that underlie auto-resistance to 1 in this clade have shown that tetrodotoxin-resistance is an ancient character that was likely present in the ancestor of all modern newts suggesting that the tetrodotoxin-bearing phenotype is ancient and that 1 has likely shaped the evolution and defensive ecology of the group for the last 50 million years [55, 57].

Chemical Ecology of the North American …

103 OH

HO H2N

O

OH N H

O OH

HN

OH

1 (tetrodotoxin)

Technical advances coupled with broad, integrative approaches over the last three decades have led to significant advances in our understanding of the chemistry and ecological impact of 1 in newts. These advances include an expanded picture of the defensive role of 1, its critical role in mediating landscape levels of coevolution with tetrodotoxin-resistant snake predators, the role of 1 at multiple trophic levels (e.g., defense against parasitism) and developmental levels (e.g., defense of newt eggs and larva), its potential role in the defensive chemical ecology of other species (e.g., mimicry and toxin sequestration by snakes), its role as a signaling molecule in species interactions, as well as a more complete understanding of the evolutionary history of the tetrodotoxin-bearing phenotype 1 in newts. The emerging picture is one that requires a broad understanding of the species interactions that are mediated by 1, including defense against parasites (e.g., [13], potential coevolutionary interactions and indirect selection on adult levels of 1 from egg predation [58], landscape level patterns of coevolution with snakes [59–61], its role in conspecific and trophic signaling [54], as well as the possibility that tetrodotoxin-toxicity is, in part, inducible rather than fixed in some populations [51]. Although the origin and biosynthesis of 1 in newts is still unclear, recent work has shed new light on the possibility that 1 is produced endogenously by newts, suggesting those that are tetrodotoxin-bearing can respond to selection through this web of species interactions [5, 62].

2 Taxonomy, Systematics, and Distribution of North American Newts North American newts comprise two genera: Taricha and Notophthalmus. There are four currently recognized species in Taricha: the roughed-skin newt (T. granulosa), the California newt (T. torosa), the red-bellied newt (T. rivularis), and the Sierra newt (T. sierrae) [63–67] (Plate 1). Three species with five subspecies are recognized

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Plate 1 Taricha. a The rough-skinned newt (T. granulosa), b the California newt (T. torosa), c the red-bellied newt (T. rivularis), and d the Sierra newt (T. sierrae). Photographs courtesy of E.D. Brodie Jr

currently in Notophthalmus: the black-spotted newt (N. meridionalis), the Texas black-spotted newt (N. meridionalis meridionalis), the striped newt (N. perstriatus), the Eastern newt (N. viridescens), the broken-striped newt (N. viridescens dorsalis), the central newt (N. viridescens louisianensis), the peninsula newt (N. viridescens piaropicola), and the red-spotted newt (N. viridescens viridescens) (Plate 2) (see [67]). The genus Taricha is distributed along the west of North America with a northern range limit that includes southwestern Alaska and northern British Columbia (Taricha granulosa) and a southern range that includes historical reports from San Diego County (California, USA) (Taricha torosa). Taricha rivularis is limited to a small region of coastal California north of the San Francisco Bay (Mendocino CO) and Taricha sierrae is limited to the foothills of the Sierra Mountains in central California [68]. Notophthalmus is distributed widely in Eastern North America with a southern range that includes coastal populations in Florida, Alabama, Mississippi, Louisiana, and Texas [67, 69]. The western edge of the range of the genus extends to the Midwest region of the USA as well as to populations in Ontario (Canada). These populations north of Lake Superior along with populations in New Brunswick and central Quebec (Canada) represent the northern range limit for Notophthalmus [67, 69].

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Plate 2 Notophthalmus. a The Eastern newt (N. viridescens viridescens), b the peninsula newt (N. viridescens piaropicola), c the broken-stripe newt (N. viridescens dorsalis), d the black-spotted (Texas) newt (N. meridionalis), e the striped newt (N. perstriatus). Photographs courtesy of E.D. Brodie Jr

3 Tetrodotoxin Structure and Stereoisomers Tetrodotoxin (1) is a complex and structurally striking molecule. It consists of a guanidinium moiety bound to a 2,4-dioxaadamantane skeleton (Fig. 1; see [49, 70] for recent reviews). The toxin typically is described as an alkaloid because the nitrogencontaining ring in the guanidinium moiety has been thought to be derived from an amino-acid precursor, but recent work has hypothesized that this core moiety of 1 present in newts (as opposed to 1 in marine species; [71, 72]) may be derived from a terpene rather than an amino acid [70, 73, 74]. Regardless, the structure and mode of action of 1 is highly unusual with structural and functional properties that are shared only with saxitoxin (15) and saxitoxin analogs [10, 75, 76].

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H2 N

H

R2

O 4

3

R3 O

N

2 1

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R7

H2N

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7

8

O

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4a

8a

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OH O

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R6

R4

R2 H OH H H H

R1 1 (tetrodotoxin) H 2 (4-epitetrodotoxin) H 3 (6-epitetrodotoxin) H 4 (8-epitetrodotoxin) H 5 (1-hydroxy-8OH epitetrodotoxin)

OH

R3 R4 OH OH H OH OH CH2OH OH OH OH OH

R5 CH2OH CH2OH OH CH2OH CH2OH

R6 H H H OH OH

R7 OH OH OH H H

6 (4,9-anhydro-tetrodotoxin)

O H2N

H2 N

H OH O

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9

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R1 R 2 H OH

7 (8-epi-5,11-dideoxytetrodotoxin) 8 (1-hydroxy-8-epi-5,11OH OH dideoxy-tetrodotoxin) 9 (8-epi-5,6,11-trideoxyH H tetrodotoxin) OH H 10 (1-hydroxy-8-epi5,6,11-trideoxy-tetrodotoxin)

R1

R1 R2 R3 R4 11 (4,9-anhydro-10-hemiketalOH CH2OH H OH 5-deoxy-tetrodotoxin) CH3 OH H 12 (4,9-anhydro-8-epi-10-hemiketal- H 5,6,11-trideoxy-tetrodotoxin)

HO 10

9

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11 8a

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H

8 7

OH 7

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13 (Cep-210)

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4

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2

O

9

H

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6

OH

14 (Tgr-288A)

Fig. 1 Chemical structures of tetrodotoxin (1), 4-epitetrodotoxin (2), and representative newt specific analogs of 1 and guanidino compounds including 6-epitetrodotoxin (3), 8-epitetrodotoxin (4) as well as possible precursors. Compounds 13 and 14 are newt-specific molecules that may represent a link between a terpene precursor and 1

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O O

H2N

H N

HN

NH2 N

NH

OH

OH 15 (saxitoxin)

Naturally occurring analogs, stereoisomers, and structurally related compounds are also present in organisms that possess 1 (Fig. 1; [10, 35, 49, 75]). Although an exhaustive survey of the chemistry and biosynthesis of 1 is beyond the scope of this chapter, variations in the naturally occurring analogs are relevant to the origin and possible biosynthesis of 1 present in Taricha and Notophthalmus (see Sect. 6) and may also be important in chemically mediated interactions of Taricha (see Sect. 7.1). Analysis of tissue extracts (primarily skin, egg, or whole animal) using highperformance liquid chromatography coupled with either fluorescence detection or liquid chromatography-mass spectrometry has demonstrated that 1 is the primary isomer present in both Notophthalmus and Taricha, but a core group of stereoisomers and/or analogs (Fig. 1) typically are also present [35, 75, 77–84]. These isomers of 1 include stereoisomers that are thought to be alternative or chemical equilibrium products of tetrodotoxin synthesis (e.g., 4-epitetrodotoxin (2) and 4,9-anhydrotetrodotoxin (6)), degradation products of 1, and other isomers/analogs that possibly represent reaction shunt products. Many of these isomers (e.g., 2) as well as chemical equilibrium analogs (e.g., 6) appear to be ubiquitous in marine and terrestrial tetrodotoxin-bearing species. However, a subset of derivatives (3–5 and 7–10) of 1 seem to be unique to newts and have only been identified in Taricha and Notophthalmus (or other newts such as Cynops), but not in marine tetrodotoxin-bearing metazoans or tetrodotoxin-producing bacteria (Fig. 1). Chief among these are the C-6 and C-8 variants: 6-epitetrodotoxin (3) and 8-epitetrodotoxin (4) and related breakdown/shunt products (Fig. 1). These isomers of 1 and the 10-hemiketal type analogs 11 and 12 of 1 (Fig. 1) currently are thought to represent alternative forms of 1 that may be generated during its biosynthesis [70, 73, 74, 85–87]. In addition to these clear tetrodotoxin analogs, work in newts has identified multiple bicyclic and tricyclic guanidino compounds (e.g., Cep-210 (13) and Tgr-288A (14); Fig. 1) that appear to be modified monoterpene geranyl derivatives. Recent work has proposed that these represent newt-specific precursors of 1 and reaction shunt products associated with terrestrial tetrodotoxin biosynthesis (see Sect. 6; [70, 73, 74, 85–87]).

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4 Pharmacology of Tetrodotoxin Tetrodotoxin (1) binds selectively to and blocks the ion-conducting pore of voltagegated sodium ion channels [30, 31, 38, 88, 89]. These channels are responsible for the initiation and propagation of action potentials in nerve and muscle cells (with some exceptions e.g., nematodes and crustaceans). Action potential generation by voltagegated sodium ion channels results from the rapid influx of sodium ions through the channel pores [90, 91]. Tetrodotoxin (1) binds to the outer portion of each pore and occludes it, thereby preventing sodium ion influx [92]. The physiological impact of 1 is rapid and can be severe [93]. In animals that use negative pressure to ventilate their lungs such as reptiles and mammals, the flaccid paralysis generated by sodium channel block typically leads to death by suffocation in non-resistant animals [26]. Rapid lethal responses (less than 1 min) to the oral intake of 1 have been documented in multiple potential predators of Taricha and Notophthalmus [26]. The rapid effect of 1 is likely to be important in the context of newt-predator interactions for two reasons: (1) it appears to allow newts to survive predation events because rapid paralysis or death of the predator interrupts attacks, and (2) it provides a mechanism for predators to “sample” and avoid toxic newts prior to predation. Evidence for (1) is primarily anecdotal but there are examples of live newts found in the esophagus or mouth of snake and avian predators [26, 94] (personal communication: E.D. Brodie Jr.). Evidence for (2) comes from studies of Thamnophis sirtalis and highly toxic Taricha granulosa demonstrating that snakes sample and reject newts with high levels of 1 [95]. Whether rejection and avoidance results from a poorly understood signal-response processing by snakes or simply from rapid onset of paralysis in the oral cavity in predators, it likely represents an important element of the defensive properties of 1 for newts. The ability of a newt to survive predation because of 1 generates a simple Darwinian model in which the presence of this compound in the skin of an individual newt directly increases its ability to survive predation and reproduce.

5 Phylogeny and Evolutionary History of Tetrodotoxin in Taricha and Notophthalmus The two genera of North American newts, Taricha and Notophthalmus, are members of a monophyletic clade (modern newts; family Salamandridae) that includes 12 genera with a global distribution [5, 96, 97]. Within the modern newt clade, tetrodotoxin (1) is present in Asian, European, and New World taxa (reviewed in [5]; [27, 35, 75, 78, 83]. Although levels of 1 vary considerably across species and genera, with some populations possessing only low levels and others (e.g., some Taricha) possessing extraordinary levels (but see Sect. 7), the consensus is that 1 is present at levels that are biologically relevant (e.g., protection from predation or possibly endoparasites) throughout the clade.

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While the evolutionary history of 1 itself in newts cannot be resolved currently because the origin (and/or genes associated with biosynthesis) of 1 in newts is still unclear, the tetrodotoxin-bearing phenotype also incorporates auto-resistance to 1 [55, 57, 98]. As such, the evolutionary history of tetrodotoxin-resistance in modern newts provides an important insight into the evolutionary history of the tetrodotoxinbearing phenotype in the clade. Tetrodotoxin (1) binds to and blocks voltage-gated sodium ion channels (see Sect. 4), which are responsible for the initiation and propagation of electric signals in animal nerve and muscle tissue. As such, the evolutionary history of tetrodotoxin-resistance in these proteins (and associated genes) can provide insight into the evolutionary history of tetrodotoxin-resistance in newts. Studies of voltage-gated sodium ion channels in modern newts have shown that resistance to 1 likely predates the use of this compound as a chemical defense in newts [55]. All members of the Salamandridae including multiple genera (e.g., Salamandra and Pleurodeles) that do not appear to possess 1, have low-level or moderate organismal resistance to 1 and possess tetrodotoxin-resistant voltage-gated sodium ion channels [55]. Extended analysis of the voltage-gated sodium ion channel gene family in Taricha and other modern newts supports this pattern and suggests that tetrodotoxinresistance evolved once in the common ancestor of modern newts [55, 57, 98]. Taken together, the genetic history of tetrodotoxin-resistance in modern newts suggests that the tetrodotoxin-bearing phenotype arose in the common ancestor of all modern newts (including Taricha and Notophthalmus) and likely played an important role in their survival and dispersal across the last 50 Ma [55].

6 Origin of Tetrodotoxin: Biosynthesis or Sequestration? A fundamental question associated with tetrodotoxin (1) in newts (as with other tetrodotoxin-bearing metazoans such as puffer fish and octopuses) is the source of 1 that is present in the organs and tissues of these animals. This question bears directly on the evolutionary ecology of 1 in newts because it centers on the ability of newts to respond to selection imposed by predation or interactions with other species. Although sequestered (as opposed to endogenously produced), 1 could still function as a defensive toxin in newts, the evolutionary frameworks of synthesized versus sequestered 1 in newts are fundamentally different. Resolution of this question is critical for understanding the evolution of tetrodotoxin toxicity in newts as well as the evolutionary ecology of their chemical defenses. Tetrodotoxin (1) is distributed widely in metazoans and has been documented in well over 60 species representing eight phyla, including Chordata, Echinodermata, Mollusca, Arthropoda, Chaetognatha, Annelida, Platyhelminthes, and Nemertea (see [5, 99, 100] for reviews). Importantly, 1 and its production have also been identified in several species of bacteria (see [49, 98] for recent discussions). The presence of 1 in multiple distantly related groups coupled with its occurrence in bacteria have led many to hypothesize that the presence of 1 in Metazoans results from symbiosis with tetrodotoxin-producing microbes and sequestration of a microbially produced

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toxin. Evidence in support of this hypothesis comes largely from work in marine taxa (see [49, 71, 72] for updated discussions) and some recent work in Taricha is also congruent with a bacterial origin [98, 101]. There is, however, a growing body of work that suggests that a “one size fits all” hypothesis for the origin of 1 may not apply to all metazoans and that newts may be able to produce their own. Evidence includes studies demonstrating that tetrodotoxin-bearing newts do not appear to harbor tetrodotoxinproducing bacteria in their skin [102], that they are able to maintain high levels of 1 in captivity [103, 104], and that tetrodotoxin-bearing newts possess newt-specific isomers of 1 not present in either tetrodotoxin-bearing marine taxa or bacteria [5, 70, 73, 74, 84–87, 99]. Furthermore, newts have elaborate secretory skin glands that play a critical role in producing and secreting toxins in amphibians [105] and recent detailed studies of the glandular structure and the staining of 1 in the skin of newts demonstrates structural and chemical variations that correlate with differences in levels of 1 [62]. A critical advance associated with the possible biosynthesis of tetrodotoxin (1) in newts are recent discoveries of newt-specific isomers of 1 and analogs that may form a newt- (or terrestrial-) specific tetrodotoxin biosynthesis pathway (Fig. 1) [70, 73, 74, 84–87]. A full examination of these data is beyond the scope of this contribution, but these new compounds include tricyclic guanidino molecules that provide a link between putative terpene precursors and 1 itself, reaction shunt products that are predicted by the reaction pathway, and intermediate molecules and analogs of 1 that provide reaction steps that explain the presence of newt-specific stereoisomers of 1, such as 6-epitetrodotoxin (3) and 8-epitetrodotoxin (4) analogs (Fig. 1; [78, 86, 87]). This work also suggests that variations in putative precursor molecules may be correlated with differences in organismal levels of tetrodotoxin (1) [74, 85, 87]. These advances in the biochemistry of 1 provide important insights into longstanding issues associated with the marine-terrestrial biosynthesis of this compound, and suggest that differences in stereoisomer profiles of newts when compared to bacteria and/or marine species may result from the presence of an endogenous biosynthesis pathway of 1 that is different from that present in bacteria [72].

7 Tetrodotoxin: Levels and Variation An often-unrecognized element of the evolutionary ecology of tetrodotoxin (1) in newts is the fact that the average levels of 1 in most species and genera of modern newts including both Taricha and Notophthalmus are relatively low [5, 24, 25, 35, 59, 60, 80, 81, 83, 106–108]. Typically, individuals from populations of Old-World newts (e.g., Cynops or Triturus) possess nanogram levels of 1 and there are no examples of populations of either Cynops or Triturus that possess it at levels approaching those seen in extreme populations of Taricha [35, 81, 108]. These patterns hold in other genera of newts that possess 1 but are less-well studied. For example, populations of Triturus have levels of 1 in the nanogram or low microgram range, which are far less than the 15 to 20 mg amount that has been identified in populations of

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Taricha granulosa [52, 59, 60, 77, 83], and the same is true for Pachytriton and other Asian modern newts [35]. Within the North American newt genera (i.e., Taricha and Notophthalmus), most surveyed populations possess either low (picogram to nanogram) levels of 1 or are non-toxic (below measurable HPLC limits) [60, 80, 81, 109, 110]. This general pattern suggests that the ancestral state of levels of 1 in this clade is likely to be low. Given the remarkable potency of 1, low levels are still likely to be biologically relevant and would be able to defend these species (see Sects. 4 and 8). Patterns of variation within and among genera of newts suggest that tetrodotoxin (1) does not segregate at the level of either genus or species [60, 80]. Specifically, there is no evidence that any one genus or species within (and among) North American newts is more toxic because of innate species level differences in the production of 1 or its bio-sequestration. Instead, it appears that there is significant variation in levels across populations of all species and all genera; with moderate, low, and non-toxic populations present in all species and genera of North American newts [24, 60, 80, 107]. However, species in the genus Taricha appear to possess a larger proportion of moderate to high toxicity populations when compared to Notophthalmus [59, 60, 80]. This pattern possibly results from multiple factors, including empirical artifacts (e.g., less sampling of Notophthalmus populations) and others that result from differences in the biotic interactions between Taricha and Notophthalmus (e.g., coevolution with garter snakes as well as tetrodotoxin-resistant caddisflies; see Sect. 8).

7.1 Geographic and Species Level Variation in Tetrodotoxin Isomers Stereoisomers of tetrodotoxin (1) do not appear to segregate among species within the modern newts; HPLC analyses of Taricha, Notophthalmus, and other tetrodotoxinbearing newts (e.g., Cynops) have not identified species-specific isomers of 1 (reviewed in [5] and see [35, 75, 76, 78–83]). However, many of the isomers of 1 identified in Taricha and Cynops are present at very low levels and can be difficult to purify and characterize if samples are limited in quantity. As a result, many of these isomers have not been confirmed in Notophthalmus [80–82]. The newt-specific isomer 6-epitetrodotoxin (3) has been identified in Taricha and Notophthalmus as well as in Old World newts [75, 79, 81, 83]. However, as with levels of 1, there is significant variation in the amount and relative proportion of this isomer among populations [35, 75–83]. Although the majority of newt-specific analogs and/or intermediates of 1 are present only in trace quantities, 3 has been shown to be a major element of the profile of 1 for some populations of newts [60, 77]. This stereoisomer is present in all tetrodotoxin-bearing newts including Old World newts such as Cynops and Triturus as well as Taricha and Notophthalmus, but work in Taricha has shown that levels of 3 are highly variable among its populations. In some of these populations, 3 is present

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at only trace levels but in others it can represent 10 to 20% of the “total 1” present in the skin of individual newts [77]. Also of note is the fact that 1:3 ratios appear to possess little variation within a population; in populations where the amount of 3 produced relative to 1 is high, all individuals seem to produce higher amounts (CTH, unpublished data). Although the biological significance of this variation is still unclear, the sodium channel binding affinity of 3 is high enough so that amounts present in some populations of newts may be biologically relevant [111]. Large-scale studies of the geographic variation of 1 isomer profiles of Notophthalmus have not been completed, but LC–MS and LC-FLD studies of this genus have also documented some spatial variation of 1 profiles among populations of this genus [80, 81]. The ecological relevance of variation in isomer profiles is still unclear, but may suggest that selection is acting on profiles of 1 or that differences in profiles have some impact on interactions with other species.

8 Tetrodotoxin as a Chemical Defense: Protection Across Multiple Life History Stages The presence of a potent defensive toxin in modern newts had been identified long before the chemical identity of the presence of a “toxin” was established (see [37] and [5] for reviews). Hubbard (1903) described garter snake avoidance of Taricha, Phisalix [42] identified the presence of defense toxins in closely related genera of Old World newts (genus Triturus) in the early twentieth century, and Stuhr (1936) described the effect of Taricha skin toxin on the frog heart rate (from Brodie [26]). Extensive work by Twitty identified the presence of a paralytic toxin in Taricha embryos and eggs as well as broader investigations in the chemical ecology of Taricha [112–115]. These early studies were followed by work that clarified the chemistry, pharmacology, and identity of this skin toxin. In the 1960’s a series of investigations by a research group at Stanford University isolated and identified the toxin present in the eggs and adult Taricha torosa and elucidated its structure and pharmacology [28, 37, 116]. Although in early work the toxin was named tarichatoxin, it was quickly recognized that its molecular formula, structure, and pharmacology are identical to tetrodotoxin (1), which had been recently isolated, purified, and described from pufferfish (Tetraodontiodae) [117]. Further work demonstrated that 1 was present in multiple species of Taricha (e.g., T. rivularis) as well as Notophthalmus and related Old World genera of newts [26, 33, 50, 56], but appeared to be a synapomorphy, limited (in salamanders) to the modern newt clade [reviewed in 5, 55]. Brodie confirmed the presence of a tetrodotoxin-like toxin in the skin of adult Taricha granulosa as well as its physiological effect on a range of potential newt predators [26]. This important paper established that the skin toxin of newts (now known to be 1) generated rapid paralysis and death in mammalian, reptilian, and avian predators as well as in fish (Fig. 1). Furthermore, this work was the first evidence that some populations of garter snake predators (in the genus Thamnophis) were highly

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resistant to Taricha skin toxins and could consume toxic newts with only minor consequences. In addition, Brodie’s work with unpurified skin extracts from a population of newts in the central Willamette Valley (Soap Creek Ponds) demonstrated that some populations of Taricha were extraordinarily toxic (see Sect. 7). Work by Brodie and the Stanford group that had first purified and described tarichatoxin/tetrodotoxin demonstrated that 1 was also present in the populations of Notophthalmus from Eastern North America [33, 50]. These “Red Efts” also possess a striking color pattern thought to be aposematic and use the combination of 1 and the coloring as defense against predation [27, 50, 56, 118, 119]. There has been considerable work to investigate 1 and its impact on species interactions involving both Taricha and Notophthalmus since this early period. These investigations include extensive exploration of multiple coevolutionary arms-races between Taricha newts and garter snake predators [60, 61, 120–122], examination of 1-egg provisioning in Taricha and its relationship to tetrodotoxin-resistant invertebrate egg predators (caddisflies) [52, 123–125], as well as its possible role in conspecific predatory signaling in Taricha [51, 54, 126–128] (Fig. 1). Corresponding work in Notophthalmus has expanded our understanding of predator avoidance associated with tetrodotoxin [27, 53, 56, 129–131], and with 1 as a defense in early life history stages [53, 132, 133], the relationship between 1 and aposematic signaling [134–136], and the potential interaction between 1 and parasite load in this genus [13]. Additional work has also explored wider ecological consequences of the toxicity of 1 in Taricha and Notophthalmus and suggests that 1 may form the core of multiple Batesian mimicry complexes [134–138] as it may even function as a “keystone molecule” (see [54] for a discussion). The emerging picture from this body of work suggests that the ecological roles of 1 in newts are complex and intertwined and that a holistic model for understanding the chemical ecology of these species is required.

8.1 General Protection Against Vertebrate Predators in Adult Newts Tetrodotoxin (1) causes rapid paralysis and death in most metazoans (see Sect. 4) and there is ample evidence that the presence of this toxin protects adult and juvenile Taricha and Notophthalmus species from predation [24, 26, 27, 50, 53, 56, 90, 117, 120, 129, 130, 135, 136, 139]. Vertebrate predators including avian species (e.g., Great Blue Heron), Carnivora (e.g., Skunks and Bobcats), as well as bony fish (e.g., Oncorhynchus) are all susceptible to 1 as are most other Reptilia and Amphibia (Fig. 1) [26, 94, 140]. Furthermore, vertebrate predators also appear to avoid tetrodotoxin-bearing newts. Together, this work demonstrates that 1 present in the skin of Taricha and Notophthalmus species protect these animals from predation. Recent evidence includes a broad study of Notophthalmus [53], as well as

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specific interactions between snapping turtles and Notophthalmus [129]. One important aspect of this defense is that the rapid paralytic nature of 1 may interrupt predation events by vertebrate predators, rather than simply causing death when consumed (see Sect. 4). Interruption of predation, in turn, allows potential predators to learn to avoid newts as well as providing a direct selective advantage for increased levels of 1 in individual adult newts because of survival in response to attack.

8.2 Coevolution Between Thamnophis Snakes and Newts The most extensively studied aspects of the chemical ecology of North American newts are the coevolutionary interactions between Taricha and garter snake predators (genus Thamnophis) in western North America (Fig. 2). Ongoing work started in the late 1960s and then continued by multiple laboratory groups has generated a textbooktype example of a coevolutionary interaction between these two genera [141]. The core dynamic governing Taricha–Thamnophis interactions is described typically as an arms-race in which predation by snakes on co-occurring newts generates selection that favors increased tetrodotoxin levels in sympatric newts (Fig. 2). In turn, increased levels of 1 in newts generates selection that drives increased tetrodotoxin-resistance in snake predators (see Thompson [142], for a general discussion, and Refs. [59, 60, 120, 122, 143]). This combination is thought to generate a positive feedback loop of escalating reciprocal selection that produces extreme phenotypes associated with tetrodotoxin-resistance in snakes and tetrodotoxin-toxicity in newts [60, 122, 142, 143]. Early work supported this model and demonstrated three important elements: (1) newt-eating Thamnophis sirtalis that are sympatric with toxic newts possess elevated tetrodotoxin-resistance [143], (2) T. sirtalis that co-occurred with low toxicity newts on Vancouver Island have reduced resistance to 1 [120], and (3) comparisons of a small number of newt-snake population pairs showed a positive correlation between newt levels of 1 and snake resistance [122]. A larger analysis of levels of 1 in the genus Taricha coupled with range-wide patterns of snake resistance supported the arms-race hypothesis but suggested that this dynamic is limited to a subset of populations across the interaction [60]. Studies that extended analysis of tetrodotoxin-resistance in Thamnophis to other members of the genus have demonstrated that organismal resistance to 1 associated with convergent tetrodotoxinresistant substitutions in voltage-gated sodium ion channels has evolved in at least three additional species of garter snakes that are sympatric with tetrodotoxin-bearing Taricha. Additionally, geographic patterns of toxicity and resistance are consistent with arms-race coevolution in at least one of these [61, 144].

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Fig. 2 Selective pathways in a simplified “toxin” web for tetrodotoxin-bearing newts. Selection on levels of 1 in adult newts and early life stages are marked with black lines. Dashed lines represent potential or identified selection for increased tetrodotoxin-resistance in predators. Question marks indicate selective pathways that are predicted or may not apply throughout the range of tetrodotoxinbearing newts

Although the repeated evolution of tetrodotoxin-resistance in garter snakes provides strong (if indirect) evidence of the importance of 1 as a chemical defense in newts, the dynamics of these interactions and their impact on the overall defensive ecology of newts is still unclear. Significant unresolved issues include: (1)

The majority of Taricha populations (across all species and the range of the genus) are characterized typically by moderate to low levels of 1 (see Sect. 7), suggesting that runaway arms-races may be limited throughout the genus [59, 60, 120].

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(2)

Phenotype matching studies between newts and snakes have identified a striking asymmetry in which snakes always appear to be “ahead” of newts when phenotypes are mismatched. This includes super-resistant populations of snakes that co-occur with high toxicity newt populations as well as populations of moderately tetrodotoxin-resistant snakes that co-occur with low or nontoxic newts [59–61]. Analyses of meta-population genetic structure coupled with phenotype analyses (e.g., tetrodotoxin-resistance and tetrodotoxin-toxicity) indicate that coevolutionary arms-races are limited to a small number of populations distributed across the range of the interaction. Furthermore, these analyses suggest that selection on newt levels of 1 associated with snake predation is also limited to a small set of populations in the interaction [59]. Dietary studies of Thamnophis are limited but suggest that although Taricha is consumed by Thamnophis sirtalis [50, 145], in parts of the sympatric ranges Taricha may not make up a significant component of the diet of T. sirtalis throughout the range in which they co-occur [146].

(3)

(4)

There is currently no evidence that populations of Notophthalmus are involved with coevolutionary arms-races with Eastern species (or populations) of Thamnophis or any natricine snake predator. A single study has identified predation on toxic Notophthalmus by Hognose snakes (Heterodon platirhinos) that possess organismal resistance to 1 but examination of Hognose voltage-gated sodium ion channels failed to identify tetrodotoxin-resistant substitutions in those channels and their mechanism of tetrodotoxin-resistance is currently unknown [109]. Notably, levels of 1 in Notophthalmus populations typically are much lower than those of Taricha, which may also provide indirect evidence of the greater impact of tetrodotoxin-resistant garter snakes on the chemical ecology of Taricha.

8.3 Predation and Tetrodotoxin Sequestration in Newt Eggs As with many chemically defended animals, North American newts appear to provision their eggs and larva with 1 (Fig. 2, Refs. [26, 28, 52, 123–125, 147], but see [54]). Eggs and juveniles of both Taricha and Notophthalmus contain levels of 1 that are physiologically significant (e.g., up to 1 µg of 1 per egg; [33, 116, 148]). As with adults, there is evidence that eggs and juvenile newts are avoided by predators because of 1 [51, 116, 119, 132, 149]. For example, dragon fly larva (genus Anax) are unable to consume eggs and larva from populations of highly toxic Taricha [150]. Marion and Hay demonstrated that eggs of Notophthalmus are rejected by a range of aquatic predators [53]. However, Wrynn and Gall [133] showed that dragon fly larvae (genus Anax) are capable of consuming larva and metamorphs of Notophthalmus, suggesting that 1 may be present at very low levels in some populations or is not as universally effective as a defense in Notophthalmus. Furthermore, work in lowtoxicity populations of Taricha torosa has shown that, although eggs and juveniles

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possess 1, larval newts did not possess 1 (reviewed in [54]). Zimmer argued that larval Taricha are not chemically defended by 1, based on the observation of intraspecific oophagy and cannibalism as well as predation by crayfish (Fig. 2). However, newts are highly resistant to 1 [55] and muscle action potentials in crayfish are propagated through voltage-gated calcium channels rather than voltage-gated sodium ion channels. A defensive role for tetrodotoxin (1) in eggs and early life history stages is an important aspect of the overall chemical ecology of newts, because newts actively provision eggs with 1 [52, 124, 125, 150], and also individual egg levels of 1 in Taricha granulosa are positively correlated with maternal levels of this compound (Fig. 2; [52]. Furthermore, 1 is concentrated in ovarian tissue of female newts [33] and one of us (CTH) has measured patterns of 1 in adult female newts that suggest that the levels of 1 are higher in newts that are preparing to oviposit (CTH, unpublished data). Taken together, these elements of 1 in newts suggest that adult levels of this compound may respond to multiple levels of selection and that early life history stage predation may drive selection, favoring increased concentrations of 1 in adult female newts (Fig. 2). Selection through maternal effects (i.e., egg predation and provisioning) may play an important role in the overall evolutionary ecology of Taricha because there is now considerable evidence that eggs of Taricha are preyed upon by tetrodotoxin-resistant caddisfly larvae (genus: Limnephilus) [58, 123] (Fig. 2). Studies on Taricha–caddisfly interactions have demonstrated that caddisfly larvae consume newt eggs laden with 1 from populations in which an individual egg can contain upwards of 1 µg of 1 [52, 58]. Caddisflies are common throughout the range of Taricha in western North America. In addition, there is evidence that Taricha has been present in Oregon since the Oligocene (32–33 Ma) and caddisflies likely date to the late Cretaceous or early Paleocene (66 Ma), suggesting that potential ecological interactions between these taxa may be very old [151, 152]. The evolution of tetrodotoxin-resistance in caddisflies coupled with their ubiquity in aquatic environments used by newts for breeding are notable. When understood in the context of the age of potential predation of eggs by caddisflies, it is suggested that egg predation is likely to play an important role in the chemical ecology of Taricha that needs to be more fully explored.

8.4 Protection from Parasites An additional component of the chemical ecology of tetrodotoxin (1) and North American newts may also include protection from both ectoparasites and endoparasites (Fig. 2) [13, 153]. The general hypothesis is that parasites represent a significant threat to amphibians and there is evidence that the presence of skin toxins and other metabolites in amphibians protect these taxa from infection and infestation [13]. Although this hypothesis has been proposed for Taricha, evidence in support of this role for the genus is mixed [13, 153, 154]. Some studies have demonstrated negative correlations between levels of 1 and parasite loads in Taricha [13], but examination

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of populations of Notophthalmus do not support this role for 1 [153, 154]. An additional question that needs to be explicitly addressed is whether voltage-gated sodium ion channels are expressed in putative endoparasites of newts (e.g., nematodes). For example, extensive genomic analysis has shown that voltage-gated sodium ion channels are absent from the genomes of nematodes and that they do not generate action potentials [155]. Since 1 binds to voltage-gated sodium ion channels with high specificity and does not interact with other targets, it is unclear what the mode of action for 1 could be in these species and thus there is a lack of a clear physiological pathway to efficacy. As a result, this role for 1 is still in the early stages of investigation and requires considerable work to assess the role it plays in defending newts from parasites.

9 Broader Ecological Impacts The chemical defenses of Taricha and Notophthalmus also have significant impact on broader species communities. Many of these impacts are poorly understood and require further exploration, but may affect the structure of the communities in which these organisms exist. Particularly interesting examples are “convergent” Batesian mimicry complexes associated with chemically defended Taricha and Notophthalmus [134–136, 138]. Work in the 1970s and 1980s documented multiple examples of mimicry associated with the distinctive red coloration of Notophthalmus efts and exposure to 1 [135, 136] (Plate 3). This work demonstrated that naïve avian predators avoided nontoxic salamanders with a red coloration after exposure to toxic efts. Repeated examination of possible salamander mimics indicates that this dynamic is not limited to a small region of the range of Notophthalmus or to a single species of mimic, but may involve a much larger portion of the range of the genus and multiple mimic species [134]. A similar mimicry system associated with Taricha may also be present in western North American species. Ensatina eschscholtzii is a small nontoxic species found throughout western North America. This species forms a well-documented ring complex in California that is marked by extreme variation in morphology and color pattern [137, 138, 156–158]. The range of this species overlaps two species of Taricha including Taricha torosa [138]. Ensatina eschscholtzii from this overlapping region has a recognized subspecies (E. eschscholtzii xanthoptica) that possesses a morphology and color patterns that are similar to T. torosa (e.g., yellow eyes as well as red dorsal coloration) (Plate 3). Furthermore, this similarity appears to protect them from predation [137, 138]. These patterns of mimicry are striking and suggest that parallel ecological dynamics associated with mimicry in Taricha and Notophthalmus are likely to be important in understanding the larger picture of toxicity in these genera. A combination of bio-sequestration and warning color may also be in play with garter snake predators of Taricha (Plate 4). Williams demonstrated that Thamnophis from populations that co-occur with highly toxic Taricha granulosa, maintains measurable levels of 1 in the liver and other tissues past a period of time that

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Plate 3 Mimics of Notophthalmus (a–e) and Taricha (f). a N. viridescens viridescens (bottom) and Pseudotriton ruber (top), b Pseudotriton ruber warning display, c Eurycea lucifuga, d Plethodon cinereus, e Gyrinophilus porphyriticus, f Ensatina eschscholtzii xanthoptica. Photographs a–e courtesy of E. D. Brodie Jr., f courtesy of R.W. Van Devender

Plate 4 Thamnophis sirtalis. a Thamnophis sirtalis attacking Taricha granulosa, b tetrodotoxinresistant Thamnophis sirtalis—note extensive red dorsal markings

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would be predicted by simple pharmacokinetic clearing after exposure [159]. There is also anecdotal evidence that these garter snakes have an increase in distinctive red coloration, which raises the possibility that these snakes are harvesting newt chemical defenses for their own use, similar to toad-eating Rhabdophis tigrinus snakes [160]. Finally, Zimmer (2007) has proposed that tetrodotoxin (1) should be viewed as a “keystone” molecule. Specifically, he argues, as with a keystone species, that 1 and other guanidine metabolites have impacts far beyond their “biomass”. This perspective emphasizes the role of 1 as a signaling molecule and identifies its role as a chemosensory cue for predation risk for larval newts and as a general signal that “mediates trophic interactions” [54]. Certainly, the broad trophic impacts of 1 described here are congruent with this perspective and further extend the likelihood that understanding the chemical ecology of North American newts requires a broad ecological perspective.

10 Conclusions Tetrodotoxin (1) plays a central role in the chemical ecology of North American newts. The potent pharmacology of this toxin is coupled with aposematic coloration and defensive postures to form a complex phenotype that defends these organisms from attack and predation throughout their life histories. This phenotype is also ancient and 1 has likely played an important role in the chemical ecology of Taricha and Notophthalmus since the origin of these genera, sometime in the mid to late Paleogene (30–40 Ma). Although the origin of 1 present in these taxa is still unresolved recent analyses of newt physiology and biochemistry are congruent with a possible endogenous biosynthesis pathway. Tetrodotoxin (1) in North American newts sits at the center of an extended web of species interactions that includes coevolution with snakes, potential egg predation by insects, complex conspecific interactions, as well as extended mimicry complexes. Levels of 1 are highly variable among populations of these genera and the dynamics and ecological interactions that give rise to this variation are still poorly understood. Although coevolution with Thamnophis sirtalis is likely to be an important determinant of levels of 1 in adult Taricha from some populations, it does not explain much of the measured variation throughout the range of the genus. Nor does it explain patterns of variation in Notophthalmus. The model proposed here suggests that developing a complete understanding of the chemical ecology of these genera will require an understanding of the impact of maternal effects associated with egg predation and provisioning, the role of 1 as a defense against parasites, and the overlapping interactions of multiple coevolutionary arms-races with other snake predators. Finally, understanding the potential value of low levels of 1 throughout the ranges of these genera will require a better understanding of the Batesian mimicry complexes that are associated with the tetrodotoxin-bearing phenotype in Taricha and Notophthalmus as well as the complex dynamics that result from the interplay of these interactions.

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Acknowledgments Sophia Hanifin provided original line illustrations for Fig. 1. Charles T. Hanifin would like to thank E. D. Brodie Jr. for extensive conversations and input associated with this chapter. In addition, he provided all photographs except the one of Ensatina eschscholtzii xanthoptica, which is due to R.W. Van Devender. Work associated with the biochemistry, pharmacology, and analysis of tetrodotoxin was funded (in part) by the Japan Society for the Promotion of Science (JSPS) through its KAKENHI Grant-in-Aid for Innovative Area, Frontier Research on Redesigning Biosynthetic Machineries (no. JP19H04636) (to Y.K.). Other support was from a Chemical Communications grant (no. JP17H06406), a grant for Scientific Research (no. JP20H02921), a grant for Exploratory Research (no. JP19K22266), grants-in-aid for Research Activity start-up (JP18H05997 and JP19K21141), and a grant for Young Scientists (JP20K15405) (all to Y.K.). This work was supported also by the Uehara Memorial Foundation (to M.Y.Y.) and the Naito Foundation (to Y.K.).

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148. Twitty VC (1966) Of scientists and salamanders. WH Freeman & Co, New York, NY, USA 149. Lehman EM (2006) Egg toxicity and egg predation in rough-skinned newts, Taricha granulosa. Master’s thesis, Indiana University, Bloomington, IN, USA 150. Gall BG, Stokes AN, French SS, Schlepphorst EA, Brodie ED III, Brodie ED Jr (2011) Tetrodotoxin levels in larval and metamorphosed newts (Taricha granulosa) and palatability to predatory dragonflies. Toxicon 57:978 151. Jacisin JJ, Hopkins SSB (2018) A redescription and phylogenetic analysis based on new material of the fossil newts Taricha oligocenica Van Frank, 1955 and Taricha lindoei Naylor, 1979 (Amphibia, Salamandridae) from the Oligocene of Oregon. J Paleontol 92:713 152. Thorp JH, Wallace JB, Georgian TJ Jr (1986) Untangling the web of caddisfly evolution and distribution. Oikos 47:253 153. Mebs D, Yotsu-Yamashita M, Seitz HM, Arakawa O (2012) Tetrodotoxin does not protect red-spotted newts, Notophthalmus viridescens, from intestinal parasites. Toxicon 60:66 154. Mebs D, Yotsu-Yamashita M, Hartmann K, Elbert C, Zehner R, Toennes SW (2020) Revisited-failure of tetrodotoxin to protect red-spotted newts, Notophthalmus viridescens, from endoparasites. Toxicon 178:77 155. Schafer W (2016) Nematode nervous systems. Curr Biol 26:R955 156. Highton R (1998) Is Ensatina eschscholtzii a ring-species? Herpetologica 54:254 157. Kuchta SR, Parks DS, Mueller RL, Wake DB (2009) Closing the ring: historical biogeography of the salamander ring species Ensatina eschscholtzii. J Biogeogr 36:982 158. Moritz C, Schneider CJ, Wake DB (1992) Evolutionary relationships within the Ensatina eschscholtzii complex confirm the ring species interpretation. Syst Biol 41:273 159. Williams BL, Brodie ED Jr (2004) A resistant predator and its toxic prey: persistence of newt toxin leads to poisonous (not venomous) snakes. J Chem Ecol 30:1901 160. Hutchinson DA, Mori A, Savitzky AH, Burghardt GM, Wu X, Meinwald J, Schroeder FC (2007) Dietary sequestration of defensive steroids in nuchal glands of the Asian snake Rhabdophis tigrinus. Proc Natl Acad Sci USA 104:2265

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Charles T. Hanifin obtained a Ph.D. degree in Ecology and Evolutionary Biology at Utah State University, where he worked on the chemical ecology of tetrodotoxin-bearing Taricha newts and their coevolution with tetrodotoxin-resistant garter snakes. Subsequently, he held a National Institutes of Health (NIH, USA) postdoctoral fellowship at the Hopkins Marine Station (Stanford University) where he examined the evolution of voltage-gated sodium channels and organismal resistance to tetrodotoxin in the Salamandridae. He is currently an Associate Professor in the Department of Biology at Utah State University (Uintah Basin Campus). His research adopts an integrative approach to understand adaptive evolution in voltage-gated sodium channels associated with the evolution of tetrodotoxin resistance in tetrodotoxin-bearing organisms. In addition, his work explores questions about the effect of variation in tetrodotoxin stereoisomers on landscape level patterns of coevolution with garter snake predators and the general defensive ecology of Taricha. His work on exploring the coevolution between Taricha newts and garter snake predators is widely disseminated and has become a textbook example of arms-race coevolution. Yuta Kudo received his Ph.D. degree in Agriculture from Tohoku University in 2016 under the supervision of Prof. Mari Yotsu-Yamashita. As a Japan Society for Promotion of Science Overseas Research Fellow, he held a postdoctoral position at the University of California, San Diego, Scripps Institution of Oceanography in the laboratory of Prof. Bradley S. Moore, where he investigated the biosynthesis of bacterial secondary metabolites. In 2018, he became a Specially Appointed Assistant Professor at the Graduate School of Agricultural Sciences, Tohoku University in the laboratory of Prof. Mari Yotsu-Yamashita. Currently, he is an Assistant Professor at the Frontier Research Institute for Interdisciplinary Sciences and Graduate School of Agricultural Sciences, Tohoku University in Prof. Yotsu-Yamashita’s laboratory. His research interests include the isolation, structural elucidation, and biosynthesis of unique natural products. His main research focus has been the screening of new tetrodotoxin-related compounds from toxic newts to elucidate the biosynthetic pathway of tetrodotoxin in terrestrial environments.

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C. T. Hanifin et al. Mari Yotsu-Yamashita received her Bachelor’s degree in Agriculture from Tohoku University in 1984 and started working as a junior researcher (technical official) at this same institution. She received her Ph.D. degree in Agriculture in 1989 from Tohoku University under the supervision of Prof. Takeshi Yasumoto. She joined the Department of Chemistry, Ohio State University, as a Visiting Scholar, to study synthesis under the direction of Prof. Viresh H. Rawal during 1994–1995. She was promoted to Associate Professor of Tohoku University in 1998 (Prof. Teruo Miyazawa’s group), and then to Full Professor in 2004. Her research is focused mainly on the chemistry of biologically active natural products, especially marine natural toxins such as tetrodotoxin, saxitoxin, domoic acid, and polycavernosides, through their isolation, structural determination, biosynthesis, and mode of action, and and also on the development of analytical methods.

The Genus Walsura: A Rich Resource of Bioactive Limonoids, Triterpenoids, and Other Types of Compounds Ninh The Son

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phytochemical Investigations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Limonoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Triterpenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Sesquiterpenoids, Sterols, and Lignans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Simple Phenols, Flavonoids, Xanthones, Anthraquinones, and Other Types of Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Biological Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Cytotoxic Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Antimicrobial Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Antidiabetic Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Anti-inflammatory Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Antioxidative Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Antifeedant and Other Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Synthesis Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

131 132 142 148 153 155 157 157 168 169 170 170 171 171 173 173

1 Introduction Plants of the genus Walsura, comprising 30 to 40 species, are distributed widely in several countries of Asia [1]. As evidenced from its traditional applications, Walsura trichostemon, locally named “Lamyai Pa”, has been used in Thai folk medicine to treat tendon disabilities, hemorrhoids, to stop excessive blood flow, and to clean N. T. Son (B) Department of Applied Biochemistry, Institute of Chemistry, Vietnam Academy of Science and Technology, 18-Hoang Quoc Viet, Caugiay, Hanoi, Vietnam © The Author(s), under exclusive license to Springer Nature Switzerland AG 2022 A. D. Kinghorn, H. Falk, S. Gibbons, Y. Asakawa, J.-K. Liu, V. M. Dirsch (eds.), Progress in the Chemistry of Organic Natural Products 118, https://doi.org/10.1007/978-3-030-92030-2_4

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N. T. Son

wounds [2]. The bark of W. piscidia has been employed for its stimulant, expectorant, emmenagogue, and emetic effects, while its fruits are used to poison fish as a trapping method [3]. In China, W. robusta leaves and twigs have been utilized as an insecticide [4]. As with other genera of the family Meliaceae, Walsura species are known for the presence of structurally diverse secondary metabolites such as simple phenols, lignans, sesquiterpenoids, and, in particular, limonoids and triterpenoids [5–7]. Walsura plant extracts and their isolated compounds have a wide spectrum of biological effects, such as cytotoxic, antimicrobial, antioxidative, antifeedant, and neuroprotective activities [8–12]. Although secondary metabolites derived from Walsura medicinal plants are well-known for their traditional uses, to date no specific reviews on the bioactive constituents of this genus have been published. The current chapter deals with almost 50 years of research on the genus Walsura, and provides a general insight into its prior phytochemical and biological testing research. In addition to consulting various journal articles, the Web of Science, the Plant List, Scopus, SciFinder, Google Scholar, PubMed, Science Direct, Chemical Abstracts, the Wiley Online Library, and the IOP Science databases were used for literature searching.

2 Phytochemical Investigations The current contribution gives an overview of the phytochemicals derived from Walsura species, as purified using various chromatographic techniques. Nine plant species, including W. chrysogyne, W. cochinchinensis, W. pinnata, W. piscidia, W. robusta, W. trichostemon, W. trifoliata, W. tubulata, and W. yunnanensis, (Fig. 1) have been the main subjects of phytochemical studies. A total of 223 secondary metabolites are compiled in Table 1 and Figs. 2, 3, 4, and 5. These isolated secondary metabolites derived from Walsura species may be categorized into a broad variety of structural classes, including limonoids, triterpenoids, sesquiterpenoids, sterols, lignans, simple phenols, flavonoids, anthraquinones, xanthones, and other types. Compounds 1–114 are limonoids [1, 5–7, 13–25]. In turn, natural products 115–187 are assigned to the triterpenoid group [2, 3, 5, 7, 10, 11, 15, 17, 20, 22, 24, 26–41]. Compounds 188–194 fall into the sesquiterpenoids [4, 12, 31, 32, 34, 39], and compounds 195–197 can be

Fig. 1 Representative Walsura species: W. cochinchinensis (a), W. robusta (b), and W. pinnata (c)

The Genus Walsura: A Rich Resource …

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Table 1 Chemical constituents of the genus Walsura No.

Compound

Species

Refs.

Limonoids 1

12α-Acetoxycedrelone

W. robusta fruits

[24]

2

11β-Acetoxydihydrocedrelone

W. robusta leaves and fruits, W. yunnanensis bark

[7, 18, 24]

3

11β-Acetoxywalsuranolide

W. yunnanensis bark

[18]

4

Anthothecol

W. robusta fruits

[24]

5

Cedrelone

W. robusta fruits, W. yunnanensis bark

[18, 24]

6

Cochinchinoid A

W. cochinchinensis twigs and leaves

[15]

7

Cochinchinoid B

W. cochinchinensis twigs and leaves

[15]

8

Cochinchinoid C

W. cochinchinensis twigs and leaves

[15]

9

Cochinchinoid D

W. cochinchinensis twigs and leaves

[15]

10

Cochinchinoid E

W. cochinchinensis twigs and leaves

[15]

11

Cochinchinoid F

W. cochinchinensis twigs and leaves

[25]

12

Cochinchinoid G

W. cochinchinensis twigs and leaves

[15]

13

Cochinchinoid H

W. cochinchinensis twigs and leaves

[15]

14

Cochinchinoid I

W. cochinchinensis twigs and leaves

[15]

15

Cochinchinoid J

W. cochinchinensis twigs and leaves

[15]

16

7-Deacetoxy-7hydroxyazadirone

W. piscidia fruits

[12]

17

Dihydrocedrelone

W. robusta fruits

[24]

18

1,2-Dihydro-7-deacetoxy-7hydroxyazadirone

W. piscidia fruits

[22]

19

1α,11β-Dihydroxy-1,2dihydroisowalsuranolide

W. yunnanensis twigs and leaves

[17]

20

11β,12α-Diacetoxycedrelone

W. robusta fruits

[24]

21

20,22-Dihydro-22,23epoxywalsuranolide

W. yunnanensis bark

[18]

22

Dysoxylumosin B

W. robusta fruits

[24] (continued)

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N. T. Son

Table 1 (continued) No.

Compound

Species

Refs.

23

Dysoxylumosin C

W. robusta fruits

[24]

24

Dysoxylumosin D

W. robusta fruits

[24]

25

Dysoxylumosin G

W. robusta fruits

[24]

26

1α-Ethoxy-11βhydroxydihydrocedrelone

W. robusta fruits

[24]

27

11β-Hydroxycedrelone

W. robusta fruits, W. [18, 24, 25] yunnanensis twigs, leaves, and bark

28

11β-Hydroxydihydrocedrelone

W. robusta leaves and fruits, W. yunnanensis bark

[7, 18, 24]

29

11β-Hydroxy-1α-methoxy1,2-dihydroisowalsuranolide

W. yunnanensis twigs and leaves

[17]

30

11β-Hydroxy-1,2dihydroisowalsuranolide

W. robusta leaves, W. yunnanensis twigs and leaves

[7, 17]

31

11β-Hydroxy-23-Omethylwalsuranolide

W. robusta leaves, W. yunnanensis twigs and leaves

[7, 17]

32

11β-Hydroxyisowalsuranolide

W. yunnanensis twigs and leaves

[17]

33

Isowalsurinolide

W. robusta leaves and fruits, W. yunnanensis bark

[7, 18]

34

1α-Methoxy-12αacetoxydihydrocedrelone

W. robusta fruits

[24]

35

1α-Methoxy-11βhydroxydihydrocedrelone

W. robusta fruits

[24]

36

11-Oxo-dihydrocedrelone

W. piscidia fruits

[22]

37

Piscidofuran

W. robusta fruits

[24]

38

Walrobsin A

W. robusta root bark

[23]

39

Walrobsin B

W. robusta root bark

[23]

40

Walrobsin C

W. robusta root bark

[14]

41

Walrobsin D

W. robusta root bark

[14]

42

Walrobsin E

W. robusta root bark

[14]

43

Walrobsin F

W. robusta root bark

[14]

44

Walrobsin G

W. robusta root bark

[14]

45

Walrobsin H

W. robusta root bark

[14]

46

Walrobsin I

W. robusta root bark

[14]

47

Walrobsin J

W. robusta root bark

[14]

48

Walrobsin K

W. robusta root bark

[14]

49

Walrobsin L

W. robusta root bark

[14]

50

Walrobsin M

W. robusta root bark

[14] (continued)

The Genus Walsura: A Rich Resource …

135

Table 1 (continued) No.

Compound

Species

Refs.

51

Walrobsin N

W. robusta root bark

[14]

52

Walrobsin O

W. robusta root bark

[14]

53

Walrobsin P

W. robusta root bark

[14]

54

Walrobsin Q

W. robusta root bark

[14]

55

Walrobsin R

W. robusta root bark

[14]

56

Walsogyne A

W. chrysogyne bark

[21]

57

Walsogyne B

W. chrysogyne bark

[19]

58

Walsogyne C

W. chrysogyne bark

[19]

59

Walsogyne D

W. chrysogyne bark

[19]

60

Walsogyne E

W. chrysogyne bark

[19]

61

Walsogyne F

W. chrysogyne bark

[19]

62

Walsogyne G

W. chrysogyne bark

[19]

63

Walsucochinone A

W. cochinchinensis bark

[6]

64

Walsucochinone B

W. cochinchinensis bark

[6]

65

Walsucochinone C

W. cochinchinensis bark

[6]

66

Walsucochinoid A

W. cochinchinensis twigs and leaves

[1]

67

Walsucochinoid B

W. cochinchinensis twigs and leaves

[1]

68

Walsucochinoid C

W. cochinchinensis twigs and leaves

[16]

69

Walsucochinoid D

W. cochinchinensis twigs and leaves

[16]

70

Walsucochinoid E

W. cochinchinensis twigs and leaves

[16]

71

Walsucochinoid F

W. cochinchinensis twigs and leaves

[16]

72

Walsucochinoid G

W. cochinchinensis twigs and leaves

[16]

73

Walsucochinoid H

W. cochinchinensis twigs and leaves

[16]

74

Walsucochinoid I

W. cochinchinensis twigs and leaves

[16]

75

Walsucochinoid J

W. cochinchinensis twigs and leaves

[16]

76

Walsucochinoid K

W. cochinchinensis twigs and leaves

[16]

77

Walsucochinoid L

W. cochinchinensis twigs and leaves

[16] (continued)

136

N. T. Son

Table 1 (continued) No.

Compound

Species

Refs.

78

Walsucochinoid M

W. cochinchinensis twigs and leaves

[16]

79

Walsucochinoid N

W. cochinchinensis twigs and leaves

[16]

80

Walsucochinoid O

W. cochinchinensis twigs and leaves

[16]

81

Walsucochinoid P

W. cochinchinensis twigs and leaves

[16]

82

Walsucochinoid Q

W. cochinchinensis twigs and leaves

[16]

83

Walsucochinoid R

W. cochinchinensis twigs and leaves

[16]

84

Walsunoid A

W. robusta leaves

[7]

85

Walsunoid B

W. robusta leaves

[7]

86

Walsunoid C

W. robusta leaves

[7]

87

Walsunoid D

W. robusta leaves

[7]

88

Walsunoid E

W. robusta leaves

[7]

89

Walsunoid F

W. robusta leaves

[7]

90

Walsunoid G

W. robusta leaves

[7]

91

Walsunoid H

W. robusta leaves and fruits

[7, 24]

92

Walsunoid I

W. robusta leaves

[7]

93

Walsuranin B

W. robusta root bark

[14]

94

Walsuranolide

W. yunnanensis bark

[18]

95

Walsuranolide B

W. yunnanensis twigs and leaves

[17]

96

Walsurin

W. robusta fruits, W. yunnanensis bark

[18, 24]

97

Walsurin A

W. robusta fruits

[24]

98

Walsurin B

W. robusta fruits

[24]

99

Walsurin C

W. robusta fruits

[24]

100

Walsurin D

W. robusta fruits

[24]

101

Walsurin E

W. robusta fruits

[24]

102

Walsuronoid A

W. robusta twigs, leaves and root bark

[14, 23]

103

Walsuronoid B

W. robusta twigs, leaves

[23]

104

Walsuronoid C

W. robusta twigs and leaves

[23]

105

Walsuronoid D

W. robusta leaves

[5]

106

Walsuronoid E

W. robusta leaves

[5] (continued)

The Genus Walsura: A Rich Resource …

137

Table 1 (continued) No.

Compound

Species

Refs.

107

Walsuronoid F

W. robusta leaves

[24]

108

Walsuronoid G

W. robusta leaves

[24]

109

Walsuronoid H

W. robusta leaves

[24]

110

Walsuronoid I

W. robusta leaves

[24]

111

Yunnanol

W. yunnanensis twigs and leaves

[17]

112

Yunnanolide A

W. yunnanensis twigs and leaves

[17]

113

Yunnanolide B

W. yunnanensis twigs and leaves

[17]

114

New unamed limonoida

W. trichostemon stems

[20]

Triterpenoids 115

7α-Acetoxy-17α-(20S),21,24- W. robusta fruits epoxy-apotirucall-14-en-3one-(23R,24S),25-triol

[24]

116

Apowalsogyne A

W. chrysogyne bark

[30]

117

Apowalsogyne B

W. chrysogyne bark

[30]

118

Betulin

W. robusta fruits

[24]

119

Betulonic acid

W. pinnata bark and stem bark, W. robusta fruits

[24, 29, 32, 39]

120

Cabraleadiol

W. chrysogyne leaves

[34]

121

Chrysura

W. chrysogyne leaves

[33, 34]

122

Cochinchinoid K

W. cochinchinensis twigs and leaves

[15]

123

Cycloart-23-ene-3β,25-diol

W. chrysogyne leaves

[34]

124

(24S,25R)-Cycloartane3β,24,25,26-tetrol

W. robusta leaves, W. yunnanensis twigs and leaves

[5, 17]

125

7-Deacetylbrujavanone E

W. trichostemon leaves

[37]

126

11,25Dideacetyltrichostemonate

W. trichostemon leaves and stems

[20, 37]

127

11α,20-Dihydroxydammar24-ene-3-one

W. trichostemon stem bark

[10]

128

Dihydroniloticin

W. robusta leaves

[7]

129

Dymalol

W. chrysogyne leaves

[34]

130

Eichlerianic acid

W. chrysogyne leaves

[34]

131

3-Epimesendanin S

W. cochinchinensis twigs and leaves, W. trichostemon roots, W. robusta leaves

[2, 7, 15]

(continued)

138

N. T. Son

Table 1 (continued) No.

Compound

132

3-Epimesendanin S 12-acetate W. trichostemon roots

Species

[2]

Refs.

133

(20R,24S)-Epoxy-25hydroxydammaran-3-one

W. chrysogyne leaves

[34]

134

(22S,23R)-Epoxytirucalla-7ene-3α,24,25-triol

W. robusta leaves

[7]

135

Foveolin

W. chrysogyne leaves

[34]

136

Friedelanone

W. trichostemon twigs

[38]

137

Grandifolinolenenone

W. robusta fruits, W. trichostemon stem bark

[10, 24]

138

Hispidol B

W. robusta fruits

[24]

139

12α-Hydroxy-3-oxooleanano- W. pinnata stem bark 28,13-lactone

140

3β-Hydroxy-5-glutinen-28oic acid

W. pinnata bark and stem bark [31, 32, 39]

141

3-Ketours-11-en-13β(28)olide

W. pinnata stem bark

[39]

142

Lupeol

W. trifoliata leaves

[35]

143

Niloticin

W. robusta leaves

[7]

144

Melianone

W. trichostemon twigs

[38]

145

Mesendanin S

W. cochinchinensis twigs and leaves

[15]

146

Mesendanin G

W. trichostemon roots

[2]

147

25-Methoxycycloart-23-en3β-ol

W. chrysogyne leaves

[34]

148

Methyl eichlerianate

W. chrysogyne leaves

[34]

149

Oleanonic acid

W. pinnata stem bark

[39]

150

3-Oxo-20(24)-epoxy-12β,25- W. pinnata bark dihydroxydammarane

[31, 32]

151

3-Oxo-lup-20(29)-en-28-oic acid

W. pinnata bark

[31]

152

3-Oxo-olean-11-en-28,13βolide

W. pinnata bark and stem bark [31, 39]

153

3-Oxo-olean-9(11),12-dien28-oic acid

W. pinnata bark

154

3-Oxoursolic acid

W. pinnata stem bark

155

Pinnatane A

W. pinnata bark

[40]

156

Piscidenone

W. piscidia leaves

[3]

[39]

[31, 32, 39]

(continued)

The Genus Walsura: A Rich Resource …

139

Table 1 (continued) No.

Compound

Species

157

Piscidinol A

W. piscidia leaves, W. trifoliata [3, 7, 32, 35] leaves, W. robusta leaves

Refs.

158

Piscidinol B

W. piscidia leaves

[27, 35]

159

Piscidinol C

W. piscidia leaves, W. trifoliata leaves

[3, 22, 27, 28, 35]

160

Piscidinol D

W. piscidia leaves, W. trifoliata leaves

[32, 37, 46]

161

Piscidinol E

W. piscidia leaves, W. trifoliata leaves

[22, 27, 35]

162

Piscidinol F

W. piscidia leaves

[27]

163

Piscidinol G

W. piscidia leaves

[3]

164

Piscidinol H

W. trifoliata leaves

[35]

165

Piscidinol I

W. trifoliata leaves

[35]

166

Piscidinol J

W. trifoliata leaves

[35]

167

Piscidinol K

W. trifoliata leaves

[35]

168

Piscidinol L

W. trifoliata leaves

[35]

169

Piscidinone A

W. trifoliata twigs and leaves

[11]

170

Piscidinone B

W. trifoliata twigs and leaves

[11]

171

Sapelin E acetate

W. trichostemon stem bark

[10]

172

Spicatin

W. robusta fruits

[24]

173

21,24,25-Triacetyl-7deacetyl-6hydroxybrujavanone E

W. trichostemon leaves

[37]

174

Trichostemonate

W. trichostemon roots

[36]

175

Trichostemonoate

W. trichostemon stem bark

[10]

176

Trichostemonol

W. trichostemon twigs

[38]

177

12β,(20S,24R)Trihydroxydammar-25-en-3one

W. trichostemon stems

[20]

178

12β,(20S),25Trihydroxydammar-23-en-3one

W. trichostemon stems

[20]

179

Walsucochin A

W. cochinchinensis twigs and leaves

[41]

180

Walsucochin B

W. cochinchinensis twigs and leaves

[41]

181

Walsurenol

W. tubulata leaves

[26] (continued)

140

N. T. Son

Table 1 (continued) No.

Compound

Species

Refs.

182

12β,(20S)Dihydroxydammar-23-en-3one derivativea

W. trichostemon stems

[20]

183

12β,(20S)Dihydroxydammar-3-one derivativea

W. trichostemon stems

[20]

184

12β,(20S)Dihydroxydammar-3-one derivative

W. trichostemon stems

[20]

185

12β,(20S)Dihydroxydammar-3-one derivative

W. trichostemon stems

[20]

186

Apotirucallane derivativea

W. trichostemon stems

[20]

187

Apotirucallane derivativea

W. trichostemon stems

[20] [32, 47]

Sesquiterpenoids 188

2(3),6(7)-Diepoxy-9humulene

W. pinnata bark

189

Ledol

W. pinnata stem bark

190

10β-Nitro-isodauc-3-en-15-al W. robusta leaves

191

Oplopanone

W. pinnata stem bark

[39]

192

10-Oxo-isodauc-3-en-15-al

W. robusta leaves

[4]

193

Turpinionoside A

W. robusta leaves and twigs

[12]

194

Viridiflorol

W. chrysogyne leaves

[34]

195

β-Sitosterol

W. trichostemon twigs and roots

[2, 38]

196

β-Sitosterol glucoside

W. pinnata bark, W. trichostemon roots

[2, 31, 32]

197

Stigmasterol

W. pinnata bark

[31, 32]

[39] [4]

Sterols

Lignans 198

Brassilignan

W. robusta leaves

[5]

199

9-Deoxyarctigenin

W. robusta leaves

[5]

200

Lariciresinol

W. cochinchinensis bark

[6]

201

(+)-Lyoniresinol 3α-O-β-d-glucopyranoside

W. robusta leaves and twigs

[12]

202

(–)-Lyoniresinol 3α-O-β-d-glucopyranoside

W. robusta leaves and twigs

[12]

203

Phyllanthin

W. cochinchinensis bark

[6] (continued)

The Genus Walsura: A Rich Resource …

141

Table 1 (continued) No.

Compound

Species

Refs.

204

[(1R,2S,3S)-1,2,3,4Tetrahydro-7-hydroxy-1-(4hydroxy-3-methoxyphenyl)3-(hydroxymethyl)-6methoxynaphthalen-2yl]methyl-α-larabinopyranoside

W. yunnanensis bark

[8]

205

[(1S,2R,3R)-1,2,3,4Tetrahydro-7hydroxy-1-(4-hydroxy-3,5dimethoxyphenyl)-3(hydroxymethyl)-6methoxynaphthalen-2yl]methyl-α-larabinopyranoside

W. yunnanensis bark

[8]

Simple phenols 206

Ferulaldehyde

W. cochinchinensis bark

[6]

207

Methyl vanillate

W. cochinchinensis bark

[6]

208

Protocatechuic acid

W. cochinchinensis bark

[6]

209

Sinapaldehyde

W. cochinchinensis bark

[6]

210

3,4,5-Trimethoxyphenyl-β-d- W. robusta twigs and leaves, glucopyranoside W. yunnanensis bark

[8, 12]

211

3,4,5-Trimethoxyphenyl W. yunnanensis bark 2-O-(α-l-fucopyranosyl)-β-dglucopyranoside

[8]

212

Vanillic acid

W. cochinchinensis bark; W. pinnata bark

[6, 31, 32]

213

Walsuraside

W. yunnanensis bark

[8]

Flavonoids 214

3,5,6,7,8,3 ,4 Heptamethoxyflavone

W. cochinchinensis bark

[6]

215

(2R,3R)-10-(3,4-Dihydroxyphenyl)-2-(3,5dihydroxyphenyl)-3,5dihydroxy-3,4,9,10tetrahydro-2H,8Hpyrano[2,3-f ]chromen-8-one

W. trifoliata bark

[47]

216

(2R,3S)10-(3,4-Dihydroxy-phenyl)2-(3,5-dihydroxyphenyl)-3,5dihydroxy-3,4,9,10tetrahydro-2H,8Hpyrano[2,3-f ]chromen-8-one

W. trifoliata leaves

[47]

(continued)

142

N. T. Son

Table 1 (continued) No.

Compound

Species

Refs.

Anthraquinone and xanthone derivatives 217

4,8,10-Trihydroxy-10-(2W. trifoliata bark hydroxyacetoxy)-9-oxo-9,10dihydroanthracene-2carboxylic acid 10-glycoside

[45]

218

Chrysophanol

W. trichostemon twigs

[38]

219

Mangostenone F

W. trichostemon bark and twigs

[10, 38]

220

α-Mangostin

W. trifoliata bark

[45]

Other Compounds 221

Asperglaucide

W. yunnanensis bark

[8]

222

Butyl α-d-fructofuranoside

W. yunnanensis bark

[8]

223

4-Hydroxy-4,8-dimethyl-1tetralone

W. pinnata bark

[31, 45]

a Compound

was unnamed

assigned as sterols [2, 31, 32, 38]. Lignans are well represented (198–205) [5, 6, 8, 12], and structures 206–213 have been categorized as miscellaneous phenol derivatives [6, 8, 12, 31, 32]. Although flavonoids are abundant in the plant kingdom they have been reported only rarely in the genus Walsura, with only the three compounds 214– 216 reported thus far [6, 42, 43]. Similarly, compounds 217–220 are anthraquinone or xanthone derivatives [10, 42, 44]. Walsura species also produce the additional compounds, 221–223 [8, 41, 45].

2.1 Limonoids Limonoids are characteristic natural products that represent a high structural diversification within the genera of the family Meliaceae. Limonoids derived from Meliaceae species can be divided into those with 35 different carbon skeletons, in which metabolites with a 4,4,8-trimethyl-17-furanyl steroid backbone are the major isolated compounds [48]. In turn, Walsura species are a rich source of limonoids. In the current subsection, a detailed list of 114 isolated compounds (1–114) is summarized in Table 1 and Fig. 2 [1, 5–7, 13–25]. Walsura chrysogyne, W. piscidia, W. robusta, W. trichostemon, W. yunnanensis, and W. cochinchinensis are the main plants found to produce limonoids. It may be noted that many of these were new natural products when first characterized. They represent various skeletons, such as cedrelone in 11β-acetoxydihydrocedrelone (2), vilasinin in cochinchinoid A (6), a seco-ring D type in cochinchinoid E (10), an azadirone in cochinchinoid H

The Genus Walsura: A Rich Resource …

143 R4

23

R1

11

9

1

R2

10

2

16

O

R3

R2 O

O

O

O

O

6

4

R1

17 14 15

8

5

3

20

13

7

O

21

12

R3

O

O

22

R4

OH

OH

1 R1 + R2 = Δ1,2, R3 = H, R4 = α -OAc 2 R1 = R2 = R4 = H, R3 = β -OAc 4 R1 + R2 = Δ1,2, R3 = α -OAc, R4 = H 5 R1 + R2 = Δ1,2, R3 = R4 = H 17 R1 = R2 = R3 = R4 = H 20 R1 + R2 = Δ1,2, R3 = β -OAc, R4 = α -OAc 25 R1 = α -OMe, R2 = R4 = H, R3 = β -OAc 26 R1 = α -OEt, R2 = R4 = H, R3 = β -OH 27 R1 + R2 = Δ1,2, R3 = H, R4 = α -OAc 28 R1 = OMe, R2 = R4 = H, R3 = β -OH 34 R1 = α -OMe, R2 = R3 = H, R4 = α -OAc 35 R1 = α -OMe, R2 = R4 = H, R3 = β -OH 37 R1 = R2 = R4 = H, R3 = C=O 91 R1 + R2 = Δ1,2, R3 = β -OAc, R4 = H

3 R1 + R2 = Δ1,2, R3 = OAc, R4 = 23-OH 31 R1 + R2 = Δ1,2, R3 = OH, R4 = OMe 32 R1 + R2 = Δ1,2, R3 = OH, R4 = α -OH 33 R1 + R2 = Δ1,2, R3 = H, R4 = 21-OH 85 R1 + R2 = Δ1,2, R3 = OH, R4 = β -OMe 86 R1 + R2 = Δ1,2, R3 = R4 = OH 89 R1 + R2 = Δ1,2, R3 = OAc, R4 = 21-OH 90 R1 = R2 = H, R3 = OAc, R4 = OH 94 R1 + R2 = Δ1,2, R3 = H, R4 = ?-OH 95 R1 + R2 = Δ1,2, R3 = OH, R4 = H

O

O R3 AcO OAc

R4

OH

O

O O

R1O

OAc

RO

OR2

O

O

O

10 R = X3 11 R = X1 12 R = X2

6 R1 = R2 = X1, R3 = H, R4 = OH 7 R1 = R2 = X2, R3 = H, R4 = OH 8 R1 = R2 = X3, R3 = R4 = H 9 R1 = X3, R2 = X4, R3 = OAc, R4 = H

O

O R5 R1

R4 O

R2 O

OR3

13 R1 + R2 = Δ1,2, R3 = X4, R4 = OAc, R5 = H 14 R1 + R2 = Δ1,2, R3 = X4, R4 = H, R5 = OAc 15 R1 = R2 = R4 = H, R3 = X4, R5 = OAc

Fig. 2 Limonoids from the genus Walsura

R1 R2 O

OR3

16 R1 + R2 = Δ1,2, R3 = H 18 R1 = R2 = R3 = H

144

N. T. Son O

O

O

O

O

O

R2

HO R1

O

19 R = OH, R = OH 29 R1 = OMe, R2 = OH 30 R1 = H, R2 = OH 87 R1 = H, R2 = OMe 88 R1 = R2 = H

O

36

21

O

O

R4 R1

OH

OH

2

O

O AcO

O

O

OH 1

O

O

O O

AcO OX3

O

HO

O

O

R3

R1O

R2 AcO

OH

R2

O

O

R4

OH

O

O

OH

R3

40 22 R1 = α -OAc, R2 = R4 = H, R3 = β-OH 23 R1 + R2 = Δ1,2, R3 + R4 = Δ11,12 24 R1 = α -OAc, R2 = H, R3 + R4 = Δ11,12 103 R1 + R2 = Δ1,2, R3 = β -OH, R4 = H 108 R1 = OEt, R2 = H, R3 = β -OH, R4 = H 109 R1 = R2 = R3 = R4 = H

38 R1 = X3, R2 + R3 = O, R4 = H 39 R1 = X2, R2 + R3 = O, R4 = H 47 R1 = X2, R2 = OH, R3 = R4 = H 48 R1 = X3, R2 = OAc, R3 = R4 = H 49 R1 = X2, R2 = OAc, R3 = R4 = H 50 R1 = X3, R2 = OH, R3 = H, R4 = OAc 51 R1 = X2, R2 = OH, R3 = H, R4 = OAc 52 R1 = X3, R2 = OAc, R3 = H, R4 = OH 53 R1 = R4 = OH, R2 = OAc, R3 = H

O O R1

O

R2

HO O

O AcO

O

OH OH

R1

R5

R2

R4

R4

41 R = R = H, R = OAc, R = R = O 42 R1 = R5 = H, R2 = OH, R3 = R4 = O 43 R1 = R4 = R5 = H, R2 = R3 = OAc 44 R1 = R4 = R5 = H, R2 = R3 = OH 45 R1 = H, R2 = OAc, R3 = R4 = O, R5 = OH 46 R1 = R3 = OH, R2 = OAc, R4 = R5 = H 93 R1 = R4 = R5 = H, R2 = OAc, R3 = OH 1

5

O

R3

O

2

Fig. 2 (continued)

3

4

OX3 O

R3

O

54 R = R = OH, R = H 55 R1 = X3, R2 = R3 = O 1

2

3

56

O

The Genus Walsura: A Rich Resource …

145 O O

O

O

O

O

O

O HO

O

HO

O

O

O

R

O

O

O

O OX3

OR

OR O

O

O

57 R = X3 58 R = X4

59 R = X3 60 R = X4

61 R = β -OH 62 R = α -OH O

O

O

OR

OR HO O

HO O

OH O

O

R1O

O

O O

O

63 R = X4 65 R = Me

64 R = X4

O

X3O

66 R1 = X5, R2 = H, R3 = Me 80 R1 = X4, R2 = H, R3 = Me 81 R1 = X4, R2 = R3 = H 82 R1 = X3, R2 = H, R3 = Me 83 R1 = X3, R2 = X4, R3 = H O

O

OR3

OR3

R2

O

OH

OR2 O

O

OH

OR3

OR2

O

R1

O

OR1

68 R1 = H, R2 = H, α -OH, R3 = Me 69 R1 = H, R2 = O, R3 = Me 70 R1 = R2 = H, α -OAc, R3 = Me 71 R1 = R3 = H, R2 = H, α -OAc 72 R1 = OH, R2 = H, α -OAc, R3 = Me 73 R1 = OH, R2 = H, α -OAc, R3 = H

67

O

74 R1 = H, R2 = OAc, R3 = Me 75 R1 = Ac, R2 = H, R3 = Me 76 R1 = R2 = H, R3 = Ac

O

O

NH OH

HO

O

HO

O O R1

R2

O

O OH

77 R1 = α -OH, R2 = α -OH 78 R1 = β -OH, R2 = α -OH 79 R1 = β -OH, R2 = O

Fig. 2 (continued)

84

O O

O OH

92

146

N. T. Son O

O

O

R6

OAc

AcO

HO

R1 R2

HO O

O OR5

O

O O

OX1

O

R 3 R4

96 R1 + R2 = Δ1,2, R3 + R4 = O, R5 = R6 = H 98 R1 = R2 = R5 = R6 = H, R3 + R4 = O 99 R1 + R2 = Δ1,2, R3 + R4 = O, R5 = H, R6 = α -OAc 100 R1 = R2 = R3 = R4 = R5 = R6 = H 101 R1 = R2 = R3 = R4 = R6 = H, R5 = Ac

97

102

O

O

O X3O

X3O

O

HO

HO O

HO O O

OH O

O

OAc

O

O OH

OH

OH

O

O

105

104

106 O

O

O

O

HO

O

HO

OH OAc

O

OH

O

OH

107

111

O

O

O

OAc

O

OAc

O

OH

OH

112

114

113

O

Fig. 2 (continued)

OH

OH

AcO

O

X =

O

O O

1

O

O O

HO

O OH

110 O

O

O

OH

O

O

OH

O

O 2

X =

3

X =

O 4

X =

O 5

X =

The Genus Walsura: A Rich Resource …

147

(13), a 5-oxatricyclo[5.4.0.11,4 ]hendecane in walrobsin B (39), a neotecleanin in walrobsin D (41), the wasurin type in walsurin B (98), and a 18(13→14)-abeo structure in walsuronoid G (108). The new structural and stereochemical variations of Walsura limonoids are due to differing patterns of hydroxylation, methoxylation, acetoxylation, and oxidation. Walsura robusta is a renowned multi-purpose herbal plant, and its various parts contain structurally diverse limonoids. A phytochemical analysis conducted by Zhang et al. revealed that, in its ethanol-H2 O (95:5, v/v) extract, most components were of the limonoid type [24]. Among 35 isolated compounds, 14 constituents were previously undescribed natural products, including the five cedrelone limonoids, 12α-acetoxycedrelone (1), 1α-ethoxy-11βhydroxydihydrocedrelone (26), 1α-methoxy-12α-acetoxydihydrocedrelone (34), 1α-methoxy-11β-hydroxydihydrocedrelone (35), and 11-oxo-dihydrocedrelone (36), and the five walsurin limonoids, walsurins A–E (97–101), in addition to four new rare 18(13→14)-abeo limonoids, walsuronoids F–I (107– 110) [24]. Phytochemical investigations of the ethanol extract of W. yunnanensis bark led to the isolation and NMR structural determination of eight new limonoids, namely, 11β-acetoxydihydrocedrelone (2), 11β-acetoxywalsuranolide (3), 20,22-dihydro-22,23-epoxywalsuranolide (21), 11β-hydroxycedrelone (27), 11β-hydroxydihydrocedrelone (28), isowalsurinolide (33), walsuranolide (94), and walsurin (96), in addition to the known analog cedrelone (5) [18]. In a search for 11β-hydroxysteroid dehydrogenase type-1 (11β-HSD1) inhibitors from plants, the ethanol (95%) extract of W. cochinchinensis twigs and leaves were reported to contain ten new limonoids, consisting of the vilasinin limonoids, cochinchinoids A–D (6–9), the three seco-ring D limonoids, cochinchinoids E–G (10– 12), and three azadirone-type ring-intact limonoids, cochinchinoids H–J (13–15) [15]. 7-Deacetoxy-7-hydroxyazadirone (16), dihydrocedrelone (17), 1,2-dihydro-7deacetoxy-7-hydroxyazadirone (18), 11β,12α-diacetoxycedrelone (20), and dysoxylumosin B–G (22–25) can be found in some other genera, but this is the first time these limonoids were observed in the genus Walsura [22, 24]. Cedrelone-type compounds seem to be the chemically predominant class among Walsura limonoids. For example, a phytochemical study on the ethanol-H2 O (95:5, v/v) extract of W. yunnanensis twigs and leaves resulted in the isolation of nine new cedrelone limonoids: 1α,11β-dihydroxy-1,2-dihydroisowalsuranolide (19), 11β-hydroxy-1α-methoxy-1,2-dihydroisowalsuranolide (29), 11β-hydroxy1,2-dihydroisowalsuranolide (30), 11β-hydroxy-23-O-methylwalsuranolide (31), 11β-hydroxyisowalsuranolide (32), walsuranolide B (95), yunnanol (111), and yunnanolides A (112) and B (113) [17]. To date, piscidofuran (36) has been the only new limonoid isolated from the fruits of W. piscidia [22]. Based on HPLC/DAD and HPLC purification work, eighteen undescribed A/B spiro-type limonoids, including seven new neotecleanin limonoids, walrobsins C–I (40–46), nine novel limonoids each bearing a 5-oxatricyclo[5.4.0.11,4]hendecane ring system, walrobsins A (38) and B (39) and J–P (47–53), and two key precursors, walrobsin Q (54) and R (55), together with the known analog walsuranin B (93), were isolated from the ethanol-H2 O (95:5, v/v) extract of W. robusta root bark [13, 14].

148

N. T. Son

Walsogyne A (56), a seco-ring C limonoid possessing a tetrahydrofuran2-ol ring, was separated and characterized structurally from the chloroform fraction of the ethanol extract of W. chrysogyne bark [24]. Moreover, the new dodecahydronaphtho[1,8-bc:5‚4-b ,c ]difuran-type limonoids, walsogynes B– E (57–60), and the dodecahydro-1H-naphtho[1,8-bc:3,4-c ]difuran-type limonoids, walsogynes F (61) and G (62) were also obtained from the bark of this species [19]. 14β,15β-Epoxidized limonoids also accumulate in Walsura species. However, the 14α,15α-epoxide ring has been found to date only in walsucochinones A–C (63–65), which are three new limonoids detected in Vietnamese W. cochinchinensis bark [6]. Another study demonstrated that the 95% ethanol extract of the twigs and leaves of the perennial Vietnamese plant W. cochinchinensis contains the novel limonoids, walsucochinoids A–Q (66–83) [16]. This group of metabolites is unlike normal limonoids, as they share a common feature of a five-membered ring fused to a sixmembered aromatic ring (Fig. 2). Walsura robusta is a further source of cedrelone-type limonoids. Utilizing HPLC, nine new 11β-HSD1 inhibitors with cedrelone backbones, namely, walsunoids A–I (84–92), were isolated and characterized structurally from an ethanol-soluble extract of W. robusta leaves [7]. Significantly, a maleimide substituent represents an unusual feature of 92. Walsuronoid A (102), a limonoid containing an unprecedented 3,4peroxide-bridged seco-A ring scaffold, the two rare 18(13→14)-abeo limonoids walsuronoids B (103) and C (104), and two new vilasinin limonoids walsuronoids D (105) and E (106), were also found to be characteristic of W. robusta [5, 14, 23]. Finally, a recent study described the isolation from a methanol extract of W. trichostemon stems of a new natural product, 114, which is also of the limonoid type [20].

2.2 Triterpenoids Plants from the genus Walsura also accumulate a large array of triterpenoids. A detailed list of 72 compounds (115–187) is summarized in Table 1 and Fig. 3. Nine plants (W. chrysogyne, W. cochinchinensis, W. pinnata, W. piscidia, W. robusta, W. trichostemon, W. trifoliata, W. tubulata, and W. yunnanensis) were found to contain this class of natural product. Walsura triterpenoids exist as several structural types, but those bearing the dammarane, tirucallane, and apotirucallane skeletons predominate. 7α-Acetoxy-17α-(20S),21,24-epoxy-apotirucall-14en-3-one-(23R,24S),25-triol (115), betulin (118), betulinic acid (119), grandifolinolenenone (137), hispidol B (138), and spicatin (172) are five known triterpenoids that were detected in W. robusta fruits [24], whereas the known triterpenoids dihydroniloticin (128), (22S,23R)-epoxytirucalla-7-ene-3α,24,25-triol (134), and niloticin (143) shown to be present in its leaves [7]. Apowalsogynes A (116) and B (117), two new 3,4-seco-apotirucallane triterpenoids, were precipitated from the methanol extract of W. chrysogyne bark [30].

The Genus Walsura: A Rich Resource …

149

OH HO O

OH O

OH

OH

R O

R2

OAc R1

O OAc

O

OAc

118 R1 = OH, R2 = CH2OH 119 R1 = OH, R2 = COOH

116 R = α -OMe 117 R = β -OMe

115

142 R1 = OH, R2 = Me 151 R1 = O, R2 = COOH OH

OH

OH O

OH

O

O

O

O HO

O

R

122

121

120 R = OH 133 R = O

OH O

OR

OH

HO

OH HO

OH OH

HO

123 R = H 147 R = Me

O

HO

OH

124 O

OH

125 OH

AcO

O

HO

HO

HO

O

R

O

OAc OH

126

128 R = OH 143 R = O

OH

127

OH O ROOC

R1

O ROOC

O

R2

HO

129 R = H 135 R = Me

130 R = H 148 R = Me

Fig. 3 Triterpenoids from the genus Walsura

131 R1 = R2 = OH 132 R1 = OAc, R2 = OH 146 R1 = OH, R2 = O

150

N. T. Son OH O

OH

O OH HO

134

137

136 OH HO

OH

O COOH

OH

HO

O

HO

OAc

O

O

HO

138

140

139 O

O O

OH OH

OH

O

O

O

HO

144

141

145 OH

OH

O

O

OH O O

O

O

150

149

152

OH

OH

O O

O

153

Fig. 3 (continued)

OH

O

O HO

154

155

The Genus Walsura: A Rich Resource …

151 OH

OH

O O

O

O

O

HO

O

AcO

OH HO

O

R

157 R = O 158 R = β -OH

O

O

156

159

OH O

O

AcO O

R

OH

O

OH

O OH

162

O

O

R3

OH

HO

R1

O

RO

R2

169 R = X4 (see Fig. 2, bottom) 170 R = X3

168 OH

HO O

HO

OAc O

OH

HO

OAc

AcO

O

HO

OH

O

OAc

O

OH

OAc

O

O

O

OH

O

164 R1 = OH, R2 = R3 = O 165 R1 = OH, R2 = R3 = α -OH 166 R1 = OX3, R2 = α -OH, R3 = O 167 R1 = H, R2 = R3 = α -OH

171 O

173

172

OAc

OH O

OH

OH

OH

AcO O

AcO

RO O

O

OAc

O

175 R = Ac 176 R = H

174

HO

Xn

177

OR 178 R = H

X6 = OH

OH

163

O

O

AcO

O

182 R = Ac 183

X7 = OH

O

O

184

X8 = O

O X9 =

Fig. 3 (continued)

OH

HO

HO

OH

160 R = α -OH 161 R = H

O

O

HO

O

OH

OH

O AcO

OH

O

O

185

R

179 R = α -OH 180 R = O

OH

152

N. T. Son OH

OH O

O

OH

OH

AcO O

AcO

HO

X3O

181

OH

186

O

AcO

X3O

OH

187

Fig. 3 (continued)

A new dammarane (20R,24S)-epoxy-25-hydroxydammaran-3-one (133), in addition to the eight known triterpenoids cabraleadiol (120), chrysura (121), cycloart23-ene-3β,25-diol (123), dymalol (129), eichlerianic acid (130), foveolin (135), 25-methoxycycloart-23-ene-3β-ol (147), and methyl eichlerianate (148), were also found in a methanol extract of W. chrysogyne leaves [33, 34]. Two new tirucallane triterpenoids, cochinchinoid K (122) and 3-epimesendanin S (131), and the known mesendanin S (145), were present in the 95% ethanol extract of W. cochinchinensis twigs and leaves [15]. In addition, walsucochins A (179) and B (180) with an unusual skeleton containing a phenylacetylene unit fused to a five-membered C-ring were also isolated from an ethanol (95%) extract of W. cochinchinensis twigs and leaves [41]. The new cycloartane, (24S,25R)-cycloartane-3β,24,25,26-tetrol (124), was the only triterpenoid found in W. yunnanensis twigs and leaves [17], whereas in a phytochemical study of the leaves of W. tubulata, walsurenol (181) was the only new triterpenoid identified [26]. Chromatographic separation of an n-hexane extract of W. pinnata stem bark resulted in the isolation of seven triterpenoids, including betulonic acid (119), 12αhydroxy-3-oxooleanano-28,13-lactone (139), 3β-hydroxy-5-glutinen-28-oic acid (140), 3-ketours-11-en-13β(28)-olide (141), oleanonic acid (149), 3-oxo-olean11-en-28,13β-olide (152), and 3-oxoursolic acid (154) [49]. W. pinnata bark also provided four triterpenoids 3-oxo-20(24)-epoxy-12β,25-dihydroxydammarane (150), 3-oxo-lup-20(29)-en-28-oic acid (151), 3-oxo-olean-9(11),12-dien-28-oic acid (153), and pinnatane A (155) [31, 32, 40]. Among them, compound 153 was new in the literature. Walsura trichostemon has been used as a traditional Thai medicinal plant to treat tendon disabilities [37]. As part of a search for bioactive compounds, three new cytotoxic apotirucallane triterpenoids, namely, 7-deacetylbrujavanone E (125), 11,25-dideacetyltrichostemonate (126), and 21,24,25-triacetyl-7-deacetyl-6hydroxylbrujavanone E (173) were isolated from an acetone extract of its leaves [37]. With a similar aim of searching for cytotoxic compounds, a new tirucallane triterpenoid, trichostemonoate (175), and the three known metabolites, 11α,20dihydroxydammar-24-en-3-one (127), grandifolinolenenone (137), and sapelin E acetate (171) were detected in an ethyl acetate-soluble extract of W. trichostemon stem bark [10]. This same species is characterized also by the presence of other

The Genus Walsura: A Rich Resource …

153

triterpenoids, in which friedelanone (136), melianone (144), and trichostemonol (176) were isolated from the twigs [38], 3-epimesendanin S 12 acetate (132), mesendanin G (146), and trichostemonate (174) were obtained from the roots [2, 36], and 12β,(20S,24R)-trihydroxydammar-25-en-3-one (177) and 12β,(20S),25trihydroxydammar-23-en-3-one (178) were purified from the stems [20]. Compounds 132, 174, and 176 were new natural products when first isolated. The two new compound groups, 182–185 and 185 and 186, which display dammarane and apotirucallane skeletons, respectively, are triterpenoids only recently observed in W. trichostemon [20]. The fruit pulp of Walsura piscidia is documented as being used in India to poison fish [3, 22]. An early phytochemical report by Purushothaman et al. described a number of triterpenoids from the leaves of this species. In particular, isolated were the two new tirucallane triterpenoids, piscidinols A (157) and B (158), and three new apotirucallanes, piscidinols C–E (159–161) [22]. In addition, two new metabolites, piscidinols F (162) and G (163) and the new piscidenone (156) were isolated also from W. piscidia leaves [3, 27]. W. trifoliata is a synonym of W. piscidia [35]. In an integrated study on the phytochemistry and biological properties of Meliaceae species and their constituents, the five new apotirucallane triterpenoids piscidinols H– L (164–168), were isolated and structurally characterized from a 95% ethanol extract of W. trifoliata leaves [35]. Also, two new apotirucallane triterpenoids (piscidinones A (169) and B (170)) were obtained from W. trifoliata leaves [11].

2.3 Sesquiterpenoids, Sterols, and Lignans Phytochemical studies on Walsura species have led to reports also of the presence of sesquiterpenoids, sterols, and lignans. Regarding sesquiterpenoids, altogether seven compounds (188–194) are summarized in Table 1 and Fig. 4. These phytochemicals were found in W. chrysogyne, W. pinnata and W. robusta [4, 12, 21, 32, 34, 39]. Apart from limonoids and triterpenoids, viridiflorol (194) was classified as a sesquiterpenoid present in W. chrysogyne leaves [34]. In turn, 2(3),6(7)-diepoxy-9humulene (188) was the only sesquiterpenoid isolated from W. pinnata bark, whereas ledol (189) and oplopanone (191) were metabolites identified in its stem bark [31, 32, 39]. Using a combination of chromatographic separation and NMR structural elucidation, two new sesquiterpenoids, 10β-nitro-isodauc-3-en-15-al (190) and 10-oxoisodauc-3-en-15-al (192), were purified and characterized from a methanol-soluble extract of W. robusta leaves [4]. It should be noted that nitro-sesquiterpenoids like 190 are very rare in both the genus Walsura and the family Meliaceae, or indeed in Nature. Therefore, it can be considered a good candidate as a chemotaxonomic marker. Turpinionoside A (193) was afforded from a methanol extract of W. robusta leaves and twigs [12]. The common chemical compound β-sitosterol (195) is a constituent of W. trichostemon twigs and roots, while its glucoside 196 can be found in W. pinnata

154

N. T. Son O

R

OH

OH

O

O

189

188

OH

O

190 R = β -NO2 192 R = C=O

191

OH

OH GlcO HO

RO

193

194

195 R = H 196 R = Glc

197

O

O

O O

HO

OH

OGlc

HO O

O

O

O

OGlc

HO

RO

OH

OH

O O

O

HO

198 R = Me 199 R = H

200

O

O

OH

O

O

O

202

HO

O

OH O

HO HO

203

O OH

201

O

O

O

O

O OH

O

OH

HO

HO

O

O OH

204

O

OH HO

OH

205

Fig. 4 Sesquiterpenoids, sterols, and lignans from the genus Walsura

bark and W. trichostemon roots [2, 31, 32, 38]. Another sterol, stigmasterol (197), was identified as one of metabolites in W. pinnata bark [31, 32]. Naturally occurring lignans are also distributed quite widely among Walsura species. To date, eight compounds of this type (198–205) have been isolated (Table 1 and Fig. 4), and they originate mainly from W. cochinchinensis, W. robusta, and W. yunnanensis [5, 6, 8, 12]. Thus, W. cochinchinensis bark produces the two known lignans, lariciresinol (200) and phyllanthin (203) [6]. Ji et al. isolated brassilignan (198) and 9-deoxyarctigenin (199) from an ethanol (95%) extract of W. robusta leaves [5]. (+)-Lyoniresinol 3α-O-β-d-glucopyranoside (201) and (–)-lyoniresinol 3α-Oβ-d-glucopyranoside (202) are two isomeric natural products that were separated by means of a chromatographic procedure from the methanol extract of W. robusta

The Genus Walsura: A Rich Resource …

155

leaves and twigs [12]. [(1R,2S,3S)-1,2,3,4-Tetrahydro-7-hydroxy-1-(4-hydroxy3-methoxyphenyl)-3-(hydroxymethyl)-6-methoxynaphthalen-2-yl]methyl-α-larabinopyranoside (204) and [(1S,2R,3R)-1,2,3,4-tetrahydro-7-hydroxy-1-(4hydroxy-3,5-dimethoxyphenyl)-3-(hydroxymethyl)-6-methoxynaphthalen-2yl]methyl-α-l-arabinopyranoside (205) are two new lignan glycosides, which were purified and characterized from the n-butanol extract of W. yunnanensis bark [8].

2.4 Simple Phenols, Flavonoids, Xanthones, Anthraquinones, and Other Types of Compounds Phytochemical work on Walsura species has revealed that several simple phenols may be present. Eight such compounds (206–213) are shown in Table 1 and Fig. 5. A phytochemical study of the Vietnamese W. cochinchinensis bark yielded four common phenolic compounds, including ferulaldehyde (206), methyl vanillate (207), protocatechuic acid (208), sinapaldehyde (209), and vanillic acid (212) [6]. Besides lignans, phytochemical investigation of the n-butanol extract of W. yunnanensis bark afforded the known phenolic 3,4,5-trimethoxyphenyl-β-dglucopyranoside (210), and the two new derivatives 3,4,5-trimethoxyphenyl 2-O(α-l-fucopyranosyl)-β-d-glucopyranoside (211) and walsuraside (214) [8]. Although flavonoids are abundant in the plant kingdom they seem to be rare among members of the genus Walsura (Fig. 5). A highly oxygenated flavonoid, namely, 3,5,6,7,8,3 ,4 -heptamethoxyflavone (214) was isolated from W. cochinchinensis bark collected in Vietnam [6]. Two new isomeric flavonols, 215 and 216, were characterized from the bark and leaves of W. trifoliata [45, 47]. Anthraquinone and xanthone derivatives occasionally occur within species of the genus Walsura. Ramana et al. isolated the new anthraquinone glycoside 4,8,10trihydroxy-10-(2-hydroxyacetoxy)-9-oxo-9,10-dihydroanthracene-2-carboxylic acid (217) and the known chrysophanol (218) from a methanol extract of W. trifoliata bark [45]. Two known xanthones, mangostenone F (219) and α-mangostin (220), were found in W. trichostemon twigs, with the latter compound (220) detected also in its bark [10, 38]. Phytochemical work has revealed also the presence of some additional structural types of secondary metabolites from Walsura species, which comprise the previously known amide asperglaucide (221) and the derivative, butyl α-d-fructofuranoside (222), in W. yunnanensis bark, as well as the aromatic derivative, 4-hydroxy-4,8dimethyl-1-tetralone (223), in W. pinnata bark [8, 31, 32].

156

N. T. Son R

O

HO

O

O

O O

RO

O

O

OH

O

O

HO

OH

HO

O HO

206 R = H 209 R = OMe

OH OH

208 R = H 212 R = Me

207

210

O O

O

O O O

OH

O

O

O

O HO

OH HO

O

O

OH

HO O

OH

O

O

OH

O

O

O

OH

OH

O

OH

O

OH

211

213

214

OH OH OH OH

O

OH

O

O

OH

COOH O

O

OH GlcO

R

OH OH

O

OH

215 R =  -OH 216 R = -OH

218

217

O OH

OH

O

O

O

O

N H

O

O HO

OH

O

O O

O

OH

220

219

H N

221 HO

OH HO O HO

O

O

OH

222

223

Fig. 5 Walsura simple phenols, flavonoids, anthraquinones, xanthones, and other types of compounds

The Genus Walsura: A Rich Resource …

157

3 Biological Activities 3.1 Cytotoxic Activity Cancer is a significant global health problem and is the second leading cause of death [46, 50]. More than 60% of anticancer drugs are derived from natural sources [51], and consequently the evaluation of bioactive agents from medicinal plants is a useful approach for their further discovery. Cytotoxic studies represent the principal biological focus of Walsura research and the results are outlined in Table 2. Walsura limonoids and triterpenoids were the main types of compounds tested in these cytotoxicity studies. Research conducted by Zhang and colleagues demonstrated that the IC 50 values of Walsura robusta limonoids against the growth of doxorubicin-resistant MCF-7 breast adenocarcinoma cells were in the following order: compound 97 (IC 50 0.52 μM) < 5 (1.04 μM) < 2 (1.60 μM) < 26 (1.86 μM) < 28 (2.06 μM) < 35 (2.23 μM) < 96 (2.65 μM) < positive control verapamil (3.75 μM) < 107 (4.36 μM) < 25 (4.42 μM) [24]. For 11β-hydroxycedrelone (27), an IC 50 value of 8.9 μM against HL-60 human myeloid leukemia cells was established [25]. Furan units are a possible reason for the difference in cytotoxicity noted between compounds 32 and 122, as compared to the cytotoxicity results of 11β-hydroxyisowalsuranolide (32) (IC 50 2.2–9.4 μM). This compound and yunnanolide A (112) exhibited IC 50 values ranging from 2.4 to 5.0 μM against the HL-60, SMMC-7721, A-549, MCF-7, SW-480 human cancer cell lines, and were compared in potency with their growth inhibitory effects on BEAS-2B normal human bronchial epithelial cells [17]. The novel seco-C limonoid walsogyne A (56) showed cytotoxicity toward P388 murine leukemia cancer cells with an IC 50 value of 5.0 μg/cm3 [21]. Among the limonoids, walsogynes B–G (57–62), compounds 58 and 62 were the most potent and both exhibited cytotoxicity to HL-60 human leukemia cells (Table 2) [19]. Walsuronoid B (103) induced G2/M phase arrest and mitochondrial and lysosomal apoptosis via the ROS/p53 signaling pathway in HepG2 and Bel-7402 cells (Fig. 6) [44]. Two new limonoids, walsuronoids D (105) and E (106), showed cytotoxic effects against the five human cancer cell lines, HL-60, SMMC-7721, A-549, MCF7, and SW-480, with IC 50 values of less than 5.0 μM, but compounds 124, 198, and 199 were regarded as being inactive (IC 50 > 40 μM) [5]. The different α- and β-orientations of the methoxy group present in compounds 116 and 117 resulted in differential cytotoxicity profiles being observed. Regarding HL-60 cell inhibition, the new 3,4-seco-apotirucallane triterpenoid apowalsogyne B (117) was more potent than its analog, apowalsogyne A (116), but 116 was superior to 117 in terms of HepG2, A-549, and MCF-7 cell growth inhibition (Table 2) [30]. The n-hexane extract of W. pinnata stem bark seems to possess promising cytotoxic activity since it inhibited the HepG2, MCF-7, HSC2, and CaSki cancer cell lines with IC 50 values of 3.0–7.0 μg/cm3 , whereas compounds 119 and 140 displayed IC 50 values of 12.0–19.0 and 18.0–25.0 μg/cm3 , respectively [39]. In another model,

158

N. T. Son

Table 2 Biological activities of Walsura isolated compounds and crude Walsura plant extracts No.

Model

Effects

Refs.

2

In vitro

IC 50 = 1.60 μM/MCF-7

[24]

5

In vitro

IC 50 = 1.04 μM/MCF-7

[24]

25

In vitro

IC 50 = 4.42 μM/MCF-7

[24]

26

In vitro

IC 50 = 1.86 μM/MCF-7

[24]

27

In vitro

IC 50 = 8.9 μM/HL-60

[25]

28

In vitro

IC 50 = 2.06 μM/MCF-7

[24]

32

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50 IC 50

35

In vitro

IC 50 = 2.23 μM/MCF-7

[24]

56

In vitro

IC 50 = 5.0

μg/cm3 /P388

[21]

57

In vitro

IC 50 = 27.7 μM/HL-60 IC 50 > 50 μM/HepG2, A-549, MCF-7

[19]

58

In vitro

IC 50 IC 50 IC 50 IC 50

= 7.7 μM/HL-60 = 37.7 μM/HepG2 = 29.9 μM/A-549 > 50 μM/MCF-7

[19]

59

In vitro

IC 50 > 50 μM/HL-60 and A-549 IC 50 = 21.7 μM/HepG2 IC 50 = 42.4 μM/MCF-7

[19]

60

In vitro

IC 50 > 50 μM/HL-60, HepG2, A-549, MCF-7

[19]

61

In vitro

IC 50 > 50 μM/HL-60, HepG2, A-549, MCF-7

[19]

62

In vitro

IC 50 IC 50 IC 50 IC 50

= 7.8 μM/HL-60 = 26.6 μM/HepG2 > 50 μM/A-549 = 18.2 μM/MCF-7

[19]

63

In vitro

IC 50 = 76.2 μg/cm3 /MCF-7

[6]

64

In vitro

IC 50 = 87.0 μg/cm3 /MCF-7

[6]

Cancer cell line cytotoxicity

= 3.6 μM/HL-60 = 2.4 μM/SMMC-7721 = 3.7 μM/A-549 = 4.2 μM/MCF-7 = 3.5 μM/SW-480 = 5.0 μM/BEAS-2B

μg/cm3 /MCF-7

[17]

65

In vitro

IC 50 = 8.1

96

In vitro

IC 50 = 2.65 μM/MCF-7

[24]

97

In vitro

IC 50 = 0.52 μM/MCF-7

[24]

[6]

(continued)

The Genus Walsura: A Rich Resource …

159

Table 2 (continued) No.

Model

Effects

Refs.

103

In vitro

G2/M phase arrest and mitochondrial and lysosomal apoptosis via the ROS/p53 signaling pathway in HepG2 and Bel-7402 cells

[44]

105

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50

= 2.7 μM/HL-60 = 3.1 μM/SMMC-7721 = 4.1 μM/A-549 = 3.1 μM/MCF-7 = 2.8 μM/SW-480

[5]

106

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50

= 3.3 μM/HL-60 = 4.1 μM/SMMC-7721 = 4.4 μM/A-549 = 4.4 μM/MCF-7 = 4.5 μM/SW-480

[5]

107

In vitro

IC 50 = 4.36 μM/MCF-7

112

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50 IC 50

= 3.1 μM/HL-60 = 2.2 μM/SMMC-7721 = 2.6 μM/A-549 = 3.9 μM/MCF-7 = 2.4 μM/SW-480 = 9.4 μM/BEAS-2B

[17]

116

In vitro

IC 50 IC 50 IC 50 IC 50

= 35.9 μM/HL-60 = 30.9 μM/HepG2 = 31.1 μM/A-549 = 32.2 μM/MCF-7

[30]

117

In vitro

IC 50 IC 50 IC 50 IC 50

= 26.9 μM/HL-60 = 68.0 μM/HepG2 = > 50 μM/ A-549 = 62.5 μM/MCF-7

[30]

119

In vitro

IC 50 IC 50 IC 50 IC 50

= 19.0 μM/HepG2 = 12.0 μM/MCF-7 = 13.0 μM/HSC-2 = 14.0 μM/CaSki

[39]

In vitro

IC 50 = 8.85 μg/cm3 /MCF-7 IC 50 = 14.35 μg/cm3 /SK-OV-3 IC 50 = 27.23 μg/cm3 /MRC-5

[24]

[32]

(continued)

160

N. T. Son

Table 2 (continued) No.

Model

Effects

Refs.

119

In vitro/in vivo

Apoptosis generated in leukemia stem cells (LSC) via BAX upregulation, with the suppression of the Bcl-2 and survivin genes, leading caspase 9 activation. Also induced apoptosis in LSC xenotransplanted zebrafish

[29]

124

In vitro

IC 50 > 40 μM/HL-60, SMMC-7721, A-549, MCF-7, SW-480

[5]

125

In vitro

IC 50 = 12.92 μg/cm3 /KB

[37]

126

In vitro

IC 50 = 3.95 IC 50 = 12.99 μg/cm3 /HeLa

127

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50

136

In vitro

IC 50 > 30 μg/cm3 /KB and HeLa

[38]

137

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50

= 41.2 μg/cm3 /KB = 24.1 μg/cm3 /HeLa = 13.6 μg/cm3 /COLO-205 = 20.4 μg/cm3 /HepG2 = 22.5 μg/cm3 /LLC

[10]

140

In vitro

IC 50 = 18.28 μg/cm3 /MCF-7 IC 50 = 14.95 μg/cm3 /SK-OV-3 IC 50 = 20.09 μg/cm3 /MRC-5

In vitro

IC 50 IC 50 IC 50 IC 50

In vitro

IC 50 = 4.5 μg/cm3 /KB IC 50 = 18.5 μg/cm3 /HeLa

144

μg/cm3 /KB

[37]

= 2.0 μg/cm3 /KB = 1.9 μg/cm3 /HeLa = 3.7 μg/cm3 /COLO-205 = 17.4 μg/cm3 /HepG2 = 3.2 μg/cm3 /LLC

[10]

= 18.0 μM/HepG2 = 25.0 μM/MCF-7 = 21.0 μM/HSC2 = 20.0 μM/CaSki

[32]

[39]

[38] (continued)

The Genus Walsura: A Rich Resource …

161

Table 2 (continued) No.

Model

Effects

Refs.

155

In vitro

IC 50 = 48.8 μM/MRC-5 IC 50 = 60.9 μM/MCF-7 IC 50 = 92.9 μM/MDA-MB-231 IC 50 = 33.9 μM/EJ-28 IC 50 = 48.0 μM/RT-112 IC 50 = 59.7 μM/HeLa S3 IC 50 = 19.0 μM/Hep3B IC 50 = 55.8 μM/HepG2 IC 50 = 50.9 μM/A-549 IC 50 = 87.2 μM/DU-145 IC 50 = 55.3 μM/PC-3 IC 50 > 100 μM/SiHa and SK-LU-1

[40]

169

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50 IC 50 IC 50 IC 50

= 34.23 μg/cm3 /HT-29 = 17.77 μg/cm3 /MCF-7 = 25.17 μg/cm3 /HeLa = 17.94 μg/cm3 /A-549 = 27.78 μg/cm3 /B-16 = 16.37 μg/cm3 /IEC-6 = 21.22 μg/cm3 /L6 = 13.62 μg/cm3 /PC-3

[11]

170

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50 IC 50 IC 50 IC 50

= 50.63 μg/cm3 /HT-29 = 24.62 μg/cm3 /MCF-7 = 27.74 μg/cm3 /HeLa = 18.48 μg/cm3 /A-549 = 46.08 μg/cm3 /B-16 = 18.52 μg/cm3 /IEC-6 = 13.52 μg/cm3 /L6 = 14.10 μg/cm3 /PC-3

[11]

171

In vitro

IC 50 = 22.3 μg/cm3 /KB IC 50 = 18.9 μg/cm3 /HeLa IC 50 = 64.0 μg/cm3 /COLO-205 IC 50 > 100 μg/cm3 /HepG2 and LLC

[10]

173

In vitro

IC 50 = 17.06 μg/cm3 /KB

[37]

μg/cm3 /KB

174

In vitro

IC 50 = 3.28 IC 50 = 0.93 μg/cm3 /HeLa

[36]

175

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50

= 4.4 μg/cm3 /KB = 3.8 μg/cm3 /HeLa = 5.3 μg/cm3 /COLO-205 = 16.2 μg/cm3 /HepG2 = 5.5 μg/cm3 /LLC

[10]

176

In vitro

IC 50 = 12.4 μg/cm3 /KB IC 50 = 25.1 μg/cm3 /HeLa

195

In vitro

IC 50 > 30 μg/cm3 /KB and HeLa

[38] [38] (continued)

162

N. T. Son

Table 2 (continued) No.

Model

Effects

Refs.

198

In vitro

IC 50 > 40 μM/HL-60, SMMC-7721, A-549, MCF-7, SW-480

[5]

199

In vitro

IC 50 > 40 μM/HL-60, SMMC-7721, A-549, MCF-7, SW-480

[5]

200

In vitro

IC 50 = 69.0 μg/cm3 /MCF-7

[6]

206

In vitro

IC 50 = 13.8 μg/cm3 /MCF-7

[6]

209

In vitro

IC 50 = 37.4

μg/cm3 /MCF-7

[6]

218

In vitro

IC 50 = 48.5 μg/cm3 /KB IC 50 > 30.0 μg/cm3 /HeLa

219

In vitro

IC 50 IC 50 IC 50 IC 50 IC 50

= 13.5 μg/cm3 /KB = 19.7 μg/cm3 /HeLa = 6.6 μg/cm3 /COLO-205 = 19.5 μg/cm3 /HepG2 = 5.1 μg/cm3 /LLC

[10]

n-Hexane extract of W. pinnata stem bark

In vitro

IC 50 IC 50 IC 50 IC 50

= 5.0 μg/cm3 /HepG2 = 6.0 μg/cm3 /MCF-7 = 3.0 μg/cm3 /HSC2 = 7.0 μg/cm3 /CaSki

[49]

132

In vitro

MIC = 128 μg/cm3 /B. cereus and B. subtilis MIC > 128 μg/cm3 /S. aureus, E. coli, P. aeruginosa, and S. sonnei

[2]

131

In vitro

MIC = 16.0 μg/cm3 /B. cereus MIC = 16.0 μg/cm3 /B. subtilis MIC > 128.0 μg/cm3 /S. aureus, E. coli, P. aeruginosa, and S. sonnei

[2]

146

In vitro

MIC = 64.0 μg/cm3 /B. cereus and P. aeruginosa MIC = 128.0 μg/cm3 /E. coli MIC > 128.0 μg/cm3 /B. subtilis, S. aureus, and S. sonnei

[2]

Methanol extract of W. robusta leaves and twigs

In vitro

11 mm (inhibitory dry disk zone)/ [12] S. mutans GS-5 10 mm/S. mutans OMGS 2482 16 mm/S. pyogenes

n-Butanol extract of W. robusta leaves and twigs

In vitro

12 mm/S. mutans GS-5 10 mm/S. mutans OMGS 2482 17 mm/S. pyogenes

[38]

Antimicrobial activity

[12]

(continued)

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163

Table 2 (continued) No.

Model

Effects

Refs.

Aqueous extract of W. robusta leaves and twigs

In vitro

11 mm/S. mutans GS-5 10 mm/S. mutans OMGS 2482 16 mm/S. pyogenes

[12]

Ethanol (95%) extract of W. robusta wood

In vitro

MIC = 7.81 mg/cm3 /E. coli [49] O26:H11, E. coli O111:NM, and E. coli ATCC 25,922 MIC = 1.95 mg/cm3 /E. coli O157:H7 MIC = 15.62 mg/cm3 /E. coli O22

n-Hexane extract of W. trifoliata In vitro young leaves

MIC > 2.0 mg/cm3 /A. niger, A. flavus, C. albicans, Botrytis cinerea, Trichophyton rubrum, and T. mentagrophytes

n-Hexane extract of W. trifoliata In vitro mature leaves

MIC > 2.0 mg/cm3 /A. niger, A. [52] flavus, and B. cinerea MIC = 1.0 mg/cm3 /C. albicans, T. rubrum, and T. mentagrophytes

n-Hexane extract of W. trifoliata In vitro bark

MIC > 2.0 mg/cm3 /A. flavus, [52] MIC = 0.5–1.0 mg/cm3 /A. niger, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

n-Hexane extract of W. trifoliata In vitro roots

MIC = 0.5–1.0 mg/cm3 /A. niger, [52] A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

Ethyl acetate extract of W. trifoliata young leaves

In vitro

MIC = 0.5–1.0 mg/cm3 /A. niger, [52] A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

Ethyl acetate extract of W. trifoliata mature leaves

In vitro

MIC = 0.5–1.0 mg/cm3 /A. niger, [52] A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

Ethyl acetate extract of W. trifoliata bark

In vitro

MIC = 0.125–0.5 mg/cm3 /A. niger, A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

[52]

Ethyl acetate extract of W. trifoliata roots

In vitro

MIC = 0.125–0.5 mg/cm3 /A. niger, A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

[52]

[52]

Methanol extract of W. trifoliata In vitro young leaves

MIC = 0.5–1.0 mg/cm3 /A. niger, [52] A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

Methanol extract of W. trifoliata In vitro mature leaves

MIC = 0.25–0.5 mg/cm3 /A. niger, A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

[52]

(continued)

164

N. T. Son

Table 2 (continued) No.

Model

Effects

Refs.

Ethyl extract of W. trifoliata bark

In vitro

MIC = 0.125–1.0 mg/cm3 /A. niger, A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

[52]

Methanol extract of W. trifoliata In vitro roots

MIC = 0.031–0.125 mg/cm3 /A. niger, A. flavus, C. albicans, B. cinerea, T. rubrum, and T. mentagrophytes

[52]

Aqueous extract W. trifoliata stem bark

In vitro

7.2 mm (inhibitory zone)/E. coli 7.7 mm/K. pneumoniae 7.5 mm/B. subtilis 7.4 mm/S. aureus

[53]

Silver nanoparticles from W. trifoliata stem bark

In vitro

14.6 mm (inhibitory zone)/E. coli [53] 16.4 mm/K. pneumoniae 13.1 mm/B. subtilis 13.6 mm/S. aureus

13

In vitro

IC 50 = 11.44 μM/human 11β-HSD1

[15]

69

In vitro

IC 50 = 13.4 μM/mouse 11β-HSD1

[16]

71

In vitro

IC 50 = 8.25 μM/mouse 11β-HSD1

[26]

91

In vitro

IC 50 = 9.9 μM/human 11β-HSD1 Not active/mouse 11β-HSD1

[7]

122

In vitro

IC 50 = 3.20 μM/human 11β-HSD1 IC 50 = 0.82 μM/mouse 11β-HSD1

[15]

128

In vitro

IC 50 = 3.8 μM/mouse 11β-HSD1

[7]

134

In vitro

IC 50 = 1.9 μM/human 11β-HSD1 IC 50 = 1.2 μM/mouse 11β-HSD1

[7]

143

In vitro

IC 50 = 0.69 μM/mouse 11β-HSD1

[7]

145

In vitro

IC 50 = 3.74 μM/human 11β-HSD1 IC 50 = 1.15 μM/mouse 11β-HSD1

[15]

157

In vivo

IC 50 = 0.88 μM/mouse 11β-HSD1

[7]

Antidiabetic activity

(continued)

The Genus Walsura: A Rich Resource …

165

Table 2 (continued) No.

Effects

Refs.

n-Hexane extract of W. trifoliata In vivo

Model

IC 50 = 50.0 μg/cm3 /α-glucosidase inhibition

[42]

Ethyl acetate extract of W. trifoliata

IC 50 = 50.0 μg/cm3 /α-glucosidase inhibition

[42]

IC 50 = 690.1 μg/cm3 /α-glucosidase inhibition

[42]

In vivo

Methanol extract of W. trifoliata In vivo

Anti-inflammatory activity (NO production inhibition) 38

In vitro

IC 50 = 9.20 μg/cm3 /RAW264.7 μg/cm3 /RAW264.7

[13, 14]

39

In vitro

IC 50 = 52.76 IC 50 > 50 μg/cm3 /BV2 and THB-1

[14]

40

In vitro

IC 50 > 50 μg/cm3 /RAW264.7, BV2, and THB-1

[14]

41

In vitro

IC 50 = 41.15 μg/cm3 /RAW264.7 [14] IC 50 = 21.19 μg/cm3 /BV2 IC 50 > 50 μg/cm3 /THB-1

42

In vitro

IC 50 > 50 μg/cm3 /RAW264.7, BV2, and THB-1

47

In vitro

IC 50 = 28.29 μg/cm3 /RAW264.7 [14] IC 50 = 15.09 μg/cm3 /BV2 IC 50 = 28.5 μg/cm3 /THB-1

48

In vitro

IC 50 = 25.69 μg/cm3 /RAW264.7 [14] IC 50 = 22.25 μg/cm3 /BV2 IC 50 = 18.01 μg/cm3 /THB-1

49

In vitro

IC 50 > 50 μg/cm3 /RAW264.7, BV2, and THB-1

50

In vitro

IC 50 = 16.58 μg/cm3 /RAW264.7 [14] IC 50 = 20.36 μg/cm3 /BV2 IC 50 = 7.96 μg/cm3 /THB-1

51

In vitro

IC 50 > 50 μg/cm3 /RAW264.7, BV2, and THB-1

52

In vitro

IC 50 = 30.72 μg/cm3 /RAW264.7 [14] IC 50 = 20.05 μg/cm3 /THB-1

53

In vitro

IC 50 = 52.46 μg/cm3 /RAW264.7 [14] IC 50 > 50 μg/cm3 /BV2 IC 50 = 15.97 μg/cm3 /THB-1

102

In vitro

IC 50 > 50 μg/cm3 /RAW264.7, BV2, and THB-1

[14]

[14]

[14]

[14]

Antioxidative activity (continued)

166

N. T. Son

Table 2 (continued) No.

Model

Effects

178

In vitro

H2 O2 -induced PC12 cell damage [41] inhibition

Refs.

179

In vitro

H2 O2 -induced PC12 cell damage [41] inhibition

193

In vitro

IC 50 = 204.9 μM/DPPH radical scavenging IC 50 = 6.25 mM/OH• radical scavenging

[12]

201

In vitro

IC 50 = 68.7 μM/DPPH radical scavenging IC 50 = 0.8 mM/O2 −• radical scavenging

[12]

202

In vitro

IC 50 = 51.5 μM/DPPH radical scavenging IC 50 = 7.4 mM/OH• radical scavenging IC 50 = 0.7 mM/O2 −• radical scavenging

[12]

204

In vitro

IC 50 = 42.4 μg/cm3 /DPPH radical scavenging

[8]

205

In vitro

IC 50 = 48.8 μg/cm3 /DPPH radical scavenging

[8]

210

In vitro

IC 50 = 68.7 μM/DPPH radical scavenging

[12]

213

In vitro

IC 50 = 43.2 μg/cm3 /DPPH radical scavenging

[8]

Methanol extract of W. robusta leaves and twigs

In vitro

IC 50 = 40.0 μg/cm3 /DPPH radical scavenging IC 50 = 1.0 μg/cm3 /OH• radical scavenging IC 50 = 1.76 mg/cm3 /O2 −• radical scavenging

[12]

Diethyl ether extract of W. robusta leaves and twigs

In vitro

IC 50 = 540.0 μg/cm3 /DPPH radical scavenging

[12]

n-Butanol extract of W. robusta leaves and twigs

In vitro

IC 50 = 20.0 μg/cm3 /DPPH radical scavenging IC 50 = 4.0 μg/cm3 /OH• radical scavenging

[12]

Aqueous extract of W. robusta leaves and twigs

In vitro

IC 50 = 60.0 μg/cm3 /DPPH radical scavenging IC 50 = 2.0 μg/cm3 /OH• radical scavenging

[12]

Antifeedant activity (continued)

The Genus Walsura: A Rich Resource …

167

Table 2 (continued) No.

Model

Effects

157

In vivo

LC 50 = 19.15 μg/insect/A. janata [35] LC 50 = 26.60 μg/insect/S. litura

Refs.

165

In vivo

LC 50 = 41.08 μg/insect/A. janata [35] LC 50 = 41.35 μg/insect/S. litura

168

In vivo

LC 50 = 22.40 μg/insect/A. janata [35] LC 50 = 30.21 μg/insect/S. litura

Methanol extract of W. trifoliata In vivo

LC 50 = 37.69 μg/insect/A. janata [35] LC 50 = 43.32 μg/insect/S. litura

Ichthyotoxicity 130

In vivo

TL M 6.7 ppm/zebrafish

[34]

194

In vivo

TL M 15.0 ppm/zebrafish

[34]

In vivo

34.5%/sperm mobility count 2.50 million/mm3 /testes 26.5 million/mm3 /cauda epididymis

[43]

Methanol extract of W. piscidia In vivo leaves (400 mg/kg/21 days/Wistar rats)

32.4%/sperm mobility count 1.70 million/mm3 /testes 15.1 million/mm3 /cauda epididymis

[43]

Apoptotic inhibition of cerebral ischemia

[9]

Antifertility activity Methanol extract of W. piscidia leaves (200 mg/kg/21 days/ Wistar rats)

Neuroprotective effect Ethanol extract of W. piscidia In vivo leaves (400 mg/kg/21 days/Wistar rats)

Fig. 6 Apoptosis produced by walsuronoid B (103)

the two triterpenoids 119 and 140 also showed moderate cytotoxicity against SKOV-3 human ovarian adenocarcinoma cells (Table 2) [32]. Betulonic acid (119) was obtained from the bark of W. pinnata and triggered intrinsic apoptosis in leukemia stem cells via BAX upregulation, Bcl-2, and survivin gene supression, as well as

168

N. T. Son

caspase 9 and downstream caspase 3/7 activation. In addition, compound 119 was able to suppress leukemia formation in a zebrafish model [29]. Tirucallane and apotirucallane triterpenoids from W. trichostemon are considered potential cytotoxic agents, and various triterpenoids such as 7-deacetylbrujavanone E (125), 11,25-dideacetyltrichostemonate (126), melianone (144), 21,24,25-triacetyl7-deacetyl-6-hydroxybrujavanone E (173), and trichostemonate (174), from this plant source, exhibited cytotoxicity against KB human epidermoid carcinoma and/or HeLa cells [36–38]. The two triterpenoids, 11α,20-dihydroxydammar-24-ene-3-one (127), trichostemonoate (175), and the xanthone, mangostenone F (219), all showed antiproliferative effects against the KB, HeLa, COLO-205, and HepG2 human cancer cell lines, and against LLC Lewis lung murine carcinoma cells, with IC 50 values of less than 20.0 μg/cm3 . Against this same cancer cell line panel, grandifolinolenenone (137) had IC 50 values of 13.6–41.2 μg/cm3 , whereas sapelin E acetate (171) was active against KB, HeLa, and COLO-205 cells, but not for HepG2 and LLC cells [38]. The triterpenoid pinnatane A (155) exhibited cancer cell line cytotoxicity potency (IC 50 values) in the order: Hep3B hepatocellular carcinoma cells (19.0 μM) > EJ-28 bladder carcinoma cells (33.9 μM) > RT-112 bladder carcinoma cells (48.0 μM) > A-549 lung adenocarcinoma cells (50.9 μM) > PC-3 prostate adenocarcinoma cells (55.3 μM) > HepG2 hepatocellular carcinoma cells (55.8 μM) > HeLa S3 cervical adenocarcinoma cells (59.7 μM) > MCF-7 breast cancer cells (60.9 μM) > DU-145 prostate carcinoma cells (87.2 μM) > MDA-MB-231 breast adenocarcinoma cells (92.9 μM) > SiHa cervical carcinoma cells, and SK-LU-1 lung adenocarcinoma cells (>100 μM, inactive) [40]. The new apotirucallane triterpenoid, piscidinone A (169), was more active against the HT-29, MCF-7, and B-16 cancer cells than piscidinone B (170) [11] (Table 2). These results possibly can be explained by the effect of the presence of a C-11-affixed tigloyl unit in compound 169, compared with a 3methylbutanoate ester group in 170. Finally, while an IC 50 value of 69.0 μg/cm3 was reported for the lignan lariciresinol (200) against MCF-7 cells, two simple phenols from the same plant source (the bark of W. cochinchinesis), ferulaldehyde (206) and sinapaldehyde (209), exhibited, in turn, IC 50 values of 13.8 and 37.4 μg/cm3 [6].

3.2 Antimicrobial Activity Pathogenic organisms are harmful to healthy individuals and cause a variety of infectious diseases. The use of traditional antibiotics, which are mostly synthetically prepared, is associated with a long duration of treatment, high costs, and drug resistance [54]. As a result, the discovery of new antibiotic drugs derived from natural sources seems necessary. Three structurally related triterpenoids 131, 132, and 146, isolated from the roots of W. trichostemon, were evaluated against four bacterial strains. Of these, 3-epimesendanin S (131) showed moderate antibacterial activity against the Grampositive Bacillus cereus with a MIC value of 16.0 μg/cm3 , while its 12-acetate 132,

The Genus Walsura: A Rich Resource …

169

a new compound, was somewhat less active (MIC value of 64.0 μg/cm3 ). However, meliasenin G (146) displayed only weak activity, with a MIC value of 128.0 μg/cm3 [2]. Compounds 131 and 146 exerted the same MIC value of 64.0 μg/cm3 against B. subtilis and the Gram-negative Pseudomonas aeruginosa, but compound 132 was inactive (MIC > 128.0 μg/cm3 ) [2]. In contrast to the negative result of the weakly polar diethyl ether extract of W. robusta leaves and twigs, three more polar extracts of this plant, produced using n-butanol, methanol, and water sequentially, inhibited Streptococcus mutans GS-5, S. mutans OMGS 2482, and S. pyogenes growth with inhibitory zones of 10–17 mm [12]. An ethanol (95%) extract of W. robusta wood demonstrated poor MIC values of 1.95 mg/cm3 against Escherichia coli O157:H7, and 7.81 mg/cm3 for E. coli O26:H11, E. coli O111:NM, E. coli ATCC 25,922, and 15.62 mg/cm3 for E. coli O22 [49]. In an antimicrobial assay against Aspergillus niger, A. flavus, Candida albicans, Botrytis cinerea, Trycophyton rubrum, and T. mentagrophytes, n-hexane, ethyl acetate, and methanol extracts of W. trifoliata roots displayed MIC values of 0.031–1.0 mg/cm3 . These values were higher than those of extracts obtained from the young and mature leaves and from the bark of the same species [52]. In addition, a silver nanoparticle formulation of an aqueous extract of W. trifoliata stem bark improved on its antimicrobial potency, since the inhibitory zones were found to measure 13.1–14.6 mm for Bacillus subtilis, Escherichia coli, Klebsiella pneumoniae, and Staphylococcus aureus, which were approximately two times greater in size than those for the unprocessed extract [53].

3.3 Antidiabetic Activity 11β-Hydroxysteroid dehydrogenase type-1 (11β-HSD1) is an NADPH-dependent enzyme that converts cortisone to its active form, cortisol. The latter may then result in metabolic changes, such as insulin suppression and hyperglycemia [55]. Hence, 11βHSD1 resistance can provide a novel therapeutic approach for finding new agents to treat type-2 diabetes mellitus [55]. From Walsura species, limonoids and triterpenoids, once again, have been the main compound types to have subjected to 11βHSD1 inhibition assays. The IC 50 values related to the human 11β-HSD1 enzyme were reported for compounds 134 (IC 50 1.9 μM) > 122 (3.20 μM) > 145 (3.74 μM) > 91 (9.9 μM) > 13 (11.44 μM) [7, 25]. In turn, for the mouse 11β-HSD1 enzyme, the order of inhibitory activity found was 143 (IC 50 0.69 μM) > 122 (0.82 μM) > 157 (0.88 μM) > 145 (1.15 μM) > 134 (1.20 μM) > 128 (3.8 μM) > 71 (8.25 μM) > 69 (13.4 μM) > 91 (not active) [7, 15, 16]. Another widely used approach to antidiabetic drug discovery is to determine effects on α-glucosidase inhibition [56]. Using this type of assay, the n-hexane and ethyl acetate extracts of W. trifoliata roots displayed the same IC 50 value of 50 μg/cm3 , and were being more potent than either its methanol extract (IC 50 690.1 μg/cm3 ) or the positive control used, acarbose (IC 50 290.9 μg/cm3 ) [42].

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3.4 Anti-inflammatory Activity There have been several attempts to investigate Walsura constituents for their potential anti-inflammatory effects. In a lipopolysaccharide (LPS) stimulated RAW264.7 macrophage cell anti-inflammatory assay, the order of NO production inhibitory activity for several limonoids tested from W. robusta was 38 (IC 50 9.2 μg/cm3 ) > 50 (16.58 μg/cm3 ) > 48 (25.69 μg/cm3 ) > 47 (28.29 μg/cm3 ) > 52 (30.72 μg/cm3 ) > 41 (41.15 μg/cm3 ) > positive control N G -monomethyl-l-arginine monoacetate (48.15 μg/cm3 ) > 52 (52.46 μg/cm3 ) > 39 (52.76 μg/cm3 ) > 40, 42, 49, 51, and 102 (>50 μg/cm3 , inactive) [14]. Furthermore, the new limonoid, walrobsin A (38), attenuated iNOS and IL-1β gene expression, as well as iNOS and COX-2 protein levels [13]. Two compounds, namely, walrobsins J (47) and K (48) induced potential anti-inflammatory activities when LPS stimulated BV2 microglial cells and Propionibacterium acnes-stimulated THB-1 human monocytic cells were utilized [14].

3.5 Antioxidative Activity The two new triterpenoids, walsucochins A (178) and B (179), at doses of 1.0, 5.0, and 10.0 μM, promoted the cell viability of PC12 cells damaged by the strong radical oxidant, H2 O2 [41]. Bioassay-guided fractionation has been conducted on extracts of W. robusta leaves and twigs with potential antioxidative constituents. As a representative example, the methanol extract displayed IC 50 values of 40 μg/cm3 , 1.0 μg/cm3 , and 1.76 mg/cm3 in capturing the 1,1-diphenyl-2-picrylhydrazyl (DPPH), OH• , and O2 −• radicals, respectively [12]. In addition, its metabolites turpinionoside A (193), (+)-lyoniresinol 3α-O-β-d-glucopyranoside (201), (–)-lyoniresinol 3α-O-βd-glucopyranoside (202), and 3,4,5-trimethoxyphenyl-β-d-glucopyranoside (210) exhibited IC 50 values in a DPPH radical scavenging test procedure that were lower than that of a positive control, butylated hydroxytoluene (IC 50 90.8 μM). Furthermore, in an O2 −• assay, compounds 201 and 202, with IC 50 values 0.7–0.8 mM proved to be more potent than the positive control trolox (IC 50 3.36 mM) [12]. In another DPPH oxidation assessment on constituents of W. yunnansensis, the two new lignans 205 and 210 and the new phenol 213 displayed IC 50 values of 42.4, 48.8, and 43.2 μg/cm3 , respectively [8]. Besides antidiabetic activity, W. trifoliata also showed antioxidative properties. Thus, the n-hexane, ethyl acetate, and methanol extracts of its roots displayed DPPH, OH• , O2 −• , and NO radical scavenging as well as inhibition of lipid peroxidation and ferric reducing antioxidant power in the concentration range of 200–1000 μg/cm3 [42].

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3.6 Antifeedant and Other Activities Antifeedant activity is a well-known property of several types of secondary metabolites of plants of the family Meliaceae. A methanol extract containing several apotirucallane triterpenoids from W. trifoliata leaves was subjected to an antifeedant activity evaluation, from which piscidinol A (157), piscidinol I (165), and piscidinol L (168) caused mortality LC 50 values of 19–41 μg/insect (castor semilooper Achea janata) and 26–43 μg/insect (tobacco caterpillar Spodotera litura), as compared with those of the methanol extract of their plant origin (38 μg/insect A. janata, μg/insect S. litura) [35]. The triterpenoid eichlerianic acid (130) and the sesquiterpenoid viridiflorol (194) were found to be active in an ichthyotoxic assay against zebrafish with a median tolerance limit (TL M ) of 6.7 and 15 ppm, respectively [34]. A methanol extract of W. piscidia leaves, at doses of 200 and 400 mg/kg, showed antifertility effects in rats, since it caused a reduction of sperm mobility, testes, and cauda epididymis during 21 days of treatment [43]. An ethanol extract of W. piscidia leads at a high dose of 400 mg/kg to reduced apoptosis caused by cerebral ischemia in Wistar rats, following a 21-day treatment [9].

4 Synthesis Aspects Due in part to their novel structural features and interesting biological values, certain Walsura constituents have been of interest in synthesis chemistry. The novel triterpenoid (±)-walsucochin B (180) may be used to illustrate a recent representative example. Its total synthesis started from farnesyl bromide (224) via several key steps, such as titanocene-mediated radical cyclization, cycloaromatization, and a Cu-mediated remote C–H hydroxylation [57]. In detail, farnesyl bromide (224) was alkylated with (cyanomethyl)cuprate to afford nitrile 225 in 68% yield (Fig. 7). Compound 225 was converted into epoxide 226 with a 60% yield with N-bromosuccinimide (NBS), via the epoxidation of the terminal alkene. Based on a titanocene-mediated epoxide opening radical cascade reaction, intermediate 226 was transformed into ketone 227 (52% yield) in two steps. Imine 228 was first formed by treatment of ketone 227 with 2-picolylamine and catalytic amounts of p-toluenesulfonic acid monohydrate (PTSA.H2 O). Then, Cu(NO3 )2 .3H2 O (2.0 equiv.) and H2 O2 (5.0 equiv.), under optimized conditions, were added to form the tricyclic ketone 229 from compound 228. It was found that the use of t-BuOK, THF, and dithioacetal substrate at 23°C for 18 h converted 229 into intermediate 230. When heated at 60°C for 2 h, this provided phenol 231 in 68% yield. Methyl ether 232 was obtained in 82% yield when 231 was treated with MeI. The C(sp2 )-SMe methylation of the thioether 232 with MeMgBr and using NiCl2 (PPh3 )(IPr) as a catalyst, resulted in 233 in 86% yield. Compound 234

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NBS, THF/H2O, MeOH, K2CO3

68%

60%

CN

CN Cp2TiCl2, THF, Zn0, r.t. TSSCI, DMF, TBSO imidazole, r.t., 16 h, 52% (two steps)

O

224

225

O

226

227

PTSA. H2O, toluene, reflux, 14 h

O

N NH2

S O

S

O TBSO

S

O

t-BuOK, THF, TBSO then 18, r.t., 18 h

OH

230

Cu(NO3)2.3H2O, H2O2 satd. aq. Na4EDTA

OH

N N TBSO

229

228

(69% single diastereomer) then 60oC 2 h, 68%

S

S

MeI, K2CO3, acetone, 82%

TBSO

O

O

OH

86%

TBSO

OH

OH

232

231

NiCl2(PPh3)(IPr),TBSO MeMgBr, toluene, 30°C, 24 h

OH

233 NIS, HFIP 98% r.t., 2 h

TMS

TMS I

O

TBSO

O

DMP, NaHCO3, DCM, r.t., 2 h 96%

O

TBSO

236

OH

235

PdCl2(PPh3)2, CuI, Et3N, 50oC, 24 h, 86%

O

TMS TBSO

OH

234

TBAF

O

O

+ HO

O

O

OH

180 ((±)-walsucochin B)

Fig. 7 Synthesis of walsucochin B (180)

was formed via regioselective iodination using N-iodosuccinimide (NIS) in hexafluoroisopropanol (HFIP). In the next step, direct Sonogashira coupling (TMSCCH, CuI, PdCl2 (PPh3 )4 , Et3 N) installed the final requisite trimethylsilylethynyl group in the secondary alcohol 235 with an excellent yield of 86%. Dess-Martin periodinane (DMP) oxidation of compound 235 to ketone 236, and finally desilylation with tetrabutylammonium fluoride (TBAF) enabled the total synthesis of (±)-walsucochin B (180).

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5 Conclusions The current contribution provides a systematic and exhaustive overview of the phytochemistry and biological activities of the secondary metabolites of plants in the genus Walsura (family Meliaceae). Nine species, comprising W. chrysogyne, W. cochinchinensis, W. pinnata, W. piscidia, W. robusta, W. trichostemon, W. trifoliata, W. tubulata, and W. yunnanensis are the main subjects of prior phytochemical investigations. Chemical constituents derived from Walsura species are characterized by the presence of various skeletons, such as anthraquinones, flavonoids, lignans, sterols, and xanthones, but limonoids and triterpenoids predominate. A total of 223 secondary metabolites have been summarized, in which limonoids (114 compounds) and triterpenoids (72 compounds) accounted for ~51 and ~32%, respectively. Significantly, more than 100 Walsura limonoids were isolated from Nature for the first time. Walsura plant constituents are being studied biologically mostly for the purposes of drug discovery. Various in vitro biological and in vivo pharmacological tests have been performed on both naturally occurring isolated compounds and plant extracts from this genus mainly focused on their cytotoxic, antimicrobial, antidiabetes, antiinflammatory, antioxidant, antifeedant, antifertility, ichthyotoxic, and neuroprotective activities. To gain greater insight into the activities of these compounds from Walsura species, more intensive investigations on their cellular mechanisms of action are necessary. Acknowledgments The writing of this chapter was supported by Vietnam Academy of Science and Technology (2021).

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Ninh The Son obtained his Ph.D. degree from Tokushima Bunri University, Tokushima, Japan, with support from the Japanese Society for the Promotion of Science (JSPS) in 2019. He worked at the Institute of Chemistry, Vietnam Academy of Science and Technology, as a chemistry researcher, and, in 2020, was a postdoctoral scientist at the University of Sao Paulo, School of Pharmaceutical Sciences, Brazil. His main research involves natural products chemistry and molecular simulation. Dr. Son received an excellence award in 2020 from VAST. He played a significant role in several projects and is a co-author of 80 scientific articles, among which he served as corresponding author of more than 50, and he has published six articles as a single author. He has also edited a book and served as a corresponding author of three book chapters. He is also active as a reviewer for various scientific publications, such as “Natural Product Research” and “Studies in Natural Products Chemistry”.