Progress in the Chemistry of Organic Natural Products 114 3030594432, 9783030594435

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Progress in the Chemistry of Organic Natural Products

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

114 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 College of Pharmacy, Columbus, OH, USA Heinz Falk , Institute of Organic Chemistry, University Linz, Linz, Austria Simon Gibbons , School of Pharmacy, University of East Anglia, Norwich, UK Jun’ichi Kobayashi, Graduate School of Pharmaceutical Sciences, Hokkaido University, Sapporo, Japan Yoshinori Asakawa , Faculty of Pharmaceutical Sciences, Tokushima Bunri University, Tokushima, Japan Ji-Kai Liu , School of Pharmaceutical Sciences, South-Central University for Nationalities, Wuhan, China 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 Verena M. Dirsch , Department of Pharmacognosy, University of Vienna, Vienna, Austria Agnieszka Ludwiczuk , Department of Pharmacognosy, Medical University of Lublin, Lublin, Poland 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 at Greensboro, 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 http://www.springer.com/series/10169

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

Progress in the Chemistry of Organic Natural Products Volume 114

With contributions by Hucheng Zhu · Chunmei Chen · Qingyi Tong · Yuan Zhou · Ying Ye · Lianghu Gu · Yonghui Zhang Nwet Nwet Win · Hiroyuki Morita Julio C. Pardo-Novoa · Carlos M. Cerda-García-Rojas Yingjie Bai · Liyun Zhang · Xiaoguang Lei

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

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

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

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

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

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

ISSN 2191-7043 ISSN 2192-4309 (electronic) Progress in the Chemistry of Organic Natural Products ISBN 978-3-030-59443-5 ISBN 978-3-030-59444-2 (eBook) https://doi.org/10.1007/978-3-030-59444-2 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 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

Progress in the Chemistry of Cytochalasans . . . . . . . . . . . . . . . . . . . . . . . . . . Hucheng Zhu, Chunmei Chen, Qingyi Tong, Yuan Zhou, Ying Ye, Lianghu Gu, and Yonghui Zhang

1

Bioactive Compounds from Medicinal Plants in Myanmar . . . . . . . . . . . . 135 Nwet Nwet Win and Hiroyuki Morita New Techniques of Structure Elucidation for Sesquiterpenes . . . . . . . . . . 253 Julio C. Pardo-Novoa and Carlos M. Cerda-García-Rojas Human Endogenous Natural Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 Yingjie Bai, Liyun Zhang, and Xiaoguang Lei

v

Progress in the Chemistry of Cytochalasans Hucheng Zhu, Chunmei Chen, Qingyi Tong, Yuan Zhou, Ying Ye, Lianghu Gu, and Yonghui Zhang

Contents 1 2

3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Structural Diversity of Cytochalasans and Their Origins . . . . . . . . . . . . . . . . . . . . . . 2.1 The Cytochalasin Group . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 The Pyrichalasin Group . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 The Chaetoglobosin Group . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 The Aspochalasin Group . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 The Alachalasin Group . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Trichalasin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological Activities of Cytochalasans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Cytotoxicity and Potential Anticancer Activities . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Antimicrobial Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 6 29 36 54 71 71 72 73 74 77

H. Zhu · C. Chen · Q. Tong · Y. Zhou · Y. Ye · L. Gu · Y. Zhang (B) Hubei Key Laboratory of Natural Medicinal Chemistry and Resource Evaluation, School of Pharmacy, Huazhong University of Science and Technology, Hangkong Road 13#, Wuhan 430030, People’s Republic of China e-mail: [email protected] H. Zhu e-mail: [email protected] C. Chen e-mail: [email protected] Q. Tong e-mail: [email protected] Y. Zhou e-mail: [email protected] Y. Ye e-mail: [email protected] L. Gu e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 114, https://doi.org/10.1007/978-3-030-59444-2_1

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H. Zhu et al.

3.3 Antiparasitic Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Anti-inflammatory Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Antiviral Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Phytotoxic Effects and Ecological Role . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7 Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Biosynthesis of Cytochalasans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Cytochalasan Gene Clusters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Investigations into the Biosynthesis of Cytochalasans . . . . . . . . . . . . . . . . . . . . . . 4.3 Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Chemical Syntheses of Cytochalasans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Synthesis of Periconiasin G . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Syntheses of Cytochalasin B and L-696,474 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Total Syntheses of Periconiasins A–E . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Total Synthesis of Aspochalasins D and B and (+)-Aspergillin PZ . . . . . . . . . . . 5.5 Total Synthesis of Asperchalasine A and Related Derivatives . . . . . . . . . . . . . . . 5.6 Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

78 79 79 80 81 82 83 86 97 98 101 101 105 108 111 115 117 118

1 Introduction Cytochalasans are a group of fungal-derived natural products characterized by the presence of a perhydro-isoindolone core fused with a macrocyclic ring, with these compounds exhibiting high structural diversity and a broad-spectrum of bioactivities. From the standpoint of biosynthesis, cytochalasans are generated via a hybrid polyketide synthase–non-ribosomal peptide synthetase (PKS–NRPS) biosynthesis pathway with certain amino acids. Cytochalasans have attracted significant attention from the chemical and pharmacological scientific communities and have already been reviewed in recent years from different vantage points, such as their chemistry and biology [1], biosynthesis [2], and total synthesis [3]. There is continued interest in cytochalasans and relevant investigations on these compounds are growing rapidly, as noted from the fact that only 100 cytochalasans were reported by 2009 [1], but by 2020, this number had grown to 500. The present contribution provides a general view of the isolation, structural determination, biological activities, biosynthesis, and total synthesis of cytochalasans. In all, 477 cytochalasans are described: some “unnatural” cytochalasans obtained by feeding or genetic manipulation and a new sub-group named “merocytochalasans” are also included. Merocytochalasans are a class of cytochalasans arising from the dimerization or polymerization of one or more cytochalasan molecules with one or more other natural product units. The term “merocytochalasan” was put forward initially in 2017 by Zhang and coworkers [4], but the first isolation and characterization of merocytochalasans may be traced back to the isolation of aspochalamins A–D (439–442) from Aspergillus niveus LU 9575 in 2004 [5, 6]. Merocytochalasans are found mainly as a group of aspochalasins, generally with an additional epicoccine moiety or peptide (amino acid). Due to their intriguing structures and potential

Progress in the Chemistry of Cytochalasans

3

bioactivities, merocytochalasans have attracted considerable interest from organic and biosynthetic chemists as well as pharmacologists. In the process of completing this work, it was noted that there is some confusion in the assignment of trivial names of certain cytochalasans, since either the same compound may have different names or else different compounds have been assigned the same name. Therefore, synonyms have been merged in this chapter, such as “cytochalasin D (2) (zygosporin A)”. In turn, for the homonyms, these have denoted by adding the family name of the corresponding author after the trivial name, such as “cytochalasin N Natori (14)” and “cytochalasin N Edwards (20).” There are also some cytochalasans reported with neither a trivial name nor a chemical name in the original literature, so their original bolded code numbers with the relevant literature information have been used to identify them, such as “compound 1 (68) Bioorg. Med. Chem. Lett. 2015, 25, 1823” [7] and “compound 15ii (80) Org. Lett. 2019, 21, 4163” [8]. A conscientious effort has been made to cover cytochalasans as comprehensively as possible in this chapter. Although not all relevant literature reports may have been included, it is hoped that this contribution will provide a good understanding of cytochalasans, and also may arouse additional wide scientific interest in their further investigation.

2 The Structural Diversity of Cytochalasans and Their Origins Based on the different amino acids involved in their construction, cytochalasans have been classified generally into five groups [1], including “cytochalasins” (with a phenylalanine residue, Sect. 2.1), “pyrichalasins” (with a tyrosine or a related derivative residue, Sect. 2.2), “chaetoglobosins” (with a tryptophan residue, Sect. 2.3), “aspochalasins” (with a leucine residue, Sect. 2.4), and “alachalasins” (with an alanine residue, Sect. 2.5). In 2012, a new type of cytochalasan named “trichalasin” (with a valine residue, Sect. 2.6) was reported by Zou and coworkers from Trichoderma gamsii, but this trichalasin group still has only one member, trichalasin A (477) [9]. Thus, there are altogether six groups of cytochalasans in terms of the different amino acids incorporated into the polyketide skeleton (Fig. 1). In this contribution, 477 cytochalasans have been included: comprising 163 cytochalasins, 46 pyrichalasins, 133 chaetoglobosins, 127 aspochalasins, seven alachalasins, and one trichalasin (Table 1). For each group of cytochalasans, they can be divided further into several sub-classes based mainly on the different types of macrocyclic ring fused to the isoindolone moiety, such as a carbocyclic ring (usually with odd-numbered carbons), a lactone ring, a carbonate ring, or a seco-ring. Merocytochalasans are listed separately as a sub-class under the groups of cytochalasins, chaetoglobosins, and aspochalasins. Besides changes occurring on ring C, additional modifications of

4

H. Zhu et al. 12

12

11 5

10

7

3

9

HN

1 O 23

OO

12

11

OH

5

10 15

13

3

HN

O

17

1

7 13

O OAc

21

5

10

15

9 21

O

11

OH

3

19

HN

N H

17

OH

1

OO

7 9 23

15

13 17

21

19

19

O

O

group 1: cytochalasin A (109)

group 2: pyrichalasin H (164)

OH

group 3: chaetoglobosin A (210)

OH B A

HN O O

C

O

HN

group 4: aspochalasin A (343)

HN O O HO

O

O O

O

O

OH

group 6: trichalasin A (477)

group 5: alachalasin A (470)

Fig. 1 Representatives of six groups of cytochalasans Table 1 Statistical data of six groups of cytochalasans (477 in total) Group

Sub-classes

Number

Subtotal

Compound number

Cytochalasins

Carbocyclic ring C

108

163

1–108

Lactone ring C Cyclic carbonate ring C Seco-ring C Pyrichalasins

Chaetoglobosins

Aspochalasins

Trichalasin

109–144

4

145–148

10

149–158

Mero-cytochalasins

5

159–163

Carbocyclic ring C

31

46

164–194

Lactone ring C

7

195–201

Cyclic carbonate ring C

7

202–208

Seco-ring C

1

209

Carbocyclic ring C

116

133

210–325

Seco-ring C

8

326–333

Mero-chaetoglobosins

9

334–342

Carbocyclic ring C

70

Lactone ring C

21

413–433

5

434–438

Seco-ring C Alachalasins

36

Mero-aspochalasins

31

Carbocyclic ring C

5

Mero-alachalasins

2

Lactone

1

127

343–412

439–469 7

470–474 475–476

1

477

Progress in the Chemistry of Cytochalasans

5

the amino acid residue and reconstruction of the isoindolone moiety may also further increase the structural diversity of the cytochalasans. As mentioned in the Introduction, almost all the reported cytochalasans have been isolated from fungi, mainly inclusive of the genera Chaetomium, Aspergillus, Phoma, Phomopsis, Xylaria, Spicaria, Periconia, Daldinia, and Hypoxylon. While the first cytochalasans were isolated from species from the genera Phoma and Helminthosporium, fungi of these genera are not the main sources of cytochalasans. The most important cytochalasan-producing fungi are members of the genera Chaetomium and Aspergillus, and nearly 200 cytochalasans, and about 40% of the cytochalasans reported to date have been isolated from these two genera. Some fungi were found to produce cytochalasans belonging to more than one structural group, such as Chaetomium globosum (Plate 1) and Aspergillus flavipes (Plate 2). These normally produce chaetoglobosins and aspochalasins, respectively, but were also found to produce pyrichalasins [10] and cytochalasins [11]. It may be noted that all seven of the alachalasins were isolated from Stachybotrys chartarum in 2008 [12], with no further analogs of this type reported subsequently. As the only member of the trichalasin group, the generation of trichalasin A (477) is perhaps questionable [9], because it was co-isolated with a number of aspochalasins from the fungus Trichoderma gamsii. Fungi are not the only source of cytochalasans, as there are also a few exceptions reported: chaetoglobosin A (210) was also been isolated from a Streptomyces sp. [13] and six oxidized derivatives of L-696,474 (26) were obtained via bioconversion using an Actinoplanes sp. [14]. Plate 1 Colonies of Chaetomium globosum

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H. Zhu et al.

Plate 2 Colonies of Aspergillus flavipes

2.1 The Cytochalasin Group Cytochalasins are a group of cytochalasans with a phenylalanine unit incorporated into the polyketide backbone (Tables 2, 3, 4, 5, 6 and 7 and Figs. 2, 3, 4, 5, 6 and 7), which is also the largest group of cytochalasans (163 out of 477 representatives). The first cytochalasin was isolated from a culture broth of a Phoma sp. (strain S-298) in 1966, which was named “phomin” [15]. Almost simultaneously, in 1967, the structurally related compounds, cytochalasins A (109) and B (110), were isolated from Helminthosporium dematioideum [16], and in the same investigation, cytochalasins C (1) and D (2) were reported from Metarrhizium anisopliae. This is the first time “cytochalasin” was used for this new class of natural products from fungi, with this name being proposed as descriptive of their biological effects (κÚτoς, kytos = cell, χαλασις, chalasis = relaxation) [16]. In fact, the structure of cytochalasin B (110) is identical to that of phomin, and the structure of cytochalasin A (109) is identical to that of dehydrophomin [15]. From then on, the name “cytochalasin” has been used widely, rather than “phomin”. To date, 163 cytochalasins with various skeleton have been reported, including carbocyclic cytochalasins, lactone, and carbonated cytochalasins, seco-cytochalasins, and mero-cytochalasins.

Progress in the Chemistry of Cytochalasans

2.1.1

7

Carbocyclic Cytochalasins

[5.6.11]-Cytochalasins and Related Derivatives This group, with 101 members, refer to cytochalasins with an 11-membered carbocyclic ring that is fused to the isoindolone moiety, leading to the [5.6.11] ring system as well as their derivatives with [5.6.6.7], [5.6.5.8], [5.6.6.6], or other skeletons (Table 2 and Fig. 2). Cytochalasins C (1) and D (2) are the first cytochalasins of

HN

HN

HN O OAc

HO

O

1 (cytochalasin C)

O OAc

O

HO 2 (cytochalasin D)(zygosporin A)

OAc

O HO 4 (dytochalasin D monoacetate) (zygosporin F)

5 (zygosporin E)

O OAc HO 6 (zygosporin G)

OH

OH

O OAc

OH

O

O OAc

HO

7 (cytochalasin H)

O

HN

HN O OAc

HO

10 (cytochalasin J) (deacetylcytochalasin H)

HO

13 (engleromycin)

O OH

HO

11 (epoxycytochalasin H)

OH

HN

HN O

HO

12 (epoxycytochalasin J) (epoxydeacetylcytochalasin H)

OH

OH

O

HO

9 (kodo-cytochalasin-2)

O

HN

O OH

O OH

HO

8 (kodo-cytochalasin-1)

OH

O

HN

HN

HN

O OH

O HO 3 (desacetylcytochalasin D) (zygosporin D)

HN

HN O OAc

O OAc

O OH

OH

HN

HN

OH

OH

OH

O OAc

HO

14 (cytochalasin N Natori)

Fig. 2 Structures of [5.6.11]-cytochalasins and related derivatives

O OH

HO

15 (cytochalasin O Natori)

8

H. Zhu et al. OH OH

OH OH

OH OH

HN

HN

HN O OAc

O OH

HO

16 (cytochalasin P Natori)

OH

O

OH OH

OH

HN O OH

HN O OAc

HO

HO

O OAc

O

20 (cytochalasin N Edwards)

19 (cytochalasin S)

OH OH

HO

18 (cytochalasin R Natori)

17 (cytochalasin Q Natori)

OH

HN

O OH

HO

HO

O

21 (cytochalasin O Edwards)

O

O O

HN

HN O OAc

HO 22 (cytochalasin P Edwards)

HN O OAc

O

HO

O OAc

O

23 (cytochalasin Q Edwards)

HO

O

24 (cytochalasin R Edwards)

HO OH

HN

HN

HN O OAc

O OAc

O OAc

HO

27 (L-696,475)

26 (L-696,474)

25 (cytochalasin U Chen) HO

OH

OH OH

HN

HN

O OAc

O OAc

28 (L-697,318)

30 (16ß,22-dihydroxy-L-696,474)

O

O HN O OAc

HO

O

31 (19,20-epoxycytochalasin Q Edwards)

Fig. 2 (continued)

OH

O O OH

OH

O OAc

29 (22-hydroxy-L-696,474)

O

HN

OH HN

HN HO

O

32 (21-O-deacetyl-19,20epoxycytochalasin Q Edwards)

O O

HO

33 ((7S*,13E,16S*,18S*,19E)-7,18dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13,19-triene1,21-dione)

Progress in the Chemistry of Cytochalasans

9 OH

OH

O O

O O

HO

34 ((7S*,13E,16S*,18R*)-7,18dihydroxy-16,18-dimethyl-10phenyl-[11]-cytochalasa-6(12),13diene-1,21-dione)

O O

35 ((7S*,13E,16S*,18S*,19R*)-7,18,19trihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13-diene-1,21dione)

HO

36 ((7S*,13E,16S*,18S*,19R*)-7,18dihydroxy-16,18-dimethyl-19-methoxy10-phenyl-[11]cytochalasa-6(12),13diene-1,21-dione)

OH

OAc O O

HN

HO

OH

HN

O

OH

HN

HN

OH

OH

HN

HN O O

HO

37 ((7S*,13E,16S*,18S*,19R*)-7,18dihydroxy-16,18-dimethyl-19-acetoxy10-phenyl-[11]-cytochalasa-6(12),13diene-1,21-dione)

O O

38 ((7S*,13E,16S*,18R*,19E)-16,18dimethyl-7-hydroxy-10-phenyl-[11]cytochalasa-6(12),13,19-triene-1,21dione)

39 ((7S*,13E,16S*,18S*)-16,18dimethyl-7-hydroxy-10-phenyl [11]-cytochalasa-6(12),13-diene1,21-dione)

OH OH

OH

HN

HN

O O

HN O O

40 ((7S*,13E,16S*,18R*,19R*)-7,19dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13-diene1,21-dione)

O O

HO

41 ((6Z,13E,16S*,18S*,19E)-18hydroxy-16,18-dimethyl-10-phenyl [11]-cytochalasa-6,13,19-triene1,21-dione)

O

O

HN

O O

OH

43 ((6Z,13E,16S*,17R*,18S*,19E)-17hydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6,13,19-triene-1,21dione)

OH

HN

HN O O

HO

42 ((6E,13E,16S*,18S*,19E)-12,18dihydroxy-16,18-dimethyl-10phenyl-[11]-cytochalasa-6,13,19triene-1,21-dione)

O O

HO

44 ((7S*,13E,16S*,18S*,19E)-6,7epoxy-18-hydroxy-16,18-dimethyl10-phenyl-[11]cytocalasa-13,19diene-1,21-dione)

OH

HO

45 ((7S*,13E,16S*,18S*,19R*)-6,7epoxy-18,19-dihydroxy-16,18dimethyl-10-phenyl-[11]cytochalasa-13-ene-1,21-dione)

O

OOH

O HN

HN O O HO

46 ((7S*,13E,16S*,18R*,19E)-7,18dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13,19-triene1,17,21-trione)

HN O O

O

O OAc O HO

OH

47 ((5Z,7S*,13E,16S*,17R*,18S*,19E)-7hydroperoxy-17-hydroxy-16,18-dimethyl10-phenyl-[11]-cytochalasa-5,13-19triene-1,21-dione)

48 (19,20-epoxycytochalasin R Edwards)

O

O

O

O

OH

O HN

HN

HN O OAc O

O

49 (18-deoxy-19,20-epoxycytochalasin R Edwards)

Fig. 2 (continued)

O OAc O

O

50 (18-deoxy-19,20-epoxycytochalasin Q Edwards)

O OAc O HO

O

51 (19,20-epoxycytochalasin N Edwards)

10

H. Zhu et al. OH

OH

HN

HN

HN O OAc O HO

O OAc O HO

O

52 (19,20-epoxycytochalasin C)

O OAc

O

53 (19,20-epoxycytochalasin D)

O

OH

OH

HN

HN O OH

HO

HO

54 (RKS-1778)

HN O OH HO

O

O O

O

56 ((6R,16S,18R,21R)-18,21dihydroxy-16,18-dimethyl-10phenyl[11]cytochalasa-(13E,19E)diene-1,7,17-trione)

55 (deacetylcytochalasin C)

HO

57 (fragiformin A)

OH O

OH

O

HN

HN

O O HO

59 ((16S*,18S*,19R*)-12,18-dihydroxy-19-methoxy-16,18-dimethyl 10-phenyl-[11]-cyto-chalasa6(7),(13E)-diene-1,21-dione)

58 (fragiformin B)

OH

60 ((6R*,7S*,16S*,18S*,19R*)-6,7epoxy-18-hydroxy-19-methoxy-16,18dimethyl-10-phenyl-[11]-cytochalasa(13E)-ene-1,21-dione)

OH

OH

O

OH

HN

HN

HN O OH

O OAc

HO

61 (cytochalasin Z10)

O OAc O O HO 63 (19,20-epoxicitocalasina)

HO

62 (cytochalasin Z11)

O

O

OH

HN

HN

HN O OAc

OO

O OH

O

64 (18-deoxycytochalasin Q Edwards)

OH

O

67 (18-methoxy cytochalasin J) (cytochalasin J1)

Fig. 2 (continued)

O 66 (cytochalasin H2)

OH

O

HN

HN

HN

OH

O

65 (21-O-deacetylcytochalasin Q Edwards)

OH

O OH

O

HN

O O HO

O O HO

O OH

O

68 (compound 1)(Bioorg Med Chem Lett (2015) 25:1823)

O OH

O

69 (compound 2)(Bioorg Med Chem Lett (2015) 25:1823)

Progress in the Chemistry of Cytochalasans

11 OH

OH

HN

HN O OH

70 (compound 3)(Bioorg Med Chem Lett (2015) 25:1823)

HN O OAc O

O

HO

O OAc

O

O

72 (18-desoxycytochalasin C)

71 (18-desoxy-19,20-epoxycytochalasin C)

COOH

COOH

O

HN

HN O OAc

O OAc

HO

73 (phomocytochalasin)

HN O OH

HO

HO 75 (phomopsichalasin E)

74 (phomopsichalasin D)

OOH

OH

HN

OH

HN O OAc HO

76 (phomopchalasin C)

HN O OH

O OAc

77 (cytochalasin J2)

78 (cytochalasin H1)

O

O

OH OH OH HN

HN

HN O OAc

O OAc

79 (fragiformin E)

HO

80 (compound 15ii)(Org Lett (2019) 21:4163

O O O AcO HO HO

81 (cytochalasin D1) OH OH

O OH OH

HN

OH

O O O AcO HO HO

HN

HN O OAc

82 (cytochalasin C1)

HO

O OAc

O

83 (6,12-epoxycytochalasin D)

O

HO

84 (6-epi-cytochalasin P) HO

OH OAc

O

O

HN

HN O OAc

HO

O

85 (7-O-acetylcytochalasin P

Fig. 2 (continued)

HN O OAc

O

86 (7-oxo-cytochalasin C)

O OAc

OH

87 (12-hydroxycytochalasin Q Edwards)

O

12

H. Zhu et al.

O

O

OH HN

HN

O

HN

O OH OH

O OH

88 (phomopchalasin B)

89 (cytochalasin J3)

O O

90 (xylastriasan A)

OH O

OH OH HN

HN O OH

O OAc

91 (phomopchalasin A)

92 (cytochalasin H2)

OH O N H O

O

O

O

O N

HO

93 (chaetoconvosin A)

OH O

O

O N H O

O

HO

O O

94 (chaetoconvosin B)

95 (chamiside A)

H N

O

OH

O

OH O

HN

HN O OH HO

OH

O

96 (curtachalasin A) H N

O OH HO O

97 (curtachalasin B) H N

O

O HO O

HO OH

Fig. 2 (continued)

98 (curtachalasin C)

O

OH HO HO O

99 (curtachalasin D)

O

100 (curtachalasin E)

O

Cl

OAc

Cl

HN O HO O

OH

101 (xylarichalasin A)

Progress in the Chemistry of Cytochalasans

13

Table 2 [5.6.11]-Cytochalasins and related derivatives Compound

Origin

References

Cytochalasin C (1)

Metarrhizium anisopliae

[16] [22]

Cytochalasin D (2) (zygosporin A)

Metarrhizium anisopliae Zygosporium masonii

[16] [22] [17] [18] [19] [20] [21]

Desacetylcytochalasin D (3) (zygosporin D)

Zygosporium masonii

[23]

Cytochalasin D monoacetate (4) (zygosporin F)

Zygosporium masonii

[23]

Zygosporin E (5)

Zygosporium masonii

[23]

Zygosporin G (6)

Zygosporium masonii

[23]

Cytochalasin H (7)

Phomopsis sp. (ZZF08)

[24] [25] [26] [27]

Kodo-cytochalasin-1 (8)

Phomopsis paspalli

[28] [26]

Kodo-cytochalasin-2 (9)

Phomopsis paspalli

[28]

Cytochalasin J (10) Deacetylcytochalasin H

Phomopsis sojae Phomopsis sp. (68-GO-1641)

[29] [30]

Epoxycytochalasin H (11)

Phomopsis sojae

[29] [30]

Epoxycytochalasin J (12) Epoxydeacetylcytochalasin H

Phomopsis sojae

[29] [30]

Engleromycin (13)

Englermyces goetzei

[31]

Cytochalasin N Natori (14)

Phomopsis sp. (68-GO-1641)

[30] [32]

Cytochalasin O Natori (15)

Phomopsis sp. (68-GO-1641)

[30] [32]

Cytochalasin P Natori (16)

Phomopsis sp. (68-GO-1641)

[30] [32]

Cytochalasin Q Natori (17)

Phomopsis sp. (68-GO-1641)

[32]

Cytochalasin R Natori (18)

Phomopsis sp. (68-GO-1641)

[32]

Cytochalasin S (19)

Phomopsis sp. (68-GO-1641)

[32]

Cytochalasin N Edwards (20)

Hypoxylon terricola Mill.

[33]

Cytochalasin O Edwards (21)

Hypoxylon terricola Mill.

[33]

Cytochalasin P Edwards (22)

Hypoxylon terricola Mill.

[33] (continued)

14

H. Zhu et al.

Table 2 (continued) Compound

Origin

References

Cytochalasin Q Edwards (23)

Hypoxylon terricola Mill.

[33]

Cytochalasin R Edwards (24)

Hypoxylon terricola Mill.

[33]

Cytochalasin U Chen (25)

Pestalotia sp. AB1942R-114

[34]

L-696,474 (26)

Hypoxylon fragiforme (ATCC 20995, MF5511)

[35]

L-696,475 (27)

Hypoxylon fragiforme (ATCC 20995, MF5511)

[35]

L-697,318 (28)

Hypoxylon fragiforme (ATCC 20995, MF5511)

[35]

22-Hydroxy-L-696,474 (29)

Actinoplanes sp. ACTT 53771

[14]

16β,22-Dihydroxy-L-696,474 (30)

Actinoplanes sp. ACTT 53771

[14]

19,20-Epoxycytochalasin Q Edwards (31)

Xylaria obovata

[36]

21-O-Deacetyl-19,20epoxycytochalasin Q Edwards (32)

Xylaria obovata

[36]

(7S*,13E,16S*,18S*,19E)-7,18Daldinia sp. Dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13,19-triene1,21-dione (33)

[37]

(7S*,13E,16S*,18R*)-7,18Daldinia sp. Dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13-diene-1,21dione (34)

[37]

(7S*,13E,16S*,18S*,19R*)-7,18,19Trihydroxy-16,18-dimethyl-10phenyl-[11]-cytochalasa-6(12),13diene-1,21-dione (35)

Daldinia sp.

[37]

(7S*,13E,16S*,18S*,19R*)-7,18Dihydroxy-16,18-dimethyl-19methoxy-10-phenyl-[11]-cytochalasa6(12),13-diene-1,21-dione (36)

Daldinia sp.

[37]

(7S*,13E,16S*,18S*,19R*)-7,18Dihydroxy-16,18-dimethyl-19acetoxy-10-phenyl-[11]-cytochalasa6(12),13-diene-1,21-dione (37)

Daldinia sp.

[37]

(continued)

Progress in the Chemistry of Cytochalasans

15

Table 2 (continued) Compound

Origin

References

(7S*,13E,16S*,18R*,19E)-16,18Dimethyl-7-hydroxy-10-phenyl-[11]cytochalasa-6(12),13,19-triene-1,21dione (38)

Daldinia sp.

[38]

(7S*,13E,16S*,18S*)-16,18Dimethyl-7-hydroxy-10-phenyl [11]-cytochalasa-6(12),13-diene-1,21dione (39)

Daldinia sp.

[38]

(7S*,13E,16S*,18R*,19R*)-7,19Daldinia sp. Dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13-diene-1,21dione (40)

[38] [39]

(6Z,13E,16S*,18S*,19E)-18Hydroxy-16,18-dimethyl-10-phenyl [11]-cytochalasa-6,13,19-triene-1,21dione (41)

Daldinia sp.

[38]

(6E,13E,16S*,18S*,19E)-12,18Daldinia sp. Dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6,13,19-triene-1,21dione (42)

[38]

(6Z,13E,16S*,17R*,18S*,19E)-17Hydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6,13,19-triene-1,21dione (43)

Daldinia sp.

[38]

(7S*,13E,16S*,18S*,19E)-6,7-Epoxy- Daldinia sp. 18-hydroxy-16,18-dimethyl-10phenyl-[11]-cytochalasa-13,19-diene1,21-dione (44)

[38]

(7S*,13E,16S*,18S*,19R*)-6,7Epoxy-18,19-dihydroxy-16,18dimethyl-10-phenyl-[11]-cytochalasa13-ene-1,21-dione (45)

Daldinia sp.

[38]

(7S*,13E,16S*,18R*,19E)-7,18Daldinia sp. Dihydroxy-16,18-dimethyl-10-phenyl[11]-cytochalasa-6(12),13,19-triene1,17,21-trione (46)

[38]

(continued)

16

H. Zhu et al.

Table 2 (continued) Compound

Origin

References

(5Z,7S*,13E,16S*,17R*,18S*,19E)-7- Daldinia sp. Hydroperoxy-17-hydroxy-16,18dimethyl-10-phenyl-[11]-cytochalasa5,13-19-triene-1,21-dione (47)

[38]

19,20-Epoxycytochalasin R Edwards (48)

Xylaria hypoxylon

[40]

18-Deoxy-19,20-epoxycytochalasin R Edwards (49)

Xylaria hypoxylon

[40]

18-Deoxy-19,20-epoxycytochalasin Q Edwards (50)

Xylaria hypoxylon

[40]

19,20-Epoxycytochalasin N Edwards (51)

Xylaria hypoxylon

[40]

19,20-Epoxycytochalasin C (52)

Xylaria hypoxylon

[40]

19,20-Epoxycytochalasin D (53)

Xylaria hypoxylon

[40]

RKS-1778 (54)

Phoma sp. SNF-1778

[41]

Deacetylcytochalasin C (55)

Metarrhizium anisopliae

[42]

(6R,16S,18R,21R)-18,21-Dihydroxy16,18-dimethyl-10-phenyl [11] cytochalasa-(13E,19E)-diene-1,7,17trione (56)

Metarrhizium anisopliae

[42]

Fragiformin A (57)

Hypoxylon fragiforme

[43]

Fragiformin B (58)

Hypoxylon fragiforme

[43]

(16S*,18S*,19R*)-12,18-Dihydroxy19-methoxy-16,18-dimethyl-10phenyl-[11]-cytochalasa-6(7),(13E)diene-1,21-dione (59)

Microporellus subsessilis

[44]

(6R*,7S*,16S*,18S*,19R*)-6,7Epoxy-18-hydroxy-19-methoxy16,18-dimethyl-10-phenyl-[11]cytochalasa-(13E)-ene-1,21-dione (60)

Microporellus subsessilis

[44]

Cytochalasin Z10 (61)

Endothia gyrosa

[45]

Cytochalasin Z11 (62)

Endothia gyrosa

[45]

19,20-Epoxicitocalasina (63)

Xylaria sp.

[46]

18-Deoxycytochalasin Q Edwards (64) Xylaria sp. SCSIO 156

[47]

21-O-Deacetylcytochalasin Q Edwards Xylaria sp. SCSIO 156 (65)

[47]

Cytochalasin H2 (66)

[48]

Xylaria sp. A23

(continued)

Progress in the Chemistry of Cytochalasans

17

Table 2 (continued) Compound

Origin

References

18-Methoxy cytochalasin J (67) (cytochalasin J1)

Phomopsis sp. IFB-E060 Phomopsis sp. (CMB-M0042F)

[49] [50]

Compound 1 (68) Cordyceps taii Bioorg Med Chem Lett (2015) 25:1823

[7]

Compound 2 (69) Cordyceps taii Bioorg Med Chem Lett (2015) 25:1823

[7]

Compound 3 (70) Cordyceps taii Bioorg Med Chem Lett (2015) 25:1823

[7]

18-Desoxy-19,20-epoxycytochalasin C (71)

KL-1.1

[51]

18-Desoxycytochalasin C (72)

KL-1.1

[51]

Phomocytochalasin (73)

Phomopsis theicola

[52]

Phomopsichalasin D (74)

Phomopsis spp. xy21 and xy22

[53]

Phomopsichalasin E (75)

Phomopsis spp. xy21 and xy22

[53]

Phomopchalasin C (76)

Phomopsis sp. shj2

[54]

Cytochalasin J2 (77)

Phomopsis sp. (CMB-M0042F)

[50]

Cytochalasin H1 (78)

Phomopsis sp. (CMB-M0042F)

[50]

Fragiformin E (79)

Hypoxylon fragiforme MUCL 51264

[55]

Compound 15ii (80) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Cytochalasin D1 (81)

Xylaria cf. curta

[56]

Cytochalasin C1 (82)

Xylaria cf. curta

[56]

6,12-Epoxycytochalasin D (83)

Xylaria longipes

[57]

6-epi-Cytochalasin P (84)

Xylaria longipes

[57]

7-O-Acetylcytochalasin P (85)

Xylaria longipes

[57]

7-Oxo-cytochalasin C (86)

Xylaria longipes

[57]

12-Hydroxycytochalsin Q Edwards (87)

Xylaria longipes

[57]

Phomopchalasin B (88)

Phomopsis sp. shj2

[54]

Cytochalasin J3 (89)

Phomopsis sp. (CMB-M0042F)

[50]

Xylastriasan A (90)

Xylaria striata

[58]

Phomopchalasin A (91)

Phomopsis sp. shj2

[54]

Cytochalasin H2 (92)

Phomopsis sp. (CMB-M0042F)

[50]

Chaetoconvosin A (93)

Chaetomium convolutum cib-100

[59]

Chaetoconvosin B (94)

Chaetomium convolutum cib-100

[59]

Curtachalasin A (95)

Xylaria curta E10

[60]

Curtachalasin B (96)

Xylaria curta E10

[60]

Chamiside A (97)

Chaetomium nigricolor F5

[61] (continued)

18

H. Zhu et al.

Table 2 (continued) Compound

Origin

References

Curtachalasin C (98)

Xylaria cf. curta

[62]

Curtachalasin D (99)

Xylaria cf. curta

[62]

Curtachalasin E (100)

Xylaria cf. curta

[62]

Xylarichalasin A (101)

Xylaria cf. curta

[63]

this group, which were isolated from Metarrhizium anisopliae in 1967 [16]. Their planar structures were determined in 1969, and the structure of cytochalasin D (2) was found to be identical with “zygosporin A” from Zygosporium masonii, for which the structure was determined by X-ray diffraction analysis [17–21]. Most of these [5.6.11]-cytochalasins share the same characteristic skeleton with an 11-membered carbocyclic ring C, and they differ from each other mainly by modifications of ring B and C. • Modification of ring B Normally, [5.6.11]-cytochalasins exhibit two methyl groups (Me-11 and Me-12), occurring at C-5 and C-6 of the isoindolone core. In terms of ring B, there are primarily two types: with or without a double bond. For the cytochalasins with a double bond in or at ring B, the double bond may be located at C-5, C-6, or between C-6 and C-12, such as in the examples cytochalasin C (1), zygosporin G (6), and cytochalasin D (2). Moreover, if the double bond is located at C-5 or between C-6 and C-12, normally, C-7 is generally an oxygenated sp3 methine (such as in 1–5 and 7–15); when the double bond is located at C-6, C-7 is correspondingly a sp2 methine (such as in 6, 25, and 27). Indeed, those cytochalasins without double bond in or at ring B are oxidized derivatives of the former type, as their double bond is oxidized to an epoxy group (such as in 20, 23, and 24) or further hydrolyzed to become two hydroxy groups (such as in 16–18, 21, and 22). Oxidation of C-12 also frequently occurs, and such compounds like 25, 28, 42, and 59 may possess a hydroxy group at C-12. In addition, compound 83 possesses an epoxy group between C-6 and C-12 [57], while compounds 74 and 75 have an unexpected carboxylic acid group at C-12 [53]. There are also several cytochalasins with other specific functional groups: (6R,16S,18R,21R)-18,21-dihydroxy-16,18-dimethyl-10-phenyl[11]cytochalasa(13E),(19E)-diene-1,7,17-trione (56) [42] and 19,20-epoxicitocalasina (63) [46] possess a carbonyl group at C-7, while phomopchalasin C (76) exhibits an unusual hydroperoxy group at C-7 [54]. • Modification of ring C Generally, [5.6.11]-cytochalasins have two methyl groups both at C-16 and C-18 and a double bond at C-13 of ring C, and occasionally, the methyl group at C-16 may be oxidized to a hydroxymethyl (29, 30, and 80) [8, 14] and the double bond at

Progress in the Chemistry of Cytochalasans

19

C-13 oxidized to an epoxy group (49) [40]. Besides the double bond at C-13, most of [5.6.11]-cytochalasins have an additional double bond at C-19; however, some may be oxidized to an epoxy (48–53) [40] or hydroxy group (35 and 40) [37–39], or even become reduced (39) [38]. Cytochalasin J2 (77) [50] is the only example from this group with three double bonds in ring C, and the additional double bond at C-17 is possibly formed by dehydration of the hydroxy group at C-18. The C-21 carbons of all [5.6.11]-cytochalasins are oxidized and most of them are oxygenated sp3 carbons, with these examples being carbonyl carbons (33–47 and 57–60). Additionally, oxidations occur normally at C-17 and C-18. Acylation is also an important aspect for the structural diversity of cytochalasins and normally, it occurs on C-21. Cytochalasin D monoacetate (4) is the only representative with an additional acetyl at C-7 [23]. It is not clear whether these acetylations are catalyzed by BGC (Biosynthetic Gene Cluster) enzymes or not. The modification of rings B and C for other groups of cytochalasans are similar to those of the cytochalasins. • Skeleton reconstruction Cytochalasin D1 (81) and cytochalasin C1 (82) [56], isolated from the liquid fermentation of fungus Xylaria cf. curta by Liu and coworkers, possess a unique 11membered macrocycle with an oxygen bridge between C-13 and C-20, forming an unusual 12-oxabicyclo[6.3.1]dodecane core. Phomopchalasins A (91) and B (88), two novel cytochalasins, were isolated from the endophytic fungus, Phomopsis sp. shj2, derived from Isodon eriocalyx var. laxiflora, by Pu and coworkers [54]. Both these compounds stand out from this group as they exhibit unexpected [5.6.5.8] and [5.6.6.7] ring systems, respectively, which are possibly derived from normal [5.6.11] cytochalasins by means of additional cyclization. Subsequently, three more cytochalasins with [5.6.5.8] and [5.6.6.7] ring systems were reported, including cytochalasin J3 (89), xylastriasan A (90), and cytochalasin H2 (92) from Phomopsis sp. (CMB-M0042F) or Xylaria striata [50, 58]. Cytochalasin J3 (89) could also be obtained from its precursor cytochalasin J (10) by an acid-mediated conversion [50], suggesting that these compounds may be artifactual in origin rather than actual natural products. In recent years, additional cytochalasins possessing unusual ring systems (93– 101) have been isolated. Chaetoconvosins A (93) and B (94) [59], with an unprecedented 6/6/5/5/7 pentacyclic ring system, were obtained from the wheat rhizospheric fungus Chaetomium convolutum cib-100. Their structures were elucidated on the basis of the analysis of their spectroscopic data and by X-ray crystallography. In the literature, the authors proposed a biosynthetic pathway starting from a phenylacetic acid unit, which enabled the construction of the six-membered ring A first with malonyl-CoA, and then by reaction with a PKS enzyme followed by Diels–Alder cyclization, Michael addition, and further modifications, to form a 6/6/5/5/7 ring system [59]. However, the present authors consider it would be more likely to afford chaetoconvosins A (93) and B (94) from cytochalasins such as phomopchalasin A (91) [54] and cytochalasin H2 (92) [50] by means of ring A expansion, considering cytochalasins are normally biosynthesized from an amino acid via PKS-NRPS hybrid

20

H. Zhu et al.

pathways [2]. Several years later, chamiside A (97) [61] was reported by Lou and coworkers from the endophytic fungus Chaetomium nigricolor F5, which shares the same ring A as that of both chaetoconvosins A (93) and B (94), while differing in the cleavage of ring C. In their paper, the authors proposed a biosynthesis pathway, and the most intriguing step was the oxidation and rearrangement in the five-membered pyrrolinone ring to generate the six-membered ring A, which also applies to chaetoconvosins A (93) and B (94). In any event, the structures of these three compounds are extraordinary and they are rare examples of cytochalasans with a six-membered ring A. Curtachalasins A (95) and B (96) were reported in 2018 by Liu and coworkers [60] from the endophytic fungus Xylaria curta E10, which are rare cytochalasans with an even-numbered (10-) carbocyclic ring C, while most other examples contain an oddnumbered (9-, 11-, and 13-) ring C. Condensation and rearrangement are proposed to be the key steps for the construction of their carbon skeleton. One year later, from the same fungus, curtachalasins C–E (98–100) [62] were identified, which share a similar ring C, but have an unprecedented bicyclo[3.3.1]lactam ring as compared with curtachalasins A (95) and B (97). From the fermented medium under optimized conditions, xylarichalasin A (101) [63], an unexpected halogenated cytochalasin, was isolated, and it has an unprecedented 6/7/5/6/6/6-fused polycyclic structure. The ring C part of xylarichalasin A (101) is similar to those of the curtachalasins A (95) and B (96), but the connection of Me-11 and the benzene ring (C-25) via a radical reaction generates a new cycloheptane ring. Moreover, the presence of two chlorine atoms located at C-10 and C-13 is rare. These findings show that the reconstruction of the isoindolone core, whether it is a bicyclo[3.3.1]lactam in 98–100 or an additional cycloheptane fused to the core structure in 101, adds a new dimension to the structural diversity of the cytochalasin family. Interestingly, some compounds of this subclass were obtained by bioconversion procedures. Chen and coworkers reported the first microbial transformation of L696,474 (26) [35], a cytochalasin isolated from Hypoxylon fragiforme ATCC 20995, using Actinoplanes sp. ATCC 53771, which led to the two new cytochalasins, 22hydroxy-L-696,474 (29) and 16β,22-dihydroxy-L-696,474 (30), and several new pyrichalasins (165–168). This work suggested that microbial biotransformation is a powerful tool to generate novel cytocalasin derivatives.

[5.6.13]-Cytochalasins and Related Derivatives [5.6.13]-Cytochalasins (Table 3 and Fig. 3) differ from [5.6.11]-cytochalasins by the occurrance of two more carbons in the macrocycle fused to the isoindolone core. In addition, [5.6.13]-cytochalasins normally exhibit only one methyl group on the macrocycle at C-16 (102–104 and 106–108), with the one exception of cytochalasin K Fex (105) [64]. The macrocycle of cytochalasin K Fex (105) is similar to those of the chaetoglobosins (Sect. 2.3), with methyl groups both at C-16 and C-18.

Progress in the Chemistry of Cytochalasans

21

Table 3 [5.6.13]-Cytochalasins Compound

Origin

References

Proxiphomin (102)

Phoma sp. S-298

[65]

Protophomin (103)

Phoma sp. S-298

[65]

Desoxaphomin (104)

Phoma sp. S-298

[66]

Cytochalasin K Fex (105)

Chalara microspora

[64] [67]

Ascochalasin (106)

Ascochyta heteromorpha

Seoxaphomin B (107)

Phoma sp.

[68]

Seoxaphomin C (108)

Phoma sp.

[68]

O

HN

HN

OH

HN OO

OO

102 (proxiphomin)

OO

103 (protophomin)

O

OH

HN

HN OO OAc

OO HO

106 (ascochalasin)

105 (cytochalasin K Fex)

OH

HN OO

O

HO

104 (desoxaphomin)

HO

107 (deoxaphomin B)

HN OO HO

108 (deoxaphomin C)

Fig. 3 Structures of [5.6.13]-cytochalasins

2.1.2

Lactone and Peroxyester Cytochalasins

Lactone cytochalasins refer to those compounds with a lactone functional group in the macrocycle (Table 4 and Fig. 4), which usually is located at C-21 or C-23 next to C-9. Biosynthetically, the lactone group could be derived from precursor cytochalasins via Baeyer–Villiger oxidation [2]. Thus, there are two main types of lactone cytochalasins, possessing a 12- or 14-membered lactone ring C derived from

22

H. Zhu et al.

Table 4 Lactone and peroxyester cytochalasins Compound

Origin

References

Cytochalasin A (109) (dehydrophomin)

Helminthosporium dematioideum

[16]

Cytochalasin B (110) (phomin)

Helminthosporium dematioideum Phoma sp. S-298

[16] [15]

Cytochalasin F (111)

Helminthosporium dematioideum

[71] [72]

Cytochalasin L (112)

Chalara microspora

[64]

Cytochalasin M (113)

Chalara microspora

[64]

Cytochalasin T (114)

Phoma exigua var. heteromorpha

[73]

Cytochalasin Z2 (115)

Pyrenophora semeniperda

[74]

Cytochalasin Z3 (116)

Pyrenophora semeniperda

[74]

Cytochalasin Z6 (117)

Phoma exigua var. heteromorpha

[75]

Cytochalasin B2 (118)

Phoma sp.

[68]

20-Deoxycytochalasin F (119)

Phoma sp.

[68]

Cytochalasin B3 (120)

Phoma sp. J08NF7

[76]

Cytochalasin B4 (121)

Phoma sp. J08NF7

[76]

Cytochalasin B5 (122)

Phoma sp. J08NF7

[76]

Cytochalasin B6 (123)

Phoma sp. J08NF7

[76]

Cytochalasin V (124)

Phoma exigua var. heteromorpha

[69]

Cytochalasin W (125)

Phoma exigua var. heteromorpha

[70]

(7S*,13E,16S*,18S*,19E)-7,18Dihydroxy-16,18-dimethyl-10phenyl-22-oxa-[12]-cytochalasa6(12),13,19-triene-1,21-dione (126)

Daldinia sp.

[38]

22-Oxa-[12]-cytochalasin-1 (127)

Rhinocladiella sp.

[77]

22-Oxa-[12]-cytochalasin-2 (128)

Rhinocladiella sp.

[77]

22-Oxa-[12]-cytochalasin-3 (129)

Rhinocladiella sp.

[77]

Rosellichalasin (130)

Rosellinia necatrix

[78]

Cytochalasin Z16 (131)

Aspergillus terreus

[79]

Cytochalasin Z17 (132)

Aspergillus terreus

[79]

Cytochalasin Z22 (133)

Spicaria elegans KLA03

[80]

Cytochalasin Z23 (134)

Spicaria elegans KLA03

[80]

Cytochalasin Z7 (135)

Spicaria elegans

[81]

Cytochalasin Z8 (136)

Spicaria elegans

[81]

Cytochalasin Z9 (137)

Spicaria elegans

[81]

Cytochalasin Z24 (138)

Aspergillus elegans ZJ-2008010

[82] (continued)

Progress in the Chemistry of Cytochalasans

23

Table 4 (continued) Compound

Origin

References

Arthriniumnin A (139)

Arthrinium arundinis

[83]

Arthriniumnin B (140)

Arthrinium arundinis ZSDS1-F3

[83]

Arthriniumnin C (141)

Arthrinium arundinis ZSDS1-F3

[83]

Arthriniumnin D (142)

Arthrinium arundinis ZSDS1-F3

[83]

Cytochalasin Z17 (143)

Aspergillus flavipes

[11]

Cytochalasin U Evidente (144)

Phoma exigua var. heteromorpha

[69]

[5.6.11]- and [5.6.13]-cytochalasins by inserting an oxygen atom at C-9. The [5.6.12] lactone cytochalasins always possess two methyl groups at C-16 and C-18, while the [5.6.14] lactone cytochalasins normally exhibit only one methyl group at C-16, except for cytochalasin L (112) and cytochalasin M (113) [64], with an additional methyl functionality at C-18. Cytochalasin U Evidente (144) [69] was isolated from liquid culture filtrates of Phoma exigua, and it possesses a unique peroxyester group. Although its structure was determined by means of spectroscopic analysis and chemical transformation, more evidence is still necessary to confirm its structural identity. For the co-isolated cytochalasin V (124) [69], it is possible that the original structure elucidation may be incorrect, because the 13 C NMR resonances at δ 142.8 and 148.3 ppm were assigned to C-20 and C-21, two adjacent carbonyl groups. The occurrence of a formyl group at C-20 in cytochalasin U Evidente (144) and at C-23 in cytochalasin V (124) is unusual, but this is found also in cytochalasin W (125) [70].

2.1.3

Cyclic Carbonate Cytochalasins

There are only four cytochalasins exhibiting cyclic carbonate groups (Table 5 and Fig. 5), comprising cytochalasin E (145), cytochalasin K Steyn (146), 6,12 cytochalasin K Steyn (147), and cytochalasin Z16 (148). The structure of cytochalasin E (145) was incorrect in the original reference and was revised later [71, 72, 84]. An epoxy group was assigned at C-6 and C-7 rather than a single hydroxy at C-7 and a double bond at C-4. To the best of our knowledge, there is no naturally occurring cytochalasan with a double bond at C-4; thus, if unusual structures are deduced when elucidating structures of cytochalasins, caution should always be expressed.

24

H. Zhu et al.

HN

O

HN

O

HN

OO

OO

109 (cytochalasin A)

HO

110 (cytochalasin B)

O

O OO

HO

O

HN

O

OH

OH

111 (cytochalasin F)

O

O

HN

OO

O

HN

OAc

O

112 (cytochalasin L)

O OO

OO HO

HO

O

113 (cytochalasin M)

114 (cytochalasin T)

OH O

HN

O

HN

OO

O

HN

OO

HO

OH

115 (cytochalasin Z2)

116 (cytochalasin Z3)

O

OH

117 (cytochalasin Z6)

O

OH

HN

O OO

HN

OO

OH

O

HN

OO

O OO

HO

O

118 (cytochalasin B2)

119 (20-deoxycytochalasin F)

HN

O

O OO

OO O

121 (cytochalasin B4)

OH

OH

OH

HN

120 (cytochalasin B3)

HO

122 (cytochalasin B5)

Fig. 4 Structures of lactone and peroxyester cytochalasins

HN

O OO O

123 (cytochalasin B6)

Progress in the Chemistry of Cytochalasans

25

OH

O O

HN

OH

OH

OH

O

O

O

HN

O

HN

O

O

O

OH

O

OH CHO

126 ((7S*,13E,16S*,18S*,19E)-7,18Dihydroxy-16,18-dimethyl-10-phenyl22-oxa-(12)-cytochalasa-6(12),13,19triene-1,21-dione)

125 (cytochalasin W)

124 (cytochalasin V)

O O

O

O

O OH

OH O

HN

OH

O

O

HN

O

O

O 127 (22-oxa-[12]-cytochalasin-1)

HN

O

O

HN

O

OH

O

O

HN

O

130 (rosellichalasin)

O

131 (cytochalasin Z16)

O

O

O

O

O

132 (cytochalasin Z17)

O

O

O OH

O O 133 (cytochalasin Z22)

HN

O

O

O O

O 136 (cytochalasin Z8)

Fig. 4 (continued)

O

HN

O 135 (cytochalasin Z7)

O

OH

O

HN

HN

O O 137 (cytochalasin Z9)

OH

O

OH

134 (cytochalasin Z23)

OH

O

OH

O

OH

HN

129 (22-oxa-[12]-cytochalasin-3)

OH

O

HN

O

O O

128 (22-oxa-[12]-cytochalasin-2 )

O

O

HN

O

OH

O O O

138 (cytochalasin Z24)

OH

26

H. Zhu et al.

OH

O

HN

O

HN

O

O

O

139 (arthriniumnin A)

O

141 (arthriniumnin C)

OH

O

HN

O

O

O

140 (arthriniumnin B)

OH

O

HN

O

O

O

O

HN

OH

O

OH

O

HN

O

O

O

O

O

O

143 (cytochalasin Z17)

142 (arthriniumnin D)

O

O

144 (cytochalasin U Evidente)

Fig. 4 (continued) Table 5 Cyclic carbonate cytochalasins Compound

Origin

References

Cytochalasin E (145)

Rosellinia necatrix

[71] [72] [84]

Cytochalasin K Steyn (146)

Aspergillus clavatus MRC 1181

[85]

6,12 -Cytochalasin K Steyn (147)

Mycotypha sp. UMF-006

[86] [84]

Cytochalasin Z16 (148)

Aspergillus flavipes

[11]

O

O

HN

OH

O

O

O

HN

OH

O

O

O OH

O O

O

146 (cytochalasin K Steyn)

145 (cytochalasin E)

OH

O

HN

O

O O

147 (Δ

O

HN OH

O 6,12

-cytochalasin K Steyn)

Fig. 5 Structures of cyclic carbonate cytochalasins

O

O O 148 (cytochalasin Z16)

O

O

Progress in the Chemistry of Cytochalasans

2.1.4

27

seco-Cytochalasins

In 2008, Gu and coworkers reported six novel cytochalasins named cytochalasins Z10 –Z15 (149–154) [87] from the marine-derived fungus, Spicaria elegans (Table 6 Table 6 Seco-cytochalasins Compound

Origin

References

Cytochalasin Z10 (149)

Spicaria elegans

[87]

Cytochalasin Z11 (150)

Spicaria elegans

[87]

Cytochalasin Z12 (151)

Spicaria elegans

[87]

Cytochalasin Z13 (152)

Spicaria elegans

[87]

Cytochalasin Z14 (153)

Spicaria elegans

[87]

Cytochalasin Z15 (154)

Spicaria elegans

[87]

Cytochalasin Z18 (155)

Aspergillus flavipes

[11]

Cytochalasin Z19 (156)

Aspergillus flavipes

[11]

Cytochalasin Z20 (157)

Aspergillus flavipes

[11]

Cytochalasin Z21 (158)

Spicaria elegans KLA03

[80]

and Fig. 6). Notably, ring C of these compounds is cleaved with a hydroxy group located at C-9. These are the first cytochalasins that contain an open chain, which is strikingly different from previously reported cytochalasins. These seco-cytochalasins possibly are derived from cyclic carbonate cytochalasins by means of hydrolysis. This same research group reported the three additional seco-cytochalasins Z18 –Z20 (155–157) [11] from Aspergillus flavipes that inhabit the mangrove plant, Acanthus ilicifolius, and two of them exhibit a carbonate group at C-9, which could be postulated as intermediates of seco-cytochalasins from cyclic carbonate cytochalasins. Later, from the work of the same group, cytochalasin Z21 (158) [80] was obtained from Spicaria elegans by the OSMAC method via adding d-tryptophan during its culturing procedure.

2.1.5

Mero-cytochalasins

As mentioned previously, mero-cytochalasans are a class of cytochalasans formed by the dimerization or polymerization of one or more cytochalasan molecules with one or more other natural product units. Thus, (7S*,16S*,18S*,19R*)7,18-dihydroxy-19-O-(4-methyl-(6E),(8E)-hexadecadienoyl)-16,18-dimethyl-10phenyl-[11]-cytochalasa-6(12),(13E)-diene-1,21-dione (159) [44], isolated from Microporellus subsessilis, could be considered as a mero-cytochalasin, as formed by the esterization between a cytochalasin and a long-chain fatty acid. Phomachalasins A–D (160–163) (Table 7 and Fig. 7) [88] are four intriguing mero-cytochalasins discovered by Evidente and coworkers from Phoma exigua var. exigua in 2011. Phomachalasin B (161), possessing an unusual [5.6.15] carbocyclic core, has an

28

H. Zhu et al. OH

OH

OH

OH OH

HN

OH

HN

HO

O

O

O

149 (cytochalasin Z10)

150 (cytochalasin Z11)

OH

OH

OH

OH OH

HN

152 (cytochalasin Z13)

OH

OH

O

OH

O

O

151 (cytochalasin Z12)

HN

OH

HN

HO

O

OH

HN

OH

O

OH

O

O

154 (cytochalasin Z15)

153 (cytochalasin Z14)

O

OH

OHO

OH O

O

HN O

O

O O

HN

O

O

O

155 (cytochalasin Z18)

O

O

O

156 (cytochalasin Z19)

O

O OH OH

OH

HN

O

O 157 (cytochalasin Z20)

OH

HN O

HO

158 (cytochalasin Z21)

Fig. 6 Structures of seco-cytochalasins

additional 1,2,3,4,6,7-hexasubstituted bicyclo[3.2.0]heptene moiety fused to the macrocycle at C-21 and C-22. Phomachalasins A (160), C (162), and D (163) [88], which share the same planar structure, could be derived from phomachalasins B (161) and related derivatives by Baeyer–Villiger oxidation. However, the relative configurations of each of phomachalasins A–D (160–163) between the isoindolone core and the 1,2,3,4,6,7-hexasubstituted bicyclo[3.2.0]heptane moiety were not determined. There was no discussion on the putative biosynthesis of these compounds, specifically from where the cyclic fragment C-27-C-33 is derived or how it is fused to the macrocycle via C-21 and C-22.

Progress in the Chemistry of Cytochalasans

29

Table 7 Mero-cytochalasins Compound

Origin

References

(7S*,16S*,18S*,19R*)-7,18-Dihydroxy-19-O(4-methyl-(6E,8E)-hexadecadienoyl)-16, 18-dimethyl-10-phenyl-[11]-cytochalasa6(12),(13E)-diene-1,21-dione (159)

Microporellus subsessilis

[44]

Phomachalasin A (160)

Phoma exigua var. exigua

[88]

Phomachalasin B (161)

Phoma exigua var. exigua

[88]

Phomachalasin C (162)

Phoma exigua var. exigua

[88]

Phomachalasin D (163)

Phoma exigua var. exigua

[88]

OH

OH

HN

HN O O O

HO

O O O O H2N HO

O

O

O HO 160 (phomochalasin A)

159 ((7S*,16S*,18S*,19R*)-7,18-dihydroxy-19-O-(4-methyl(6E,8E)-hexadecadienoyl)-16,18-dimethyl-10-phenyl-[11]cytochalasa-6(12),(13E)-diene-1,21-dione)

OH

OH

OH

HN

HN O O O H2N

O

HO

HN O

O O O O H2N HO

HO 162 (phomachalasin C)

O O

O

O HO 161 (phomachalasin B)

O O O O H2N HO

HO 163 (phomachalasin D)

Fig. 7 Structures of mero-cytochalasins

2.2 The Pyrichalasin Group Pyrichalasins normally include cytochalasans with a tyrosine unit incorporated into the polyketide backbone. However, broadly speaking, some related derivatives with a substituted benzene ring are also included in this category, no matter what the nature of the substituent groups (e.g., hydroxy, methoxy) or where the substituent groups are located (para-, meta-, or ortho). Thus, pyrichalasins represent a compound group closely related to the cytochalasins (in fact, some pyrichalasins were named “cytochalasins”, as they were usually co-isolated with compounds of the latter type), and the only difference is that pyrichalasins have additional substituents on the benzene

30

H. Zhu et al.

ring. The pyrichalasin group in total contains 46 members (Tables 8, 9 and 10 and Figs. 8, 9 and 10), including several compounds obtained by bioconversion or gene manipulation procedures.

2.2.1

Carbocyclic Pyrichalasins

This type of pyrichalasin contains a tricyclic [5.6.11] core structure (Table 8 and Fig. 8), and is similar to the [5.6.11]-cytochalasins—and therefore the most common type of pyrichalasins. Pyrichalasin H (164) [89], the first example of a pyrichalasin, was isolated from Pyricularia grisea IFO7287 by Nukina in 1987, when the name “pyrichalasin” was first introduced for this type of compound. As mentioned previously, bioconversion of L-696,474 (26) using Actinoplanes sp. ATCC 53771, provided a series of new pyrichalasins, including 22,29-dihydroxy-L696,474 (165), 22,28-dihydroxy-L-696,474 (166), 22,28,29-trihydroxy-L-696,474 (167), and 16β,22,28-trihydroxy-L-696,474 (168) as well as two new cytochalasins [14]. Thus, this bioconversion process introduced hydroxy groups at C-16 and C-21 and also on the benzene ring of L-696,474 (26). During the investigation of the biosynthesis pathway of pyrichalasin H (164), Skellam and coworkers obtained a series of new pyrichalasins (175–189) from Magnaporthe grisea NI980, using a gene knockout strategy [8], and this study also revealed that the non-proteinogenic amino acid O-methyltyrosine is the true precursor of pyrichalasin H (164). Interestingly, both compounds 181 and 184 exhibit Table 8 [5.6.11]- and [5.6.13]-Pyrichalasins Compound

Origin

References

Pyrichalasin H (164)

Pyricularia sp. IFO 7287

[89]

22,29-Dihydroxy-L-696,474 (165)

Actinoplanes sp. ATCC 53771

[14]

22,28-Dihydroxy-L-696,474 (166)

Actinoplanes sp. ATCC 53771

[14]

22,28,29-Trihydroxy-L-696,474 (167)

Actinoplanes sp. ATCC 53771

[14]

16β,22,28-Trihydroxy-L-696,474 (168)

Actinoplanes sp. ATCC 53771

[14]

4 -Hydroxy-deacetyl-18-deoxycytochalasin

H Trichoderma harzianum

[95]

(169) Phomopsichalasin F (170)

Phomopsis spp. xy21 and xy22

[53]

Phomopsichalasin G (171)

Phomopsis spp. xy21 and xy22

[53]

Compound 19 (172) Org Lett (2019) 21:8756

Hypoxylon fragiforme MUCL 51264

[55]

Compound 20 (173) Org Lett (2019) 21:8756

Hypoxylon fragiforme MUCL 51264

[55]

Compound 21 (174) Org Lett (2019) 21:8756

Hypoxylon fragiforme MUCL 51264

[55] (continued)

Progress in the Chemistry of Cytochalasans

31

Table 8 (continued) Compound

Origin

References

Compound 7 (175) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 8 (176) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 9 (177) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 11 (178) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 12 (179) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 13 (180) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 14 (181) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 15i (182) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 16 (183) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 17 (184) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 18 (185) Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 19 (186) magnachalasin H Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 26 (187) 4 -Cl-pyrichalasin H Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 29 (188) 4 -F-pyrichalasin H Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Compound 30 (189) 4 -Br-pyrichalasin H Org Lett (2019) 21:4163

Magnaporthe grisea NI980

[8]

Phenochalasin C (190)

Hypoxylon fragiforme

[96]

Phenochalasin D (191)

Hypoxylon fragiforme

[96]

Diaporthichalasin (192) phomopsichalasin

Phomopsis sp. Diaporthe sp.

[90] [91] [92] [93]

Diaporthalasin (193)

Diaporthaceae sp. PSU-SP2/4

[94]

Armochaetoglasin A (194)

Chaetomium globosum TW1-1

[10]

32

H. Zhu et al. OH

OH

OH HN

O

HN

HO

O OAc

O OAc

OH

165 (22,29-dihydroxy-L-696,474)

164 (pyrichalasin H)

OH

OH HO

OH HN

HO

OH HN

HO

O OAc

O OAc

166 (22,28-dihydroxy-L-696,474)

167 (22,28,29-trihydroxy-L-696,474)

OH

OH HO

OH HN O OAc

OH

HN

HO

O OH

168 (16ß,22,28-trihydroxy-L-696,474)

169 (4'-hydroxy-deacetyl-18deoxycytochalasin H)

O

OH

OH

OH HN

HO

HN

HN

HO

O OH

O OH

170 (phomopsichalasin F)

171 (phomopsichalasin G)

O OH 172 (compound 19) (Org Lett (2019) 21:8756)

O

O OH

HO

HN

HN

O

O OAc

O OAc

OH

174 (compound 21) (Org Lett (2019) 21:8756)

173 (compound 20) (Org Lett (2019) 21:8756)

HO

O

HN O O

O OH

175 (compound 7) (Org Lett (2019) 21:4163)

Fig. 8 Structures of [5.6.11]- and [5.6.13]-pyrichalasins

HN O O 176 (compound 8) (Org Lett (2019) 21:4163)

OH

Progress in the Chemistry of Cytochalasans

33

OH HN

O

HN

HO

O O

O OH

OH

178 (compound 11) (Org Lett (2019) 21:4163)

177 (compound 9) (Org Lett (2019) 21:4163)

OH

OH

OH HN

O

HN

HO

O OAc

O OAc 180 (compound 13) (Org Lett (2019) 21:4163)

179 (compound 12) (Org Lett (2019) 21:4163)

HO OH

OH HN

O

HN

O

O OAc

O OAc

OH

OH

182 (compound 15i) (Org Lett (2019) 21:4163)

181 (compound 14) (Org Lett (2019) 21:4163)

HO

HN

HO

O OAc

HN

O

O OAc

OH

OH

184 (compound 17) (Org Lett (2019) 21:4163)

183 (compound 16) (Org Lett (2019) 21:4163) OH

O

HN

HO

O OAc

HN

188 (compound 29) (4'-F-pyrichalasin H) (Org Lett (2019) 21:4163)

Fig. 8 (continued)

HN O OAc

Br OH

OH

HN O OAc 189 (compound 30) (4'-Br-pyrichalasin H) (Org Lett (2019) 21:4163)

OH

187 (compound 26) (4'-Cl-pyrichalasin H) (Org Lett (2019) 21:4163)

OH

HN O OAc

OH

186 (compound 19) (magnachalasin H) (Org Lett (2019) 21:4163)

OH

F

Cl

O OAc

OH

185 (compound 18) (Org Lett (2019) 21:4163)

OH

HO OH

HN O O 190 (phenochalasin C)

34

H. Zhu et al.

HO

HO

HN O O

HN

HO

HO

OO

192 (diaporthichalasin, phomopsichalasin)

191 (phenochalasin D)

HN OO

193 (diaporthalasin)

O OH

HO

HN OO HO

O

194 (armochaetoglasin A)

Fig. 8 (continued)

an unusual hydroxy group at C-10 while compounds 187–189 possess an unexpected halogen atom (Cl, F, and Br, respectively) [8]. Subsequently, this research group investigated the function of cytochalasan cytochrome P450 monooxygenases on the pyrichalasin BGC from Hypoxylon fragiforme MUCL 51264, and found that CYP3 catalyzed a site-selective epoxidation and led to the isolation of three new pyrichalasins (172–174) [55]. Phomopsichalasin [90], a new cytochalasin-like secondary metabolite, was isolated in 1995 from an endophytic fungus, Phomopsis sp., by Horn and coworkers, and possesses an unexpected 5/6/5/6/6 ring system and has an additional methyl group at C-8. Twelve years later, in 2007, another structurally related compound, diaporthichalasin (192) [91] was reported to share the same planar structure as that of phomopsichalasin. In addition, the relative configuration of diaporthichalasin (192) was confirmed by X-ray crystallographic analysis. After this, diaporthichalasin (192) and phomopsichalasin were deemed as being a pair of diastereomers. In 2012, Hoye and coworkers [92] analyzed the proton NMR data of phomopsichalasin and realized that the relative configuration proposed for this structure was likely to be wrong. Therefore, they further studied its relative configuration using both empirical and computational methods, and found that phomopsichalasin has an identical structure to that of diaporthichalasin (192). Then, Uchiro and coworkers [93] completed the total synthesis of diaporthichalasin (192) using the intramolecular Diels–Alder reaction of an α,β-unsaturated γ-hydroxylactam in an aqueous medium, which determined the absolute configuration of diaporthichalasin (192). Two years later, another analog, diaporthalasin (193) [94], was isolated from a marine-derived fungus, Diaporthaceae sp. PSU-SP2/4, and found to share the same absolute configuration of diaporthichalasin (192). Notably, with a β-oriented C-9-C-1 bond, the configuration of diaporthichalasin (192) and diaporthalasin (193) at C-9 differed from any other cytochalasans. However, the evidence was not sufficient, as their

Progress in the Chemistry of Cytochalasans

35

absolute configurations were determined only by the comparison of optical rotations. Thus it is recommended that a further investigation should be made concerning the unusual configuration assigned at C-9. Recently, Zhang and coworkers reported an unexpected pyrichalasin named armochaetoglasin A (194) [10] by feeding l-tyrosine into the culture medium of the endophytic fungus, Chaetomium globosum TW1-1. This represented the first tyrosine-derived cytochalasan alkaloid characterized by a [5.6.13] ring system, as found only rarely in both pyrichalasin and cytochalasin structures. Table 9 Lactone pyrichalasins Compound

Origin

References

Cytochalasin Z24 (195)

Eutypella sp. D-1

[97]

Cytochalasin Z25 (196)

Eutypella sp. D-1

[97]

Cytochalasin Z27 (197)

Xylaria sp. XC-16

[98]

Cytochalasin Z28 (198)

Xylaria sp. XC-16

[98]

Cytochalasin Z1 (199)

Pyrenophora semeniperda

[74]

Cytochalasin Z4 (200)

Phoma exigua

[75]

Cytochalasin Z5 (201)

Phoma exigua

[75]

HO

HN

O

O

O

HN

HO

O

O

O 196 (cytochalasin Z25)

195 (cytochalasin Z24)

OH

HO

O

HN

OH

O

O

O 198 (cytochalasin Z28)

OH

OH

O

HN

O OO

OO

HO 199 (cytochalasin Z1)

O

O

O

HN

O

HN

HO

197 (cytochalasin Z27)

HO

OH

O

200 (cytochalasin Z4)

Fig. 9 Structures of lactone pyrichalasins

OH

HO

HN

O OO HO

201 (cytochalasin Z5)

36

2.2.2

H. Zhu et al.

Lactone Pyrichalasins

This group of pyrichalasins contains seven members, including four with a 12membered lactone ring and three with a 14-membered lactone ring (Table 9 and Fig. 9). Of the four 12-membered lactone ring pyrichalasins, cytochalasins Z24 (195) and Z25 (196) [97] were isolated from the fungus Eutypella sp. D-1 when collected from the soil at a high latitude from the Arctic, while cytochalasins Z27 (197) and Z28 (198) [98] were obtained by bioassay-guided isolation of Xylaria sp. XC-16, an endophyte of Toona sinensis. The three 14-membered lactone ring pyrichalasins, cytochalasins Z1 (199) [74], Z4 (200), and Z5 (201) [75], were all discovered by Evidente and coworkers, from Pyrenophora semeniperda and Phoma exigua var. heteromorpha, respectively.

2.2.3

Cyclic Carbonate and Seco-Pyrichalasins

There are seven cyclic carbonate pyrichalasins (202–208) (Table 10, Fig. 10) that share the same 13-membered ring C with almost the same substitution pattern and differ mainly in their isoindolone moiety. Scoparasin C Deng (209), isolated from a marine fungus, Eutypella scoparia 1–15, is the only one belonging to the secopyrichalasins [99]. It also possesses a carbonate group, indicating that this secopyrichalasin is derived from a cyclic carbonate pyrichalasin.

2.3 The Chaetoglobosin Group The chaetoglobosins are a group of cytochalasans with a tryptophan unit in the core structure. Within this group, 133 chaetoglobosins are included in this Section (Tables 11, 12, 13 and 14 and Figs. 11, 12, 13 and 14). As described in Sects. 2.1 and 2.2, a [5.6.11] carbon ring system is the main type for both the cytochalasins and Table 10 Cyclic carbonate and seco pyrichalasins Compound

Origin

References

Phenochalasin A (202)

Phomopsis sp. FT-0211

[100]

Phenochalasin B (203)

Phomopsis sp. FT-0211

[100]

Scoparasin A (204)

Eutypella scoparia PSU-D44

[101]

Scoparasin B (205)

Eutypella scoparia PSU-D44

[101]

Cytochalasin Z26 (206)

Eutypella sp. D-1

[97]

Scoparasin C Rukachaisirikul (207)

Eutypella scoparia PSU-H267

[102]

Scoparasin D (208)

Eutypella scoparia 1–15

[99]

Scoparasin C Deng (209)

Eutypella scoparia 1–15

[99]

Progress in the Chemistry of Cytochalasans

37 O

HO

O

HN

O

O OH

O

HN

O

O

203 (phenochalasin B)

OH OH

OH

O

HN

OH O

202 (phenochalasin A)

O

O

O

O O

O

O OH

O

O

HN

O

O

O

O

OH

O O

204 (scoparasin A)

205 (scoparasin B)

O

HO

O

HN

O

O

O

HN

HO

O

O

O

OH

O O

O 206 (cytochalasin Z26)

207 (scoparasin C Rukachaisirikul)

O

OH

HO

O

O

HN

O

O

O OH

O

O

O

O

O

O 208 (scoparasin D)

O

HN

O

O 209 (scoparasin C Deng)

Fig. 10 Structures of cyclic carbonate and seco-pyrichalasins

pyrichalasins. In contrast, chaetoglobosins have a [5.6.13] carbon ring system in most cases (210–321), with only compounds 322–325 possessing a [5.6.11] ring system. Another feature concerning the structural diversity of chaetoglobosins emanates mainly from modifications within rings B and C. In terms of ring C, almost all of the chaetoglobosins possess a double bond at C-13, two methyl groups at C-16 and C-18, and a carbonyl group at C-23, and, most commonly, have additional double bonds at C-17 and C-21 and a carbonyl or hydroxy group at C-19 and C-20.

2.3.1

Carbocyclic Chaetoglobosins

[5.6.13]-Chaetoglobosins and Related Derivatives Chaetoglobosins A (210) and B (211) [103], isolated from Chaetomium globosum in 1973 by Natori and coworkers, are the first examples of the chaetoglobosin group

38

H. Zhu et al.

Table 11 [5.6.13]-Chaetoglobosins and derivatives Compound

Origin

References

Chaetoglobosin A (210)

Chaetomium globosum

[103] [110] [107] [104] [105] [109] [13]

Chaetoglobosin B (211)

Chaetomium globosum

[103] [110] [107] [109]

Chaetoglobosin C (212)

Chaetomium globosum

[106] [110] [111] [112] [108] [109]

Chaetoglobosin D (213)

Chaetomium globosum

[106] [110] [111] [107] [109]

Chaetoglobosin E (214)

Chaetomium globosum

[106] [110] [111] [108] [109]

Chaetoglobosin F (215)

Chaetomium globosum

[106] [110] [111] [108] [109]

Chaetoglobosin G (216)

Chaetomium globosum

[113] [106] [108] [109]

Chaetoglobosin J (217)

Chaetomium globosum

[113] [106] [108] [109]

19-O-Acetylchaetoglobosin B (218)

Chaetomium globosum

[114]

19-O-Acetylchaetoglobosin D (219)

Chaetomium globosum

[114]

19-O-Acetylchaetoglobosin A (220)

Chaetomium globosum

[114]

Prochaetoglobosin I (221)

Chaetomium subaffine

[115] (continued)

Progress in the Chemistry of Cytochalasans

39

Table 11 (continued) Compound

Origin

References

Prochaetoglobosin II (222)

Chaetomium subaffine

[115]

Prochaetoglobosin III (223)

Chaetomium subaffine

[115] [116]

Prochaetoglobosin IV (224)

Chaetomium subaffine

[115]

Prochaetoglobosin IIIed (225)

Chaetomium subaffine

[117] [116]

Isochaetoglobosin J (226)

Chaetomium subaffine

[117]

Chaetoglobosin Fex (227)

Chaetomium subaffine

[118]

20-Dihydrochaetoglobosin A (228)

Chaetomium subaffine

[118]

Chaetoglobosin O Sakamura (229)

Cylindrocladium floridanum

[119]

Penochalasin E (230)

Penicillium sp. OUPS-79

[120]

Penochalasin F (231)

Penicillium sp. OUPS-79

[120]

Penochalasin G (232)

Penicillium sp. OUPS-79

[120]

Penochalasin H (233)

Penicillium sp. OUPS-79

[120]

Chaetoglobosin Q (234)

Chaetomium globosum

[121]

Chaetoglobosin R (235)

Chaetomium globosum

[121]

Chaetoglobosin T (236)

Chaetomium globosum

[121]

Cytoglobosin B (237)

Chaetomium globosum QEN-14

[122]

Cytoglobosin C (238)

Chaetomium globosum QEN-14

[122]

Cytoglobosin D (239)

Chaetomium globosum QEN-14

[122]

Cytoglobosin E (240)

Chaetomium globosum QEN-14

[122]

Cytoglobosin F (241)

Chaetomium globosum QEN-14

[122]

Cytoglobosin G (242)

Chaetomium globosum QEN-14

[122]

Chaetoglobosin Kanokmedhakul (243)

Chaetomium elatum ChE01

[116]

Chaetoglobosin Y (244)

Chaetomium globosum (No. 64-5-8-2) [123]

Armochaetoglobin F (245)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin G (246)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin H (247)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin I (248)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin S (249)

Chaetomium globosum TW1-1

[125]

7-O-Acetylarmochaetoglobin S (250)

Chaetomium globosum TW1-1

[125]

Armochaetoglobin T (251)

Chaetomium globosum TW1-1

[125]

Armochaetoglobin U (252)

Chaetomium globosum TW1-1

[125]

Armochaetoglobin V (253)

Chaetomium globosum TW1-1

[125]

Armochaetoglobin W (254)

Chaetomium globosum TW1-1

[125]

Cytoglobosin H (255)

Chaetomium globosum

[126]

Cytoglobosin I (256)

Chaetomium globosum

[126] (continued)

40

H. Zhu et al.

Table 11 (continued) Compound

Origin

References

Isochaetoglobosin D (257)

Chaetomium globosum

[127] [108]

Armochaetoglasin D (258)

Chaetomium globosum TW1-1

[10]

Armochaetoglasin E (259)

Chaetomium globosum TW1-1

[10]

Armochaetoglasin F (260)

Chaetomium globosum TW1-1

[10]

Armochaetoglasin G (261)

Chaetomium globosum TW1-1

[10]

Armochaetoglasin H (262)

Chaetomium globosum TW1-1

[10]

Armochaetoglasin I (263)

Chaetomium globosum TW1-1

[10]

Oxichaetoglobosin A (264)

coculture of Aspergillus flavipes and Chaetomium globosum

[128]

Oxichaetoglobosin B (265)

[128]

Oxichaetoglobosin C (266)

[128]

Oxichaetoglobosin D (267)

[128]

Oxichaetoglobosin E (268)

[128]

Oxichaetoglobosin F (269)

[128]

Chaetomadrasin A (270)

Chaetomium madrasense 375

[129]

Armochaetoglosin A (271)

Chaetomium globosum TW1-1

[130]

Armochaetoglosin B (272)

Chaetomium globosum TW1-1

[130]

Chaetoglobosin K (273)

Diplodia macrospora

[131] [132]

Chaetoglobosin L (274)

Diplodia macrospora

[133]

Chaetoglobosin N (275)

Phomopsis leptostromiformis

[134] [135]

Chaetoglobosin P (276)

Discosia sp. TCF 9535

[136]

Chaetoglobosin O Cherton (277)

Phomopsis leptostromiformis ATCC 26115

[137]

Chaetoglobosin-510 (278)

Phomospis asparagi

[138]

Chaetoglobosin-540 (279)

Phomospis asparagi

[138]

Chaetoglobosin-542 (280)

Phomospis asparagi

[138]

Chaetoglobosin M (281)

mutated Diplodia macrospora

[139]

5’-F-Chaetoglobosin J (282)

Chaetomium globosum 1C51

[140]

5’-Cl-Chaetoglobosin J (283)

Chaetomium globosum 1C51

[140]

5’-Br-Chaetoglobosin J (284)

Chaetomium globosum 1C51

[140]

5’-F-Chaetoglobosin A (285)

Chaetomium globosum 1C51

[140]

5’-Cl-Chaetoglobosin A (286)

Chaetomium globosum 1C51

[140]

5’-Br-Chaetoglobosin A (287)

Chaetomium globosum 1C51

[140]

5’-F-Chaetoglobosin B (288)

Chaetomium globosum 1C51

[140] (continued)

Progress in the Chemistry of Cytochalasans

41

Table 11 (continued) Compound

Origin

References

5 -Cl-Chaetoglobosin B (289)

Chaetomium globosum 1C51

[140]

5 -Br-Chaetoglobosin B (290)

Chaetomium globosum 1C51

[140]

Chaetoglobosin Fa (291)

Chaetomium globosum

[141]

Chaetoglobosin W (292)

Chaetomium globosum IFB-E041

[142]

Armochaetoglobin J (293)

Chaetomium globosum TW1-1

[124]

Oxichaetoglobosin G (294)

coculture of Aspergillus flavipes and Chaetomium globosum

[128]

Oxichaetoglobosin H (295)

coculture of Aspergillus flavipes and Chaetomium globosum

[128]

Oxichaetoglobosin I (296)

coculture of Aspergillus flavipes and Chaetomium globosum

[128]

Penochalasin A (297)

Penicillium sp.

[143]

Penochalasin B (298)

Penicillium sp.

[143]

Penochalasin C (299)

Penicillium sp.

[143]

Penochalasin D (300)

Penicillium sp.

[120]

Armochaetoglobin K (301)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin L (302)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin M (303)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin N (304)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin O (305)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin P (306)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin Q (307)

Chaetomium globosum TW1-1

[144]

Armochaetoglobin R (308)

Chaetomium globosum TW1-1

[144]

Chaetoglobosin U (309)

Chaetomium globosum IFB-E019

[145]

Cytoglobosin A (310)

Chaetomium globosum QEN-14

[122]

Chaetoglobosin V Tan (311)

Chaetomium globosum IFB-E041

[142]

Chaetoglobosin Vb (312)

Chaetomium globosum

[146]

Armochaeglobine C (313)

Chaetomium globosum TW1-1

[147]

Armochaetoglobin X (314)

Chaetomium globosum TW1-1

[125]

Armochaetoglobin Y (315)

Chaetomium globosum TW1-1

[125]

Armochaetoglobin Z (316)

Chaetomium globosum TW1-1

[125]

Cytoglobosin Ab (317)

Chaetomium globosum

[148]

Armochaetoglosin C (318)

Chaetomium globosum TW1-1

[130]

Chaetomadrasin B (319)

Chaetomium madrasense 375

[129]

Armochaeglobine A (320)

Chaetomium globosum TW1-1

[147]

Armochaeglobine B (321)

Chaetomium globosum TW1-1

[147]

42

H. Zhu et al.

Table 12 [5.6.11]-Chaetoglobosins Compound

Origin

References

Cytochalasin G (322)

Nigrosabulum sp. (CMI 171,019)

[149]

Cytochalasin X (323)

Pseudeurotium zonatum

[150]

Cytochalasin Y (324)

Pseudeurotium zonatum

[150]

Cytochalasin Z (325)

Pseudeurotium zonatum

[150]

Table 13 Seco- and other chaetoglobosins Compound

Origin

References

Armochaetoglobin A (326)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin B (327)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin C (328)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin D (329)

Chaetomium globosum TW1-1

[124]

Armochaetoglobin E (330)

Chaetomium globosum TW1-1

[124]

Armochaetoglasin B (331)

Chaetomium globosum TW1-1

[10]

Armochaetoglasin C (332)

Chaetomium globosum TW1-1

[10]

Yamchaetoglobosin A (333)

Chaetomium globosum YNH-16

[151]

Table 14 Mero-chaetoglobosins Compound

Origin

References

MBJ-0038 (334)

Chaetomium sp. f24230

[152]

MBJ-0039 (335)

Chaetomium sp. f24230

[152]

MBJ-0040 (336)

Chaetomium sp. f24230

[152]

Cytochathiazine A (337)

coculture of Aspergillus flavipes and Chaetomium globosum

[154]

Cytochathiazine B (338)

coculture of Aspergillus flavipes and Chaetomium globosum

[154]

Cytochathiazine C (339)

coculture of Aspergillus flavipes and Chaetomium globosum

[154]

Aureochaeglobosin A (340)

Chaetomium globosum

[153]

Aureochaeglobosin B (341)

Chaetomium globosum

[153]

Aureochaeglobosin C (342)

Chaetomium globosum

[153]

and their planar structures were determined mainly through chemical reactions. In 1976, Silverton and coworkers [104] completed the first X-ray diffraction analysis of chaetoglobosin A (210) and determined its relative configuration, and further X-ray analysis by the same group in 1978 confirmed its absolute configuration [105]. After that, a series of chaetoglobosins have been elucidated [106–109], and biosynthesis studies on these compounds were also carried out [109].

Progress in the Chemistry of Cytochalasans

43

O

N H

HN

N H

OO

HN

OH

O

N H

OO O

211 (chaetoglobosin B)

OH

OH

HN

N H

OO

OO

O

HN OO

O

214 (chaetoglobosin E)

HO

HN

OH

N H

OO

HN

O

HN

HN

O

O

HN OO

OAc

220 (19-O-acetylchaetoglobosin A)

221 (prochaetoglobosin I)

O

N H

HN

N H

OO

O 222 (prochaetoglobosin II)

O

HN

N H

OO

HN OO

O

223 (prochaetoglobosin III)

224 (prochaetoglobosin IV )

OH

OH

N H

HN OO

O 225 (prochaetoglobosin III ed)

OAc

218 (19-O-acetylchaetoglobosin B)

N H

OO

OAc

O

219 (19-O-acetylchaetoglobosin D)

OO

O

N H

OO

HN

OH

217 (chaetoglobosin J )

OH

N H

N H

OO

O

O

216 (chaetoglobosin G)

O

215 (chaetoglobosin F)

OH

N H

O

O

N H HO

OO

212 (chaetoglobosin C)

HN

OH

O

213 (chaetoglobosin D)

HN

OH

210 (chaetoglobosin A)

N H

O

OH

N H

HN

N H

OO O

O

226 (isochaetoglobosin J)

Fig. 11 Structures of [5.6.13]-chaetoglobosins and derivatives

HN OO

O HO 227 (chaetoglobosin Fex)

44

H. Zhu et al. O

N H

O

OH

HN

N H

OO

HN

OH

HO

228 (20-dihydrochaetoglobosin A)

N H

OO

HN OO

230 (penochalasin E)

O

N H

OH

HN

N H

OO

HN

HN

OH

HN

N H

OO

OH

OH

OH

N H

OO HO

HN

OO OH

236 (chaetoglobosin T)

N H

OO

OH

237 (cytoglobosin B)

HN

OH

O

235 (chaetoglobosin R)

HN

OH

O 233 (penochalasin H)

234 (chaetoglobosin Q)

N H

OO

HO

N H

OO

HN

OH

O 232 (penochalasin G)

OH OH

O

N H

OO

OH

O 231 (penochalasin F)

N H

OH

O

OH

O

229 (chaetoglobosin O Sakamura)

HN OO OH

HO

OH

HO 238 (cytoglobosin C)

239 (cytoglobosin D)

OH

OH

OH

N H

HN

N H

OO O

240 (cytoglobosin E)

Fig. 11 (continued)

O

HN

N H

OO O

241 (cytoglobosin F)

OH

HN OO HO

242 (cytoglobosin G)

OH

Progress in the Chemistry of Cytochalasans

45 HO O

OH

HN

N H

N H

OO

HN

N H

OO

OO

O

HO

243 (chaetoglobosin V Kanokmedhakul)

HN

244 (chaetoglobosin Y)

O

245 (armochaetoglobin F)

HO

HN

N H

N H

OO HO

HN

O

246 (armochaetoglobin G)

OO HO

247 (armochaetoglobin H)

N H

OO

HN

O HN OO

O

HO

O

249 (armochaetoglobin S)

O

N H

OO

250 (7-O-acetylarmochaetoglobin S)

OH

248 (armochaetoglobin I)

OH OAc

HN

HO

HN

OH

O

OH OH

N H

N H

OO

HO

O

251 (armochaetoglobin T) HO

OH OH

HO

HN

N H

N H

OO O

OH

HN

253 (armochaetoglobin V)

OH OH

N H

HN

HO

HN

HN OO

HO

HO

N H

OH

HN OO O

O

256 (cytoglobosin I)

HN OO

O

254 (armochaetoglobin W)

N H

OO

O

255 (cytoglobosin H)

N H

OO

OH OH

N H

OO

HN

O

HO

O

252 (armochaetoglobin U)

N H

OO

O

257 (isochaetoglobosin D)

N H

HN OO HO

O 258 (armochaetoglasin D)

Fig. 11 (continued)

HO 259 (armochaetoglasin E)

260 (armochaetoglasin F)

46

H. Zhu et al. O

O

O

HN

N H

N H

OO

O HO 261 (armochaetoglasin G)

O HN

O O 262 (armochaetoglasin H)

O

N H

HN

N H

OO

HN

HN OO HO

O

HO

O

HO

264 and 265 (oxichaetoglobosins A and B)

OH

O

N H

OO

OO

O O 263 (armochaetoglasin I)

OH

O

HO

HN

N H

OO

266 and 267 (oxichaetoglobosins C and D)

O

268 and 269 (oxichaetoglobosins E and F) OH

O

N H

HN

N

OO

O O 270 (chaetomadrasin A)

HN OO

N

OH 271 (armochaetoglosin A)

O

N H

HN

N H

OO

272 (armochaetoglosin B) O

HN

N H

OO

OH O 274 (chaetoglobosin L)

HN OO

O O 275 (chaetoglobosin N)

O

O

HN

N H

OO O

OO

OH

OH O 273 (chaetoglobosin K)

N H

HN

HN

HO

OH

276 (chaetoglobosin P)

N H

OO

HN OO

O

277 (chaetoglobosin O Cherton)

278 (chaetoglobosin-510) O

N H

HN

N H

OO O

OH

279 (chaetoglobosin-540)

Fig. 11 (continued)

HN OO

OH OH 280 (chaetoglobosin-542)

N H

HN OO O

O

281 (chaetoglobosin M)

Progress in the Chemistry of Cytochalasans

F

47

Br

Cl HN

N H

HN

N H

OO OH

O

O

Cl

Br

HN

HN

N H

OO

O N H

O

O HN NH2

HN OO

HN

NH O

OO

O

HN

NH O

O

HO

O

297 (penochalasin A)

296 (oxichaetoglobosin I)

O

OH

N H

HN

NH O

O

O

293 (armochaetoglobin J)

N H

OO

OH

N H

HN

OH

O

294 and 295 (oxichaetoglobosins G and H )

OH

O

HO

292 (chaetoglobosin W)

OH

HO

N H

OO

O

291 (chaetoglobosin Fa ) O

O

OH

HN

N H

HN

N O H

O

298 (penochalasin B)

Fig. 11 (continued)

O

299 (penochalasin C)

OH

O

290 (5'-Br-chaetoglobosin B)

O

O

OO

OH

O

289 (5'-Cl-chaetoglobosin B)

N H

HN

N H

OO

OH

O

HN OF

HN

N H

OO

N H

OH Br

HN

288 (5'-F-chaetoglobosin B)

OH

O

287 (5'-Br-chaetoglobosin A)

OH Cl

O

OO

OH

O

286 (5'-Cl-chaetoglobosin A)

OH F

HN

N H

OO

OH

O

285 (5'-F-chaetoglobosin A)

N H

284 (5'-Br-chaetoglobosin J)

O

F

OH

O

283 (5'-Cl-chaetoglobosin J)

O

N H

OO

OH

O

282 (5'-F-chaetoglobosin J)

HN

N H

OO

O

300 (penochalasin D

48

H. Zhu et al. OH

O OH

HN

N H

NH

HN

N H

O

NH

O

NH

HN

NH

O

305 (armochaetoglobin O)

O

NH

N H

O

HN

NH

HN

O 308 (armochaetoglobin R)

OH O 310 (cytoglobosin A)

HN

OO

OH

O

HN OO O

OH

311 (chaetoglobosin V Tan)

N H

OO

HN

O

OH

313 (armochaeglobine C)

O

316 (armochaetoglobin Z)

OO O

315 (armochaetoglobin Y) O

OH

N H

OO

HN

OH

314 (armochaetoglobin X)

OH

HN

O

N H

OO

HN

N

OO O

OH

317 (cytoglobosin Ab)

OH

312 (chaetoglobosin Vb)

OH OH

HN

Fig. 11 (continued)

HN

OH O 309 (chaetoglobosin U)

N H

OO

O

N H

O

OH

N H

OO

O

NH O

O 306 (armochaetoglobin P )

N H

O

OH

N H

HN

OH

O 307 (armochaetoglobin Q)

N H

O

N H

O

O

HN

303 (armochaetoglobin M)

OH

N H

O

304 (armochaetoglobin N)

N H

O

302 (armochaetoglobin L)

OH

HN

NH O

O

301 (armochaetoglobin K)

N H

HN

N H

O

HN OO O

318 (armochaetoglosin C)

Progress in the Chemistry of Cytochalasans

49

OOH

HO N H

O

O

OH

HN O

N H

OO

HN

N H

OO

OO O

O

OH

O

HN

319 (chaetomadrasin B)

O

OH

321 (armochaeglobine B)

320 (armochaeglobine A)

Fig. 11 (continued)

O

N H

OH

HN O O

O

N H

HN O O

323 (cytochalasin X)

322 (cytochalasin G)

O

OH

N H

OH

HN O O

O

HO

324 (cytochalasin Y)

N H

O

HN O O

O

325 (cytochalasin Z)

Fig. 12 Structures of [5.6.11]-chaetoglobosins

• Methylated chaetoglobosins The first 10,11-dimethyl-chaetoglobosin, chaetoglobosin K (273) [131], was reported from Diplodia macrospora from infected corn plants (Zea mays) by Horace Cutler and coworkers, which exhibited significant inhibition of the growth of wheat coleoptiles and toxicity to day-old chicks. Soon after this, the structure of chaetoglobosin K (273) was confirmed by X-ray diffraction analysis [132]. Subsequently, chaetoglobosin L (274) was isolated from the same fungus [133], and then chaetoglobosins P (276) [136] and M (281) [139] were isolated. Chaetoglobosin M (281) is the first cytochalasan obtained from a mutated Diplodia macrospora specimen. 10,11-Dimethyl-chaetoglobosins were also reported from fungi of the genus Phomopsis, including chaetoglobosin N (275) [134, 135], chaetoglobosin O Cherton (277) [137], chaetoglobosin-510 (278) [138], chaetoglobosin-540 (279) [138], and chaetoglobosin-542 (280) [138]. Armochaetoglosins A (271) and B (272) [130] represent the first examples of 1 -N-methyl-chaetoglobosins, which possibly may be biosynthesized from the additive 1-methyl-l-tryptophan rather than from tryptophan, because both compounds

50

H. Zhu et al. O

O

N H

HN

NH

O

N H

HN

OH

OO OH

N H

OO O

O

HN

N H

OO O O

O

HN OO O O

O

330 (armochaetoglobin E)

329 (armochaetoglobin D)

O

328 (armochaetoglobin C)

O

HN

OH

HN

OH

327 (armochaetoglobin B)

O

N H

N H

OO

O

326 (armochaetoglobin A)

O

O O

331 (armochaetoglasin B) OH

N H

HN

N H

OO O O

O

332 (armochaetoglasin C)

HN OO O O

O

333 (yamchaetoglobosin A)

Fig. 13 Structures of seco- and other chaetoglobosins

were isolated from Chaetomium globosum TW1-1 by feeding 1-methyl-l-tryptophan during the fermentation process. • Halogenated-chaetoglobosins In 2011, Tan and coworkers reported a series of halogenated-chaetoglobosins by precursor-fed (halogenated-tryptophan) cultivation of endophytic Chaetomium globosum 1C51 [140], which afforded nine novel “unnatural” halogenatedchaetoglobosins including 5 -F-chaetoglobosin J (282), 5 -Cl-chaetoglobosin J (283), 5 -Br-chaetoglobosin J (284), 5 -F-chaetoglobosin A (285), 5 -Cl-chaetoglobosin A (286), 5 -Br-chaetoglobosin A (287), 5 -F-chaetoglobosin B (288), 5 -Clchaetoglobosin B (289), and 5 -Br-chaetoglobosin B (290). Besides these “unnatural” natural compounds, chemically synthesized halogenated-chaetoglobosin were also reported. Gao and coworkers obtained chaetoglobosin Fa (291) [141], containing a fluorine atom at C-23 and an oxolane ring, by the treatment of chaetoglobosin F (215) with (diethylamino) sulfur trifluoride (DAST) in dichloromethane. These compounds are also rare examples of halogenated-cytochalasans, in addition to the previously mentioned halogenated-cytochalasins 4 -Cl-pyrichalasin H (187), 4 -F-pyrichalasin H (188), and 4 -Br-pyrichalasin H (189) [8].

Progress in the Chemistry of Cytochalasans

51

O

N H

O

O O

HN O

HN

N H

O HO

O

OH

HO

336 (MBJ-0040)

N H

OO S

HN

N H

OO S

O

N O

O O

337 (cytochathiazine A)

N H

N H

N H

O

HN

OH

O OH O

O OH O HO

HO

340 (aureochaeglobosin A )

N H

O

341 (aureochaeglobosin B)

O

O 339 (cytochathiazine C)

OH

O

O

N

O

O 338 (cytochathiazine B)

O

HN

OO S

O

N H

O

HN

O

N O

N H

O

OH

HN

OH

HO

335 (MBJ-0039)

OH

N H

O

HO HO

334 (MBJ-0038)

O

OH

HO

O O

HN

N H HO

O

OH

HO

O O

O

HN O

O OH O HO

342 (aureochaeglobosin C)

Fig. 14 Structures of mero-chaetoglobosins

• Unusually oxygenated-chaetoglobosins There are some chaetoglobosins that possess an unusual oxygen bridge. Chaetoglobosin W (292) [142], from Chaetomium globosum IFB-E041, was the first chaetoglobosin with an oxolane ring formed via an oxygenated bridge between C-3 and C-6. After this, additional chaetoglobosins with an oxolane ring were reported by Zhang and coworkers, including armochaetoglobin J (293) from Chaetomium globosum TW1-1 [124] and oxichaetoglobosins G–I (294–296) from the coculture of Aspergillus flavipes and Chaetomium globosum [128]. Furthermore, chaetoglobosin Fa (291) [141], as mentioned previously, has an unusual oxygen bridge between C-20

52

H. Zhu et al.

and C-23, forming a ring system similar to that of pyrrole-chaetoglobosins described later. Certain oxidations at selected positions and functional groups are being presented here. Armochaetoglasin G (261) [10] and oxichaetoglobosins A–F (264–269) [128] are rare chaetoglobosins with a C-10 substituent that has been oxidized to a hydroxy or methoxy group. More interestingly, the indole group of armochaetoglobin T (251) [125] is oxidized and represents the first chaetoglobosin characterized by a 2 ,3 epoxy-indole moiety. The same oxidation is also found in several other compounds that include armochaetoglasin H (262) [10] and oxichaetoglobosins G (294) and H (295) [128]. Additionally, it is notable that oxichaetoglobosin I (296) [128] is the first example of a naturally occurring chaetoglobosin possessing a 2-nor-indole group that is derived from chaetoglobosin W (292) [142] by oxidation. • Pyrrole-chaetoglobosins This is a group of chaetoglobosins with an unusual pyrrole ring fused to the macrocyclic scaffold of chaetoglobosins that are possibly formed via transamination, cyclization, and dehydration reactions [144]. Penochalasins A–C (297–299) [143], which represent the first pyrrole-chaetoglobosins, were isolated from a Penicillium sp. fungus separated from a marine alga Enteromorpha intestinalis in 1996 by Numafa and coworkers. Their structures were elucidated by NMR spectroscopic analysis and chemical transformations. A few years later, another pyrrolechaetoglobosin, penochalasin D (300) [120], was discovered from the same fungus, in which the pyrrole ring is partially reduced. Then, for some time, there were no reports of new pyrrole-chaetoglobosin derivatives. However, in 2015 Zhang and coworkers isolated eight new pyrrole-chaetoglobosins named armochaetoglobins K– R (301–308) [144], from Chaetomium globosum TW1-1, which inhabits the medicinal terrestrial arthropod Armadillidium vulgare. They also proposed a biosynthesis pathway for pyrrole-chaetoglobosins from the precursor of chaetoglobosin A (210) by reactions including transamination, cyclization, and dehydration. The structure of armochaetoglobin K (301) was determined by X-ray diffraction analysis, and this was the first time the skeleton of a pyrrole-chaetoglobosin was confirmed using this technique. Isochaetoglobosin Db [148], separated from Chaetomium globosum SNSHI-5, was assigned with an unusual structure having a pyrrole ring that differed from other pyrrole-chaetoglobosins. However, its 1 H and 13 C NMR data are closely related to those of penochalasin C (299) [143], and it appears that their structures are identical. Since their NMR data were measured in different solvents and there was no detailed structural elucidation description for isochaetoglobosin Db , this supposition is not yet confirmed. • [5.6.10.5]-Chaetoglobosins Chaetoglobosins with a [5.6.10.5] ring system are a specific group derived from normal [5.6.13]-chaetoglobosins by a nucleophilic reaction, leading to the connection of C-17 and C-21. The first [5.6.10.5]-chaetoglobosin chaetoglobosin U (309) [145] was isolated by Tan and coworkers from the solid culture of Chaetomium globosum

Progress in the Chemistry of Cytochalasans

53

IFB-E019. Following this, ten chaetoglobosins (310–319) belonging to this same group have been reported, with all of these isolated from the fungus Chaetomium globosum. Moreover, chaetomadrasin B (319) [129] from Chaetomium madrasense 375 represents the first example of a chaetoglobosin characterized by possessing both a hydroxy and a carbonyl group on the indole moiety. • [5.6.7.5]- and [5.6.12]-Chaetoglobosins Armochaeglobines A (320) and B (321) [147], two chaetoglobosins with unprecedented carbon skeletons, were also obtained from the fungus Chaetomium globosum TW1-1, of which armochaeglobine A (320) features a unique tetracyclic 5/6/7/5 fused ring system and armochaeglobine B (321) possesses a rare 12-membered carbon scaffold. Both compounds represent the first examples of chaetoglobosins with an even-numbered (10- and 12-) carbocyclic ring C, differing from those of other cytochalasans by having a 9-, 11-, 13-, or 15-membered carbocyclic ring C. In particular, armochaeglobine A (321) possesses an unusual peroxide functionality in ring B. Their biosynthesis pathways were proposed starting from chaetoglobosin A (210), which undergoes oxidation, Michael addition, electrophilic addition, decarboxylation and peroxidation, and aldol condensation reactions.

[5.6.11]-Chaetoglobosins Both [5.6.11]-cytochalasins and [5.6.11]-pyrichalasins are the main types in their respective groups; however, [5.6.11]-chaetoglobosins (Table 12 and Fig. 12) represent a minor variant among the chaetoglobosin analogs. Four chaetoglobosins have been found to possess the [5.6.11] ring system, namely, cytochalasin G (322) [149] and cytochalasins X–Z (323–325) [150]. Cytochalasin G (322) [149] was the first [5.6.11]-chaetoglobosin to be isolated and was obtained by Cameron and coworkers from a Nigrosabulum sp. in 1974. Almost 30 years later, in 2002, Munro and coworkers re-isolated this compound from Pseudeurotium zonatum, along with three new [5.6.11]-chaetoglobosins named cytochalasins X–Z (323–325) [150]. Nearly 20 years since then have passed, and still today, these four compounds remain the only naturally occurring chaetoglobosins containing an 11-membered macrocycle.

2.3.2

seco-Chaetoglobosins

The first seco-chaetoglobosins, armochaetoglobins A–E (326–330) [124], were reported by Zhang and coworkers in 2015 from the fungus Chaetomium globosum TW1-1. From this same fungus, a series of pyrrole-chaetoglobosins was also isolated, as mentioned previously. Interestingly, armochaetoglobin A (326), as an unprecedented 19,20-seco-chaetoglobosin, also contains a pyrrole ring. All these compounds possibly are derived from normal chaetoglobosins or pyrrole-chaetoglobosins by the

54

H. Zhu et al.

oxidative cleavage of the C-19-C-20 bond. Later, the same group discovered two new seco-chaetoglobosins, armochaetoglasins B and C (331 and 332) [10], from the above-mentioned fungal source. Moreover, Ding and coworkers also isolated a 19,20seco-chaetoglobosin named yamchaetoglobosin A (333) [151] from Chaetomium globosum YNH-16. Thus, all of these seco-chaetoglobosins were discovered from the fungus Chaetomium globosum of different origins.

2.3.3

Mero-chaetoglobosins

In 2013, Shin-ya and coworkers reported three novel metabolites including MBJ0038 (334), MBJ-0039 (335), and MBJ-0040 (336) from Chaetomium sp. f24230 collected from a soil sample [152]. The structures of these compounds are closely related to those of normal chaetoglobosins; however, they have an additional unit (known as epicoccine) fused to C-21 and C-22, exhibiting a complicated ring system. They are the first examples of mero-chaetoglobosins, but regrettably, no biosynthesis pathway was proposed for these metabolites and their configurations were not determined. In 2018, aureochaeglobosins A–C (340–342) [153], three novel [4 + 2] cycloaddition heterodimers of chaetoglobosin and aureonitol derivatives, were obtained from a culture of the endophytic fungus Chaetomium globosum, which also represent a new type of mero-chaetoglobosins. Their structures were elucidated by extensive spectroscopic data analysis as well as by single-crystal X-ray diffraction and the use of the modified Mosher’s method. In the same year, cytochathiazines A–C (337–339) [154], which represent a new type of mero-chaetoglobosins, were isolated from a coculture of Chaetomium globosum and Aspergillus flavipes. These are the first natural products featuring an unprecedented 2H-1,4-thiazine functional group, and a plausible biosynthesis pathway with a chaeto-globosin molecule and a dipeptide as the main constitutional units was proposed [154].

2.4 The Aspochalasin Group Aspochalasins are the third-largest group of cytochalasans that feature the incorporation of leucine into a polyketide backbone (Tables 15, 16, 17, 18 and 19 and Figs. 15, 16, 17, 18 and 19). In this section, 127 aspochalasins are included (343–469), and, similar to the cytochalasins and pyrichalasins, most aspochalasins possess a [5.6.11] tricyclic ring system. In terms of their ring C, aspochalasins normally possess a double bond at C-13, a methyl group at C-14, and a carbonyl group at C-21. Xylarisin (360) [155] is the only example of an aspochalasin exhibiting a [5.6.11] ring system, and is without a methyl group at C-14. This was isolated from the marine-derived fungus Xylaria sp. PSU-F100 by Rukachiaisirikul and coworkers in 2009.

Progress in the Chemistry of Cytochalasans

55

Table 15 [5.6.11]-Aspochalasins and related derivatives Compound

Origin

References

Aspochalasin A (343)

Aspergillus microcysticus

[156] [157]

Aspochalasin B (344)

Aspergillus microcysticus

[156] [157]

Aspochalasin C (345)

Aspergillus microcysticus

[156] [157]

Aspochalasin D (346)

Aspergillus microcysticus

[156] [157] [158]

Aspochalasin E (347)

unidentified fungal strain (FA2277)

[159]

Phomacin C (348)

Phoma sp. (ATCC 74342)

[160]

Aspochalasin G (349)

Aspergillus sp. FO-4282

[161]

TMC-169 (350)

Aspergillus flavipes

[162]

Aspochalasin H (351)

Aspergillus sp. AJ117509

[163]

Aspochalasin Z (352)

Aspergillus niveus LU 9575

[5] [6]

Aspochalasin K (353)

Aspergillus flavipes

[164]

Aspochalasin L (354)

Aspergillus flavipes

[165]

Aspochalasin M (355)

Spicaria elegans

[166]

Aspochalasin N (356)

Spicaria elegans

[166]

Aspochalasin O (357)

Spicaria elegans

[166]

Aspochalasin P (358)

Spicaria elegans

[166]

Aspochalasin Q (359)

Spicaria elegans

[166]

Xylarisin (360)

Xylaria sp. PSU-F100

[155]

Aspochalasin R (361)

Spicaria elegans

[167]

Aspochalasin S (362)

Spicaria elegans

[167]

Aspochalasin T (363)

Spicaria elegans

[167]

Aspochalasin U (364)

Aspergillus sp. F00685

[168]

Trichalasin G (365)

Trichoderma gamsii

[169]

Aspochalasin V (366)

Aspergillus sp. CCTCC No. M2013631

[170]

Aspochalasin W (367)

Aspergillus sp. CCTCC No. M2013631

[170]

Flavichalasine F (368)

Aspergillus flavipes

[171]

Flavichalasine G (369)

Aspergillus flavipes

[171]

Flavichalasine H (370)

Aspergillus flavipes

[171] (continued)

56

H. Zhu et al.

Table 15 (continued) Compound

Origin

References

Flavichalasine I (371)

Aspergillus flavipes

[171]

Flavichalasine J (372)

Aspergillus flavipes

[171]

Flavichalasine K (373)

Aspergillus flavipes

[171]

Flavichalasine L (374)

Aspergillus flavipes

[171]

Flavichalasine M (375)

Aspergillus flavipes

[171]

18-Oxo-19,20-dihydrophomacin C (376) (Phomacin D)

Westerdykella dispersa Parastagonospora nodorum

[172] [173]

18-Oxo-19-methoxy-19,20-dihydrophomacin C (377)

Westerdykella dispersa

[172]

18-Oxo-19-hydroxy-19,20-dihydrophomacin C (378)

Westerdykella dispersa

[172]

19,20-Dihydrophomacin C (379)

Westerdykella dispersa

[172]

19-Methoxy-19,20-dihydrophomacin C (380)

Westerdykella dispersa

[172]

19-Hydroxy-19,20-dihydrophomacin C (381)

Westerdykella dispersa

[172]

Aspochalasinol A (382)

coculture of Aspergillus flavipes and Chaetomium globosum

[128]

Aspochalasinol B (383) Aspochalasinol C (384) Aspochalasinol D (385)

[128] [128] [128]

Phomacin E (386)

Parastagonospora nodorum

[173]

Phomacin F (387)

Parastagonospora nodorum

[173]

Spicochalasin A (388)

Spicaria elegans

[166]

Trichoderone A (389)

Trichoderma gamsii

[174]

Trichoderone B (390)

Trichoderma gamsii

[174]

Trichalasin H (391)

Trichoderma gamsii

[169]

Aspergilluchalasin (392)

Aspergillus sp. PSU-RSPG185

[175]

Flavichalasine A (393)

Aspergillus flavipes

[171]

Flavichalasine B (394)

Aspergillus flavipes

[171]

Flavichalasine C (395)

Aspergillus flavipes

[171]

Flavichalasine D (396)

Aspergillus flavipes

[171]

Flavichalasine E (397)

Aspergillus flavipes

[171]

Aspergillin PZ (398)

Aspergillus awamori

[176]

16-Hydroxymethylaspergillin PZ (399)

Westerdykella dispersa

[177]

Trichodermone (400)

Trichoderma gamsii

[178]

Aspochalazine A (401)

Aspergillus sp. Z4

[179]

Flavipesine A (402)

Aspergillus flavipes

[180]

Flavipesine B (403)

Aspergillus flavipes

[180]

Progress in the Chemistry of Cytochalasans

57

Table 16 [5.6.9]- and [5.6.7]-Aspochalasins and related derivatives Compound

Origin

References

Periconiasin A (404)

Periconia sp. F-31

[182]

Periconiasin B (405)

Periconia sp. F-31

[182]

Periconiasin C (406)

Periconia sp. F-31

[182]

Periconiasin I (407)

Periconia sp. F-31

[183]

Periconiasin D (408)

Periconia sp. F-31

[184]

Periconiasin E (409)

Periconia sp. F-31

[184]

Periconiasin F (410)

Periconia sp. F-31

[184]

Periconiasin J (411)

Periconia sp. F-31

[183]

Periconiasin G (412)

Periconia sp. F-31

[185] [186]

Table 17 Lactone aspochalasins Compound

Origin

References

Phomacin A (413)

Phoma sp. (74342)

[160]

Phomacin B (414)

Phoma sp. (74342)

[160]

Aspochalasin F (415)

Aspergillus sp. (FO-4282)

[161]

Aspochalasin I (416)

Aspergillus flavipes

[164]

Aspochalasin J (417)

Aspergillus flavipes

[164]

Trichalasin C (418)

Trichoderma gamsii

[188]

Trichalasin D (419)

Trichoderma gamsii

[188]

Trichalasin B (420)

Trichoderma gamsii

[9]

Aspochalasin A1 (421)

Aspergillus elegans

[82]

Trichalasin E (422)

Trichoderma gamsii

[169]

Trichalasin F (423)

Trichoderma gamsii

[169]

Flavichalasine N (424)

Aspergillus flavipes PJ03-11

[187]

Amiaspochalasin A (425)

Aspergillus micronesiensis

[189]

Amiaspochalasin D (426)

Aspergillus micronesiensis

[189]

Amiaspochalasin E (427)

Aspergillus micronesiensis

[189]

Amiaspochalasin H (428)

Aspergillus micronesiensis

[189]

16α-Methylaspochalasin J (429)

Westerdykella dispersa

[177]

Iizukine D (430)

Aspergillus iizukae

[190]

Amiaspochalasin B (431)

Aspergillus micronesiensis

[189]

Amiaspochalasin C (432)

Aspergillus micronesiensis

[189]

Flavichalasine O (433)

Aspergillus flavipes PJ03-11

[187]

58

H. Zhu et al.

Table 18 seco-Aspochalasins Compound

Origin

References

Pericoannosin A (434)

Periconia sp. F-31

[184]

Secochalasin A (435)

Aspergillus micronesiensis

[191]

Secochalasin B (436)

Aspergillus micronesiensis

[191]

Amiaspochalasin F (437)

Aspergillus micronesiensis

[189]

Amiaspochalasin G (438)

Aspergillus micronesiensis

[189]

2.4.1

Carbocyclic Aspochalasins

[5.6.11]-Aspochalasins and Related Derivatives The first aspochalasins were aspochalasins A–D (343–346) (Table 15 and Fig. 15) [156], which were discovered from the fungus Aspergillus microcysticus in 1979. Their structures were elucidated by extensive spectroscopic studies and chemical degradation. The skeleton and stereochemistry at C-17 and C-18 of aspochalasin C (345) were then determined by X-ray diffraction analysis of its 17,18-di-O-acetyl derivative [157], and aspochalasin D (346) was considered to be C-18 epimer of aspochalasin C (343). In 2001, Hayakawa and coworkers re-isolated aspochalasin D (346) from Aspergillus sp. AJ1 17509, and revised the previous stereochemical assignment indicating that aspochalasin D (346) is the C-17 epimer of aspochalasin C (343) [158]. Many aspochalasins have been found to possess additional functional groups besides the normal structural modifications. Phomacin C (348) [160], which was isolated from the fungus Phoma sp. in 1997, possesses a unique hydroxymethyl substituent at C-16 that had not then been observed among other aspochalasins. In 2017, Yang and coworkers isolated six more aspochalasins with a hydroxymethyl group at C-16 (376–381) [172] from the marine sediment-derived fungus Westerdykella dispersa, and determined the structure of 18-oxo-19,20-dihydrophomacin C (376) by single-crystal X-ray diffraction. Aspochalasins N (356) and O (357) [166], isolated from the fungus Spicaria elegans, were obtained by employing the OSMAC approach via changing the culture conditions. Both aspochalasin N (356) and O (357) possess an additional acetonyl group at C-20, but whether they are naturally occurring products or not was not discussed biosynthetically. Recently, overexpression of the transcription factor gene phmR encoded in the phm gene cluster resulted in the production of a congener, phomacin F (387) [173]. Aspochalasins V (366) and W (367) [170] were isolated from the culture broth of Aspergillus sp., which was found in the gut of a marine isopod Ligia oceanica. Both compounds are unusual as a methylthiol group was found to occur at C-20 of their macrocyclic ring. This is the first report on methylthio-substituted aspochalasins, and also on the first methylthio-substituted cytochalasans, which possibly are formed by methylthiotransferases [181].

Progress in the Chemistry of Cytochalasans

59

HN

HN O O

HN O O

O

O

343 (aspochalasin A)

O O

OH

O

344 (aspochalasin B)

OH

HO

345 (aspochalasin C)

OH OH HN

HN

HN O O

O O

OH

HO

346 (aspochalasin D)

OH

O O

HN O O

O

349 (aspochalasin G)

HO

348 (phomacin C)

347 (aspochalasin E)

HN

HN

O O

OH

O O

OH

O

OH

OH

351 (aspochalasin H)

350 (TMC-169)

OH OH HN

O

HN O O

HN O O

352 (aspochalasin Z)

353 (aspochalasin K)

HN

HN O O

OH

O

355 (aspochalasin M)

O O

OH

O

OH

OH

354 (aspochalasin L)

HN O O

O O

O

O 356 (aspochalasin N)

OH

O

O 357 (aspochalasin O)

O OH HN

HN O O

O

O O

OH

358 (aspochalasin P)

HN O

O

OH

361 (aspochalasin R)

O O

O

359 (aspochalasin Q)

HN O O

OH

HN

360 (xylarisin)

HN O O HO

O

362 (aspochalasin S)

Fig. 15 Structures of [5.6.11]-aspochalasins and related derivatives

O O HO

O

363 (aspochalasin T)

OH

60

H. Zhu et al.

OH OH

OH HN

HN O O

HN O O

O 365 (trichalasin G)

OH

364 (aspochalasin U)

O O

OH

S

O

366 (aspochalasin V)

OH OMe NH

NH

HN O O

S

O O

HO

O

O O

OH

368 (flavichalasine F)

367 (aspochalasin W)

NH

NH O O

OH

370 (flavichalasine H)

O O

O

HO

371 (flavichalasine I)

O O

NH O O

OH OH O

373 (flavichalasine K)

O

O

O O

OH

374 (flavichalasine L)

O O

OH

NH O O

O

376 (18-oxo-19,20-dihydrophomacin C) (phomacin D)

O O

OH

O O

O

377 (18-oxo-19-methoxy-19,20dihydrophomacin C)

HO

379 (19,20-dihydrophomacin C)

Fig. 15 (continued)

NH O O

380 (19-methoxy-19,20dihydrophomacin C)

OH O

378 (18-oxo-19-hydroxy-19,20dihydrophomacin C)

OH OH

HO

OH

OH NH

OH

O NH

OH O

375 (flavichalasine M)

O NH

OH OH O

372 (flavichalasine J)

NH

NH

OH

369 (flavichalasine G)

NH O O

HO

NH O O

OH HO

381 (19-hydroxy-19,20-dihydrophomacin C)

Progress in the Chemistry of Cytochalasans

61 HO

HO

HO

NH

NH

NH O O

O O

O

HO

382 (aspochalasinol A)

HO

O O

O

OH

O

384 (aspochalasinol C)

383 (aspochalasinol B)

HO

OH

OH O

NH

NH O O

OH

O

385 (aspochalasinol D)

NH O O

O O

O

O

O

387 (phomacin F)

386 (phomacin E)

O

O

HN

OH

O O

O O

OH O 388 (spicochalasin A)

O

HN

HN

O O

OH

390 (trichoderone B)

389 (trichoderone A )

OH O

HN

HN O O

391 (trichalasin H)

NH O O

OH

O O

OH OH

392 (aspergilluchalasin)

OH O

OH

393 (flavichalasine A )

OH O

NH

NH O O

O O

OH

394 (flavichalasine B)

O

NH O O

OH OH

OH O

396 (flavichalasine D)

395 (flavichalasine C)

OH O

NH O O

OH OH OH 397 (flavichalasine E )

Fig. 15 (continued)

O

NH O O

398 (aspergillin PZ)

O

NH OH

O O 399 (16-hydroxymethylaspergillin PZ)

62

H. Zhu et al. OH

O O

OH

O H N

HN

HN

O OH

O O 400 (trichodermone)

OH O

HN O

OH

401 (aspochalazine A)

O OH

O

HN OH

402 (flavipesine A)

O OH 403 (flavipesine B)

Fig. 15 (continued)

• [5.6.6.7]-Aspochalasins This is a group of aspochalasins with a [5.6.6.7] ring system, derived from [5.6.11]aspochalasins by the connection of C-13 and C-19. After spicochalasin A (388) [166] was isolated initially from the marine-derived fungus Spicaria elegans by Gu and coworkers in 2009, to date altogether 12 [5.6.6.7]-aspochalasins have been reported (389–400), and most of them exhibit an oxygen bridge between C-14 and C-17 or C-18. Trichoderone A (389) [174], similar to some chaetoglobosins such as chaetoglobosin W (292), possesses an oxygen bridge between C-3 and C-6. Interestingly, the double bond of trichoderone A (389) at C-13 remains after the linkage of C-13 and C-19. This compound has an unprecedented double bond between C-7 and C-8, which is possibly the only example among the cytochalasans. 16Hydroxymethylaspergillin PZ (399) [177] was characterized after purification from a solid fermentation culture of Westerdykella dispersa isolated from marine sediments, which has an additional hydroxymethyl group at C-16 as compared with the usual common [5.6.6.7]-aspochalasins. Trichodermone (400) [178], reported from Trichoderma gamsii, is a spirocytochalasan with an unprecedented tetracyclic nucleus, with its formation proposed by the oxidative cleavage of the C-6 and C-7 bond followed by esterification between C-7 and C-14. • [5.6.7.6]-Aspochalasins Interestingly, aspochalazine A (401) [179], which was isolated from a culture broth of Aspergillus sp. Z4 derived from the gut of the marine isopod Ligia oceanica, is the first aspochalasin with an unusual [5.6.7.6] ring system and it possesses a nitrogen bridge between C-18 and C-21. Soon after the report of the occurrence of compound 401, flavipesines A (402) and B (403) [180], which share the same [5.6.7.6] skeleton, but with an oxygen bridge between C-18 and C-21 rather than a nitrogen bridge, were isolated and characterized from Aspergillus flavipes. Compared with [5.6.6.7]-aspochalasins, these [5.6.7.6]-aspochalasins represent a much smaller group, which could be explained by a different cyclization mechanism of formation.

Progress in the Chemistry of Cytochalasans

63 OH

HN

HN OO

HN OO

HO

404 (periconiasin A)

HN O H

OH

O

408 (periconiasin D)

HN OO

HO

OO O

405 (periconiasin B)

406 (periconiasin C)

HO

407 (periconiasin I)

OH

OH HN O

HN O HO

OH HN O

HO

410 (periconiasin F )

409 (periconiasin E)

411 (periconiasin J)

(R)

HN OO

412 (periconiasin G)

Fig. 16 Structures of [5.6.9]- and [5.6.7]-aspochalasins and related derivatives

[5.6.9]- and [5.6.7]-Aspochalasins and Related Derivatives All the aspochalasins of this class were reported by Dai and coworkers (Table 16 and Fig. 16). In 2013 and 2016, this research group reported successively the isolation of periconiasins A–C (404–406) [182] and periconiasin I (407) [183], four new aspochalasins with an unprecedented 5/6/9 tricyclic ring system, from the endophytic fungus Periconia sp. F-31. The structure and absolute configuration of periconiasin A (404) were confirmed by X-ray crystallographic analysis. Not long after this, periconiasins D–J (408–411) [183, 184] were isolated from the same fungus, with these analogs containing a rare 5/6/6/5 tetracyclic ring system that presumably is derived from these above-mentioned [5.6.9]-aspochalasins. Periconiasin G (412) [185] is the only derivative with a [5.6.7] ring system that possesses a seven-membered ring C, and is possibly the simplest cytochalasan. Its proposed configuration at C-14 was revised as (R) by total synthesis [186]. Details of the total synthesis of periconiasin G (412) are provided in Sect. 5.

2.4.2

Lactone Aspochalasins

Most lactone aspochalasins have a [5.6.12] lactone ring system present (Table 17 and Fig. 17). Phomacins A (413) and B (414) [160], isolated from a Phoma sp., the first examples of lactone aspochalasins, differ from other members by possessing an

64

H. Zhu et al.

OH O

HN

OH

O O

O

HN

O O

OH

413 (phomacin A )

O

HN

O 414 (phomacin B)

O

O

OH

415 (aspochalasin F)

OH

O

HN

OH

O O

OH

O

O

OH

O

OH

OH

O

O

HN

O

O O

O

O

419 (trichalasin D)

OH

418 (trichalasin C)

O

HN

OH

O

417 (aspochalasin J)

O O

HN

O O

416 (aspochalasin I)

HN

O

HN

OH

421 (aspochalasin A1)

420 (trichalasin B)

OOH

O

HN

OH

O O

O HO

OH O

OH

O O

O

O

O O

HN

O

O

HN O

OH

O O

OH

OH

432 (amiaspochalasin C)

Fig. 17 Structures of lactone aspochalasins

O

OH O

OH

430 (iizukine D)

O

HN

O

OH

O

OH

429 (16a-methylaspochalasin J)

O O

O 427 (amiaspochalasin E)

O

HN

431 (amiaspochalasin B)

O

HN

OH

O

OH

428 (amiaspochalasin H)

HN

O

HN

O

424 (flavichalasine N4)

426 (amiaspochalasin D)

425 (amiaspochalasin A)

HN

O HO

OH

423 (trichalasin F)

O

OH

O

OH

O

O

HN

O

422 (trichalasin E)

HN

O

HN

O

N H

OH HO

O

433 (flavichalasine O)

Progress in the Chemistry of Cytochalasans

65

additional methyl or hydroxymethyl group at C-16 of ring C. Methylation at C-16 was also observed in amiaspochalasin A (425) [169] and 16α-methylaspochalasin J (429) [177]. Moreover, amiaspochalasin A (425) is the only derivative in this general class with no methyl group at C-13, while amiaspochalasins B (431) and C (432) possess an unusual [5.6.6.8] ring system [169]. Trichalasin E (422) [169] was isolated from the plant endophytic fungus Trichoderma gamsii, and represents the first cytochalasan with an unusual hydroperoxy group. Flavichalasine O (433) [187] was proposed as the first example of a cytochalasan possessing a nitrogen-oxygen heterocycle in the macrocyclic ring moiety. However, more evidence is necessary for the confirmation of the structure postulated, as its NMR data resemble closely those of the co-isolated lactone-aspochalasin, flavichalasine N (424).

2.4.3

seco-Aspochalasins

Pericoannosin A (434) (Table 18 and Fig. 18) [184], with an unusual hexahydro-1Hisochromen-5-isobutylpyrrolidin-2-one skeleton, has a quite different biosynthesis pathway as compared with normal cytochalasans. It shares the same precursor (1,5dihydro-pyrrol-2-one) with several co-isolated aspochalasins, with the 1,5-dihydropyrrol-2-one unit undergoing specific reduction, Claisen, and hemiacetal reactions to yield a hexahydro-1H-isochromen-5-isobutylpyrrolidin-2-one skeleton, and with subsequent dihydroxylation, thus producing pericoannosin A (434). No characteristic Diels–Alder reaction occurred to construct rings B and C; thus, although it is considered to be a seco-aspochalasin, essentially, it may be regarded as a related compound with the same precursors as aspochalasin rather than a real seco-aspochalasin. In total, four members of the seco-aspochalasin group were reported recently from Aspergillus micronesiensis, including secochalasins A (435) and B (436) [191]

O OH

O HN

HO O

HN

OH O O

HN

O

O

435 (secochalasin A)

434 (pericoannosin A)

O

O O O

O

436 (secochalasin B)

OH

OH

O

O OH

HN O

OH

O

437 (amiaspochalasin F)

Fig. 18 Structures of seco-aspochalasins

OH

HN O

438 (amiaspochalasin G)

O

66

H. Zhu et al.

and amiaspochalasins F (437) and G (438) [189]. This is the first report of actual seco-aspochalasins, with ring C cleaved between C-17 and C-18 or between C-9 and C-21.

2.4.4

Mero-aspochalasins and Aspochalasin Homodimers

Besides the merocytochalasans mentioned in Sects. 2.1 and 2.3, most merocytochalasans are generated by involving an aspochalasin unit (Table 19 and Fig. 19). The discovery of structurally complex merocytochalasans within this group may be considered to be a milestone in research on the cytochalasans, because of their fascinating ring systems and scaffolds with the presence of multiple chiral centers. The first type of mero-aspochalasins is formed by the connection of a peptide or an amino acid moiety to an aspochalasin unit. Aspochalamins A–D (439–442) [5, 6], four mero-aspochalasins constituted by the connection of a tripeptide-like moiety to an aspochalasin unit at C-19, were isolated in 2004 from an endosymbiotic fungus Aspergillus niveus, collected from the gut of a woodlouse species by Fiedler and coworkers. These were the first examples of merocytochalasans, although the term “merocytochalasan” had not yet been put forward at the time. However, to date, insufficient attention to this group has been paid, and so their relative configurations at C-17, C-18, and C-19 have still not determined. Recently, two new cysteine residuecontaining mero-aspochalasins, cyschalasins A (443) and B (444) [191] were isolated from the endophytic fungus Aspergillus micronesiensis, which represent a new type of merocytochalasan featuring a modified cysteine residue with an unusual sulfur atom. A plausible biosynthesis pathway was proposed, and possibly this finding may lead to additional attention from organic and biosynthesis chemists. Recently, Liu and coworkers isolated a mero-aspochalasin named iizukine C (445) [190] from the saline soil fungus Aspergillus iizukae, and it is the first aspochalasin featuring a unique 1,2,4-triazole functionality at C-19. Its structure was determined by extensive spectroscopic analysis inclusive of the 1 H-15 N HMBC spectrum. Another type of mero-aspochalasin is formed by the dimerization of one or more aspochalasins with one or more epicoccine units. Spicarins A (446) and B (447) [192], the first mero-aspochalasins with a epicoccine unit, were isolated from the fungus Spicaria elegans KLA03 in 2016. The structure of spicarin A (446) was determined by X-ray diffraction analysis. In the several years following, Zhang and coworkers reported a series of mero-aspochalasins (448–468) belonging to this type. The term “merocytochalasan” was put forward initially in 2017, and “heterodimer”, “heterotrimer”, and “heterotetramers” have also gradually come into use for these merocytochalasans. Asperchalasine A (455) [193], the first cytochalasan heterotrimer featuring an unusual decacyclic 5/6/11/5/5/6/5/11/6/5 ring system with as many as 20 chiral centers, was isolated from the solid culture broth of Aspergillus flavipes, together with three biogenetically related intermediates, asperchalasines B–D (448–450). The structure of asperchalasine A (455), with multiple quaternary carbon atoms and chiral

Progress in the Chemistry of Cytochalasans

67

Table 19 Mero-aspochalasins and aspochalasin-homodimer Compound

Origin

References

Aspochalamin A (439)

Aspergillus niveus LU 9575

[5] [6]

Aspochalamin B (440)

Aspergillus niveus LU 9575

[5] [6]

Aspochalamin C (441)

Aspergillus niveus LU 9575

[5] [6]

Aspochalamin D (442)

Aspergillus niveus LU 9575

[5] [6]

Cyschalasin A (443)

Aspergillus micronesiensis PG-1

[191]

Cyschalasin B (444)

Aspergillus micronesiensis PG-1

[191]

Iizukine C (445)

Aspergillus iizukae

[190]

Spicarin A (446)

Spicaria elegans KLA03

[192]

Spicarin B (447)

Spicaria elegans KLA03

[192]

Asperchalasine B (448)

Aspergillus flavipes

[193]

Asperchalasine C (449)

Aspergillus flavipes

[193]

Asperchalasine D (450)

Aspergillus flavipes

[193]

Asperchalasine E (451)

Aspergillus flavipes

[180]

Asperchalasine F (452)

Aspergillus flavipes

[180]

Asperchalasine G (453)

Aspergillus flavipes

[180]

Asperchalasine H (454)

Aspergillus flavipes

[180]

Asperchalasine A (455)

Aspergillus flavipes

[193]

Amichalasine D (456)

Aspergillus micronesiensis PG-1

[194]

Amichalasine E (457)

Aspergillus micronesiensis PG-1

[194]

Amichalasine A (458)

Aspergillus micronesiensis PG-1

[195]

Amichalasine B (459)

Aspergillus micronesiensis PG-1

[195]

Amichalasine C (460)

Aspergillus micronesiensis PG-1

[195]

Epicochalasine A (461)

Aspergillus flavipes

[196]

Epicochalasine B (462)

Aspergillus flavipes

[196]

Aspergilasine A (463)

Aspergillus flavipes QCS12

[197]

Aspergilasine B (464)

Aspergillus flavipes QCS12

[197]

Aspergilasine C (465)

Aspergillus flavipes QCS12

[197]

Aspergilasine D (466)

Aspergillus flavipes QCS12

[197]

Asperflavipine B (467)

Aspergillus flavipes QCS12

[4]

Asperflavipine A (468)

Aspergillus flavipes QCS12

[4]

Dimericchalasine A (469)

Aspergillus micronesiensis PG-1

[194]

68

H. Zhu et al.

19

HN O O O

HN

HN O O

OH

O

N H

O

NH

439 (aspochalamin A) N H 440 (aspochalamin B)

N H

441 (aspochalamin C) R = OH 442 (aspochalamin D) R = H

NH

HN O O

S

O O

OH

O

R

N H

O

NH

HN

HN

S

O O

OH

O

N

OH

N

OH

N NH

O

HO O

O 443 (cyschalasin A)

445 (iizukine C)

444 (cyschalasin B)

HN

NH

HN O O

O OAc

O O

O O

OAc

O

O

OAc AcO

O

O

OH

O

HO

O

OH

HN O O O

O

OH

OH

OH

449 (asperchalasine C)

OH

448 (asperchalasine B)

NH O O

O

HO

OAc

447 (spicarin B)

NH

O

OAc

OAc

446 (spicarin A)

O O

OAc

HO

O

450 (asperchalasine D (450)

Fig. 19 Structures of mero-aspochalasins and aspochalasin-homodimers

OO O

O

OH

HO O

OH

451 (asperchalasine E)

Progress in the Chemistry of Cytochalasans

HN

HN OO

OH

HO O

O

O

O

HN O

O O HN

O

O HN

OH

O

O OH

O HN

O

O

O HN

OH

OH

O OH

O

O OH

O O

OH

O HN

464 (aspergilasine B)

Fig. 19 (continued)

OH

O

O OH

OH

465 (aspergilasine C)

O OH

OH

463 (aspergilasine A)

O

O O HO

OH

OH

462 (epicochalasine B)

O

O

O

O O

O O HO

O

O

OH

HN O

OH OH O 460 (amichalasine C)

O

O

O

NH

O

O OH

OH

O

O O OH

O

O

NH O O H O

O

O O

O

OH

457 (amichalasine E)

459 (amichalasine B)

O

O O

O

HN O

O OH O OH

OH

461 (epicochalasine A)

HO

O

O

O 458 (amichalasine A)

OH

HO O HO

NH

O

NH O O

O

O OH O OH

O

OO O

OH

456 (amichalasine D)

O

OH

O

O

455 (asperchalasine A)

NH O O

O O O

HO

NH

NH

OO O

O OO

OH HO 454 (asperchalasine H)

NH OH

O

HO

OH

OH HO 453 (asperchalasine G)

O OO O

OH

O

O

HO

452 (asperchalasine F)

HO

OO

OH

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O

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HN OO

OH

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O

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O HN

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O

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OH O O O OH

O OH

HN

O

O

O O

O O OH

467 (asperflavipine B)

OH O O O HN O OH O O NH O HO

O

HN

HN

O OO OO

OH O OH

O OH OH OH

468 (asperflavipine A)

469 (dimericchalasine A)

Fig. 19 (continued)

centers, was elucidated unequivocally by extensive NMR spectroscopic and X-ray diffraction data analysis. Asperchalasine A (455) induced G1-phase cell cycle arrest in cancerous cells, but showed almost no negative effects on normal cells. Unsurprisingly, it has attracted considerable interest in terms of its organic synthesis and biosynthesis [2, 3, 198, 199] (for details, see Sect. 5). Continuing research has led to the isolation of two novel heterotrimers, epicochalasines A (461) and B (462) [196], from the same fungus (Aspergillus flavipes), using an OSMAC strategy with the liquid culture broth. Both compounds, containing an aspochalasin and two epicoccine units, exhibit a hendecacyclic 5/6/11/5/6/5/6/5/6/6/5 caged ring system, but their fusion patterns differ, resulting in two different carbon skeletons. Their absolute configurations were determined by both X-ray crystallographic and calculated ECD spectroscopic data analysis. Their biogenetic pathways were proposed to start from aspochalasin D (346), and to involve a Diels–Alder reaction and [3 + 2] cycloaddition as the key steps. Further studies on Aspergillus flavipes isolated from a soil sample led to the purification of the first cytochalasan heterotetramer, asperflavipine A (468) [4], which contains two cytochalasan moieties and two epicoccine moieties, uniquely defined by 5/6/11/5/6/5/6/5/6/5/5/11/6/5-fused tetradecacyclic rings. Then, aspergilasine A (468) [197], possessing an unexpected cyclobutane ring, together with three new biosynthetic intermediates of epicochalasine B (462), aspergilasines B–D (464–466), were isolated from the same fungus by using a liquid culture broth. Besides merocytochalasans from the fungus Aspergillus flavipes, Zhang and coworkers also discovered amichalasines A–C (458–460) [195] from Aspergillus micronesiensis PG-1, which represent a new type of cytochalasan heterotrimer. Amichalasines A (458) and B (459) possess an undecacyclic 5/6/11/5/5/6/6/5/11/6/5 ring system while amichalasine C (460) has an additional furan ring with a dodecacyclic 5/6/11/5/5/6/6/5/5/11/6/5 ring system. Amichalasines D (456) and E (457) exhibit a new type of cytochalasan heterotrimer, with a decacyclic 5/6/11/5/5/6/5/12/6/5 ring system, which were also reported recently from

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Aspergillus micronesiensis PG-1. Their structures were determined from the interpretation of the extensive spectroscopic data obtained and by single-crystal X-ray diffraction. Plausible biosynthesis pathways of amichalasines D (456) and E (457) were proposed involving a lactone aspochalasin unit. This is also the first report of merocytochalasans containing a [5.6.12] lactone aspochalasin unit. Dimericchalasine A (469) [194] is a unique aspochalasin homodimer fused with a C-20/C-20 single bond. This compound was also isolated from Aspergillus micronesiensis PG-1 along with other mero-aspochalasins. Its structure and absolute configuration were determined by means of X-ray crystallographic analysis. As far as is known by the present authors, it is also the first cytochalasan homodimer.

2.5 The Alachalasin Group The alachalasin group is a small group of cytochalasans (Table 20 and Fig. 20), with only seven members, namely, alachalasins A–G (470–476) [12]. Alachalasins A–G (470–476) were isolated from the cultures of Stachybotrys charatum by Che and coworkers in 2008. This type of cytochalasan contains an alanine residue, and therefore, C-10 these compounds is a methyl group without any substitution. Alachalasins F (475) and G (476) [12] possess an adenine moiety connected to C-19 of ring C, and thus, they could also be considered to be merocytochalasans.

2.6 Trichalasin From the culturing of Trichoderma gamsii isolated from the traditional Chinese medicinal plant P. notoginseng, Zhou and coworkers isolated a series of cytochalasans belonging to the aspochalasin group. Unexpectedly, an unusual cytochalasan named trichalasin A (477) (Table 20 and Fig. 20) [9] was also isolated, which represents Table 20 Alachalasins and trichalasin Compound

Origin

References

Alachalasin A (470)

Stachybotrys charatum

[12]

Alachalasin B (471)

Stachybotrys charatum

[12]

Alachalasin C (472)

Stachybotrys charatum

[12]

Alachalasin D (473)

Stachybotrys charatum

[12]

Alachalasin E (474)

Stachybotrys charatum

[12]

Alachalasin F (475)

Stachybotrys charatum

[12]

Alachalasin G (476)

Stachybotrys charatum

[12]

Trichalasin A (477)

Trichoderma gamsii

[9]

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OH

HN

OH

HN O O HO

470 (alachalasin A)

HN O O

O

O

O

O

472 (alachalasin C)

N

HN O O N

N

N

474 (alachalasin E)

O

N

O

473 (alachalasin D)

O

HN O O

O O HO

O

OH

HN O O

HN O O

O

471 (alachalasin B)

OH

HN

OH

O

N

O O O

OH

N

N

NH2

NH2

475 (alachalasin F)

476 (alachalasin G)

477 (trichalasin A)

Fig. 20 Structures of alachalasins and trichalasin

the first cytochalasan that originates biogenetically from valine. Since it features a unique valine-derived structure, and although there is only one compound of this type, it was assigned as the sixth group of the cytochalasan family.

2.7 Future Prospects As mentioned previously, there still remain some uncertainties about the structures of several cytochalasans, such as the planar structures of cytochalasin U Evidente (144) and cytochalasin V (124) and configurations of diaporthichalasin (192) and diaporthalasin (193). Moreover, there are also many cytochalasans with relative configurations that have not been determined completely, such as the merocytochalasans phomachalasins A–D (160–163) and MBJ-0038-MBJ-0040 (334–336). On the other hand, the discovery of the merocytochalasans as a group has greatly enriched the documented structural diversity of the cytochalasans, and it is recommended that further investigations on these compounds should be carried out in a strenuous manner. Thus, the identification of novel cytochalasans of diverse origin as well as the structural re-determinations of some already known cytochalasans are urgent matters to be addressed for both natural product chemists and organic chemists.

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3 Biological Activities of Cytochalasans To understand the biological activity of cytochalasans, first the cytoskeleton must be considered. The cytoskeleton is a network extending throughout a cell, which not only supports the cell, but also provides its shape, and organizes and tethers the organelles, and has roles in molecular transport, cell division, and cell signaling. A cytoskeleton occurs in all cells, including eukaryotic and prokaryotic cells, although the proteins that it is made of vary between organisms [200, 201]. Eukaryotic cells are complex cells that have a nucleus and organelles, and cells of plants, animals, fungi, and protists are all eukaryotic. The eukaryotic cytoskeleton consists of three types of filaments, which are elongated chains of proteins: microfilaments, intermediate filaments, and microtubules. Prokaryotic cells are less complex, with no true nucleus or organelles except for ribosomes, and they are found in the single-celled organisms, namely, bacteria and archaea. It was once thought that prokaryotic cells did not possess cytoskeletons, but advances in visualization technology and structure determination led to the discovery of filaments in these cells in the early 1990s [201]. A series of discoveries revealed that functional analogs of actin (MreB), tubulin (FtsZ) and intermediate filaments (crescentin) occur in prokaryotes. Moreover, it is now generally accepted that eukaryotic microtubules and actin filaments originate from these prokaryotic homologs [202]. Microfilaments are composed primarily of polymers of fibrous actin (F-actin) and the monomeric form of this protein, globular actin (G-actin), and these two forms exist in equilibrium in the cell. The microfilaments are present in bundles and form a three-dimensional (3D) intracellular meshwork. There is extensive intracellular binding and cross-linking with other intracellular proteins, such as myosin, lamin and spectrin [203]. Of the three types of protein fibers in the cytoskeleton, microfilaments are the narrowest, and they have a diameter of about 6 nm. The functions of microfilaments involve cell membrane motility, cytokinesis, endo- and exocytosis, secretion, vesicle transfer, cell shape maintenance, cell contractility, and affording mechanical stability. As mentioned earlier, cytochalasans are a group of structurally diverse fungal metabolites, and their name is derived from the Greek “kytos-chalasis”, meaning cell relaxation [204]. In a pioneering paper from 1972, actomyosin from rabbit muscle— the active protein complex of actin and myosin, was identified as a direct binding partner of cytochalasin B (110). Later on, studies by several groups revealed that cytochalasins B (110) and D (2) inhibit, but not completely arrest, actin filament elongation, and they specifically interact with the actin filament network by capping the barbed ends, and thereby altering the dynamic properties of microfilaments. In 2008, Trybus and coworkers achieved the crystallization of cytochalasin D (2) in a complex with actin and the exact binding relationship between cytochalasin and actin was clarified (Plate 3) [205, 206]. Thus, cytochalasans are known as microfilamenttargeting molecules, which exhibit a wide range of biological activities, by interfering with several cellular processes involving cytoskeleton formation.

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Plate 3 X-ray structure of the complex between cytochalasin (2) and actin (Protein Data Bank: 3EKU)

3.1 Cytotoxicity and Potential Anticancer Activities Cytotoxicity is the property of being toxic to cells, including inhibiting active growth and division, and inducing cellular apoptosis or necrosis. The first isolation of cytochalasans was guided by an observation that the crude extract of fungal metabolites had a cytotoxic action on cancer cells in vitro [207]. Subsequently, many researchers have found that cancer cells are more sensitive to cytochalasans than normal cells, because there is a substantial difference between the microfilaments of cancer and normal cells. These differences include alterations of actin polymerization and actin remodeling mediated by the activation of oncogenic actin signaling pathways (e.g., Ras and Src), or inactivation of several important actinbinding proteins that have tumor-suppressor functions [208]. Growing evidence has shown that these differences play a pivotal role in regulating the morphological and phenotypic events of cancer cells, which are related to key cancer characteristics, including altered adherence, anchorage-independent growth, invasiveness, and altered plasma membrane cytoskeletal interactions. Moreover, cytochalasans appear to damage preferentially malignant cells, as shown by their minimal effects on normal epithelial and immune cells. Thus, the cytotoxicity of cytochalasans to cancer cells and their potential as anticancer agents have attracted a great deal of attention and interest. Many studies have demonstrated the potent cytotoxicity of cytochalasans against panels of murine and human cancer cell lines [1]. In particular, the antineoplastic potentials of cytochalasins B (110) and D (2) have been investigated widely, and they have demonstrated promise using a variety of cell lines and mouse models, including murine Madison

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109 lung carcinoma and B16 melanoma. In addition, chaetoglobosins A–G (210– 216) and J (217) showed cytotoxicity against HeLa cells with IC 50 values ranging from 3 to 20 μg/cm3 [108], while chaetoglobosins A (210), B (211), D (212), J (217), Q (234), and T (236) as well as prochaetoglobosins I (221) and II (222) [115] displayed cytotoxicity against the P388 murine leukemia cell line [121]. Phomacins A–C (413, 414, and 348) from Phoma sp. exhibited inhibitory activities against HT29 colon adenocarcinoma cells, with IC 50 values of 0.6, 1.4, and 7.4 μg/cm3 , respectively [160]. Cytoglobosins C and D (238 and 239) each displayed inhibitory activity toward the A-549 cell line (IC 50 2.26 and 2.55 μM) [122]. Recently, many new cytochalasans having cytotoxic activities against cancer cells have been reported. Amichalasines A (458) and B (459), which represent a new type of cytochalasan heterotrimer, were isolated from Aspergillus micronesiensis PG1, and their IC 50 values determined against HL60 cells were 1.71 and 3.74 μM, respectively [195]. Asperflavipine A (468), a structurally complex merocytochalasan isolated from Aspergillus flavipes, was found to possess moderate cytotoxicity against seven cancer cell lines (MDA-MB-231, RKO, Hep-3B, HCT116, Jurkat, NB4 and HL60), with IC 50 values ranging from 12.7 to 27.6 μM [209]. Cytochathiazine B (338), which represents a new type of merocytochalasan isolated by the coculturing of Chaetomium globosum and Aspergillus flavipes, showed moderate antiproliferative activities against NB4 and HL-60 cells, with IC 50 values of 9.6 and 12.5 μM, respectively [154]. Flavichalasines N (424) and O (433), also isolated from Aspergillus flavipes PJ03-11, exhibited cytotoxic activities against three cancer cell lines (THP1, HL-60, and PC3), having IC 50 values ranging from 3.0 to 15.1 μM [187]. Similarly, cytoglobosins H (255) and I (256) showed antiproliferative effects for the LNCaP and B16F10 cell lines [126]. As the cytoskeletal structure acts at a specific time point in the cell cycle, the induction of cell cycle arrest at a specific checkpoint, and thereby inducing apoptosis, is a common mechanism for the cytotoxic effects of established anticancer drugs, as well as cytochalasans. Mechanically, cytochalasans initially alter the dynamic properties of microfilaments and inhibit cytokinesis of the cell without an effect on nuclear division; after this, normal cells enter the G0 resting state until sufficient actin levels have been attained for successful cytokinesis, while the fast-growing cancer cells typically continue to progress through the cell cycle, resulting in the formation of enlarged, multinucleated cells. A multinucleation phenomenon is observed almost exclusively in cancer cells, potentially representing a cancer-targeting attribute of cytochalasans [207]. It has been reported that many cytochalasins arrest the cell cycle and then induce apoptosis cell death. Treatment with cytochalasin D (2) led to arrest of the G1-to-S transition and induced apoptosis in cells retaining wild-type p53, while cells with inactivated p53 showed a partial rescue effect [210]. Cytochalasin B (110) induced G2/M phase cell cycle arrest, and activated ROS signaling and the Ca2+ -associated mitochondrial apoptotic pathways, using ZR-75-1 human breast cancer cells [211]. Chaetoglobosin A (210) showed preferential induction of apoptosis in chronic lymphocytic leukemia cells through targeting filamentous actin, and it thereby induces G2/M cell-cycle arrest [212]. Chaetoglobosin K (273) has been shown to inhibit cytokinesis, promote apoptosis, as well as to prevent

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oncogene-induced inhibition of gap junction-mediated cellular communication in ras-transformed epithelial and human lung carcinoma cells [213]. Chaetoglobosin K (273) also induced apoptosis and G2 cell cycle arrest through the p53-dependent pathway in cisplatin-resistant ovarian cancer cells [214], and it inhibited effectively tumor angiogenesis through downregulation of VEGF-binding HIF-1a in vivo [215]. Asperchalasine A (455) induced significant G1-phase cell cycle arrest by selectively inhibiting cyclin A, CDK2, and CDK6 in cancerous, but not normal cells, highlighting it as a potentially selective cell cycle regulator against cancer cells [193]. Epicochalasines A and B (461 and 462) both induced significant G2/M-phase cell cycle arrest and induced apoptosis in leukemia cells through the activation of caspase-3 and the degradation of PARP [196]. Moreover, the unique microfilament-directed mechanisms of cytochalasans have made them of interest for drug synergy investigations with existing drug therapy for treating cancer. Early studies have shown drug synergy between cytochalasin B (110) and cytarabine as well as vincristine. These have provided compelling evidence that cytochalasin B (110) and its reduced congener may have clinically applicable synergistic potential [216]. More specifically, concomitant administration of cytochalasin B (110) and vincristine substantially reduced the clonogenicity potential of U937 human monocytic leukemia cells, thereby potently inhibiting their propensity to proliferate [217]. Both cytochalasin B (110) and its congener 21,22-dihydrocytochalasin B (DiHCB) demonstrated considerable drug synergy with doxorubicin (ADR) against ADR-resistant P388/ADR leukemia cells in vitro. In turn, cytochalasins B (110) and D (2) substantially increased the life expectancy of mice challenged with P388/S and P388/ADR leukemia xenografts in vivo [218]. Further, due to cytokinesis inhibition potentiated by the cytochalasans, normal or multidrugresistant neoplastic cells become enlarged and multinucleated, which makes them suitable targets for microtubule-directed (paclitaxel), nucleic acid-directed (doxorubicin) agents, or other physicochemical treatment modalities, such as experimental sonodynamic therapy (SDT), an ultrasound therapy that preferentially damages cells based on their size. For example, pulsed low-frequency ultrasound in the 20–40 kHz range is able to induce preferential destruction of neoplastic cells enlarged by treatment with cytoskeletal-directed agents [217, 219]. The most well-known mechanism of action is the concomitant perturbation of the cell cycle via microtubule-targeting G2/M arrest and microfilament-targeting cytokinesis inhibition. For multidrugresistant neoplastic cells, Trendowski et al., using the multidrug-resistant SKVLB1 cell line as a model, demonstrated that cytochalasans appear to inhibit the activity of P-glycoprotein (P-gp) and potentially other ABC transporters, and may have a novel type of activity against multidrug-resistant neoplastic cells that overexpress drug efflux proteins [220]. In conclusion, the actin cytoskeleton is vital for carcinogenesis and subsequent pathology, and microfilament-targeting cytochalasans have raised considerable interest as cancer-related drug candidates [160, 221]. When used as a single agent, the narrow therapeutic index of cytochalasin derivatives prevents their clinical use, and to date no microfilament-directed agent has entered a clinical trial [222].

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However, there is no denying that cytochalasans offer novel mechanisms for exploitation in cancer therapy that may improve the efficacy of treatments against cancers refractory to standard chemotherapeutic protocols. In the near future, it is possible that cytochalasans could be used to supplement current chemotherapeutic measures to improve efficacy rates, as well as to decrease the prevalence of drug resistance in the clinical setting.

3.2 Antimicrobial Activities In addition to their potential antineoplastic properties, cytochalasans also have been found to possess considerable antimicrobial effects. The antimicrobial activities of cytochalasins A (109) and D (2) were disclosed in the first report on the biological characteristics of cytochalasans. Thus, cytochalasin A (109) was found to inhibit the growth of Bacillus subtilis and Escherichia coli, as well as to exert antifungal activity against Botrytis cinerea. In contrast, cytochalasin B (110) did not show any antifungal activity against the test organisms and cytochalasin D (2) was only reported to have antimycotic activity. Moreover, other cytochalasans such as diaporthichalasin (192, phomopsichalasin), scoparasin B (205) and chaetoglobosins A (210) and C (212), isolated from endophytic Chaetomium globosum, have been reported to have antimicrobial activity [223–225]. Diaporthichalasin (192, phomopsichalasin) isolated from a culture of an endophytic Phomopsis species also has antimicrobial effects [90]; this compound, in which the common macrocycle is replaced by a 13-membered tricyclic system, showed antibacterial activity in disk diffusion assays against Bacillus subtilis, Salmonella gallinarum and Staphylococcus aureus, as well as antifungal activity against the yeast Candida tropicalis. Scoparasin B (205), a cytochalasin isolated from the culture broth of the endophytic fungus Eutypella scoparia PSU-D44, was reported have antifungal activity against the dermatophyte Microsporum gypseum SH-MU-4 (MIC 30.3 μM) [101]. Chaetoglobosins A (210) and B (211), inhibited the growth of Staphylococcus aureus, methicillin-resistant S. aureus, and Mycobacterium tuberculosis H37Ra [226]. Further, Aouiche and coworkers reported that the MIC values for chaetoglobosin A (210) were between 30 and 75 μg/cm3 for yeasts, 50 and 75 μg/cm3 for filamentous fungi, and 20 and 30 μg/cm3 for Gram-positive bacteria [13]. Also, several cytochalasans have shown significant inhibition of Staphylococcus aureus biofilm formation at subtoxic levels, including cytochalasins A (109) and C (1), chaetoglobosin A (210), 19,20-epoxycytochalasin C (52) and L-696,474 (26). Among them, chaetoglobosin A was the most potent, as it inhibited 70–91% of biofilm formation in Staphylococcus aureus [96]. Studies also provided insights into the mechanism of action and structure–activity relationships of the antimicrobial activities of cytochalasans. For example, cytochalasin A (109) is a bacteriostatic agent that inhibits a variety of physiological processes in Gram-positive bacteria, such as enzyme induction, transportation, and respiration of exogenous substrates. The presence of an α,β-unsaturated carbonyl group in the

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macrolide moiety and an unsubstituted lactam ring proved to be a prerequisite for activity against Gram-positive bacteria. The presence of a C-7 hydroxy group, which has been proposed as a pharmacophore for activity in eukaryotes, does not appear to be necessary for bacterial transport antagonism.

3.3 Antiparasitic Activities Actin is also the major protein constituent in cells of parasites, and since actin filaments play a pivotal role in the differentiation and infection processes, the effects of cytochalasans on parasites have been studied widely. The influence of various cytochalasans on the growth and differentiation of Entamoeba invadens, a reptile parasite used as a model system to study such processes in human parasites that cause amebiasis, was investigated by Makioka and coworkers. At a 10 μM concentration level, cytochalasins B (110), D (2), E (145) and dihydrocytochalasin B significantly inhibited the growth of the parasite. Furthermore, at 1 μM, cytochalasin D (2) was found to inhibit cyst wall formation, which enables parasites at the worm stage to survive inside or outside (encystation) the host, whereas cytochalasins B (110) and E (145) and dihydrocytochalasin B were active only at 10 μM. Cytochalasins D (2) and E (145) also affect excystation [227]. Mori and coworkers screened 431 compounds and 6,900 samples of microbial broth extracts, and identified aspochalasin B (344), chaetoglobosin A (210) and prochaetoglobosin III (223) with l-cysteine-dependent antiamebic activity at 100 μg/cm3 [228]. Cytochalasin A (110) showed LC 50 values of 0.0854 ± 0.0019 μg/mm3 against Aedes aegypti larvae, which is the major vector of the arboviruses responsible for Dengue fever, one of the most devastating human infectious diseases [229]. Cytochalasans also showed antiparasitic activities on plant parasites. Meloidogyne spp., one of the most important groups of plant-parasitic nematodes, has caused great damage to a wide range of crops worldwide. Hu and coworkers studied the nematocidal activity of Chaetomium globosum NK102, culture filtrates, and purified chaetoglobosin A (210). The results showed that C. globosum NK102 significantly repelled second-stage juveniles (J2s). Both the culture filtrates and 210 demonstrated strong adverse effects on J2 mortality with 99.8% at 300 μg/cm3 (LC 50 = 77.0 μg/cm3 ) at 72 h. Compound 210 and the culture filtrates did not affect egg hatching until 72 h of exposure. All filtrate treatments inhibited the penetration of J2 even at a 12.5% dilution treatment. Similarly, 210 (300 and 30 μg/cm3 ) showed significant inhibitory effects on J2 penetration. The number of eggs per plant was significantly reduced using a treatment of 30 mg/kg soil, by 63% relative to control plants, indicating its apparent negative effect on reproduction of M. incognita. This study demonstrated the nematocidal activity of 210 and suggested that it could be a potential biocontrol agent for the integrated management of Meloidogyne incognita [230]. The nematicidal activity of 210 may be due to its inhibition of the polymerization of monomeric actin (G-actin) to polymeric form (F-actin), and thereby inhibit cell functions requiring cytoplasmic microfilaments.

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3.4 Anti-inflammatory Effects The anti-inflammatory effects of cytochalasans also have been reported. Chaetoglobosin Fex (227), isolated from the marine-derived endophytic fungus Chaetomium globosum QEN-14, inhibited significant LPS-induced inflammatory mediator production both in the RAW264.7 cell line and in mouse peritoneal macrophages. Its anti-inflammatory property was explained by inhibiting NF-κB and MAPKs activation in LPS-stimulated macrophages, and by blocking membrane-associated CD14 (mCD14) expression [231]. Compound 227 also exhibited an immunosuppressive effect on mouse bone marrow-derived DCs (BMDCs) via TLR3 signaling, which suggested its potential application in the treatment of autoimmune inflammatory diseases [232]. Chaetoglobosin F (215), obtained from an EtOAc extract of a solid culture of Chaetomium globosum IFB-E019, also showed anti-inflammatory effects on BMDCs. Compound 215 inhibited the CpG-induced DC maturation and function by suppressing the expression of surface molecules (CD40, CD80, CD86 and MHCII), reducing the production of cytokines and chemokines (IL-12 and CXCL-10), inhibiting CpG-induced DCs-elicited allogeneic T-cell proliferation, and impairing the migration ability of chemokines. Additionally, compound 215 inhibited CpGinduced-activation of MAPKs (p38 and JNK, but not ERK) and the nuclear translocation of NF-κB and STAT1. Furthermore, 215 was able to suppress TLR9 expression of CpG-induced DCs. Collectively, these immunosuppressive properties of 215 may prove useful in controlling DC-associated autoimmune and/or inflammatory diseases [233].

3.5 Antiviral Activities The antiviral activities of cytochalasin derivatives also have been reported. Thus, HIV-1 protease plays a crucial role in viral maturation: its inhibition results in the reduced spread of infectious viruses. Goetz and coworkers isolated a novel cytochalasin L-696,474 (26), and found it to be a competitive inhibitor of HIV-1 protease that did not possess any apparent cytotoxicity [14]. In 2004, 19,20-epoxycytochalasin Q Edwards (31) was found to be an antagonist of the chemokine receptor CCR5, which is implicated in HIV infection [234]. Aspochalasin L (354), isolated from a fermentation broth of Aspergillus flavipes, has been reported with activity against HIV integrase, with an IC 50 of 71.7 μM [165]. Che and coworkers reported that alachalasin A (471), with an unusual adenine substituent isolated from Podospora vesticola, displayed an inhibitory effect on HIV-1 replication in C8166 cells [12]. In 2015, Zhang and coworkers reported that armochaetoglobins L (302), M (303), N (304), Q (307), R (308), and penochalasin B (298), as isolated from a culture broth of Chaetomium globosum TW1-1, showed significant anti-HIV activities, with EC 50 values ranging from 0.11 to 0.55 μM [144]. These results suggest that certain cytochalasans could be of interest for development as anti-HIV agents. However,

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research studies on the antiviral activities of cytochalasans have been carried out in vitro, and there are no in vivo data available as yet. Moreover, the exact mechanisms involved in the antiviral activities of the cytochalasans are not yet understood. The effects on HIV-1 protease, integrase, and CCR5 imply that there seems to be a mechanism or mechanisms of action independent of microfilaments, but there is a lack of in-depth research on this area and no definitive conclusions can be drawn. Considering that some research has been conducted in cells infected with a virus, it may be surmised that the microfilament-targeting effects of cytochalasans still play an important role in their antiviral activities: they may affect microfilaments of the host cells and then interfere with the process of the virus infecting the host cells, and thereby inhibit the activity of virus-related enzymes and hence overall viral replication.

3.6 Phytotoxic Effects and Ecological Role Early studies demonstrated that several plant cells are susceptible to cytochalasan treatment. The addition of cytochalasin B (110) to the pollen tubes of Lilium longiflorum resulted in immediate cessation of cytoplasmic streaming and eventually led to inhibition of tip growth. Similar effects were observed in Caulerpa prolifera rhizoids and Acetabularia mediterranea [1]. Then, researchers discovered that low-molecularweight compounds secreted from phytopathogenic fungi will weaken or kill the plant, inclusive of cytochalasans. In particular, the genus Phoma includes several pathogenic representatives, which are responsible for plant diseases with characteristic symptoms, such as lesions on leaves, stems, blossoms, and pods. Among other secondary metabolites, cytochalasans have been proposed as mediators of virulence. Several cytochalasans have been isolated from plant-pathogenic fungi. Cytochalasin A (109) and B (110) can be isolated from Phoma exigua var. exigua, the causative agent of potato gangrene. Evidente and coworkers purified several cytochalasans from a culture of Phoma exigua var. heteromorpha grown on wheat kernels and determined their phytotoxic effects on germinating tomato seedlings [75]. Later on, cytochalasins B (110), F (111), Z2 (115), Z3 (116), and desoxaphomin (104) were found to be produced by Phoma exigua var. exigua isolated from Cirsium arvense (Canada thistle or creeping thistle) and Sonchus arvensis (perennial sowthistle), and their phytotoxic activity against both plants was investigated. The results suggested that a carbocyclic or a lactone macrocycle fused to an unaltered perhydroisoindolyl residue as well as a secondary hydroxy group at C-7 are essential to confer biological activity [88]. Targeting the early stages of weed growth is a strategy of herbicidal development that may be applied to natural metabolites of microbial and plant origin. Cytochalasins E (145) and B (110) were tested on Striga seed germination, and the results showed only cytochalasin E (145) to inhibit Striga germination by 50%, while cytochalasin B (110) had no inhibitory effects. The herbicidal effects mediated by cytochalasins A (109) and B (110) in inducing necrosis in the tissues of weeds

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also have been reported [235]. In 2014, Cimmino and coworkers found that cytochalasans inhibited the germination of GR24-treated broomrape seeds. The growth of broomrape radicles was potently inhibited by compounds belonging to the cytochalasan class. Broomrape radicles treated with epi-sphaeropsidone developed a layer of papillae while radicles treated with cytochalasans turned necrotic. These findings enabled potential new natural herbicides to be identified for the management of parasitic weeds [236]. The herbicidal activity of cytochalasans against selected plant species suggests their possible use as biocontrol agents for weed management. The production of secondary metabolites enables an organism to compete with other organisms in its natural environment. In this case, the production of a toxins such as the cytochalasans might play a major role in deterring predators or competitors. It is believed that endophytes living inside plant tissues obtain nutrition and protection from the host. In return, they may enhance the fitness of the host plant by producing certain functional metabolites [237]. Thus, the secretion of antimicrobial cytochalasans might foster the close association of the fungus and the host plant by protecting the plant from herbivores or plant pathogens, thus maintaining the delicate equilibrium between the antagonism of a host and a fungus. A similar ecological function might be inferred for cytochalasin F (111), isolated from an endophyte of Teucrium scorodonia that was shown to possess the algicidal activity and to inhibit photosynthesis.

3.7 Future Prospects Cytochalasans have a wide range of distinctive biological activities. The most welldocumented one is their influence on cellular processes based on interference with the microfilament network formation. Therefore, they have become useful compounds for the investigation and understanding of microfilament-involved cellular processes like cell division, multinucleation, and migration. As a result of their effects on such cellular processes, many cytochalasans also exhibit a range of cytotoxic properties and evidence of antineoplastic effects, as well as antimicrobial, antiparasitic, antiviral, and anti-inflammatory activities. Therefore, selected cytochalasans potentially could be developed into drugs. Among these compounds, cancer-related activity is the most promising. Many cytochalasans exhibit cancer cell inhibitory activity, when used either alone or when combined with established cancer chemotherapeutic drugs, as indicated by numerous laboratory studies conducted over the last two decades. Thus, more focus should be put on the antineoplastic investigations of cytochalasans in the future, not only limited to compounds that have already been well studied such as cytochalasin B (110) and D (2), but also less well-known compounds such as amichalasines A (109) and B (110). In addition, research on cellular targets and the exact mechanisms of action by cytochalasans are areas that need to be strengthened in the future. Previous studies have shown that some cytochalasans inhibit cholesterol synthesis, interfere with glucose transport by binding to high-affinity sites on glucose-transporter proteins,

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Plate 4 X-ray structure of human glucose transporter hGLUT1 bound to cytochalasin B (110) (Protein Data Bank: 5EQI)

and inhibit thyroid secretion or release of growth hormones, all of which do not seem to be directly related to actin-binding [1]. In 2016, the crystal structure of the human glucose transporter hGLUT1 bound to cytochalasin B (110) was determined (Plate 4) [238], which also indicated a cellular target that is not just a microfilament. These findings suggested that more potential mechanisms of cytochalasans need to be explored. Furthermore, with the application of new techniques such as highthroughput screening and structural pharmacology, it is possible to use trace amounts of newly discovered natural cytochalasans to screen a wider range of biological activities, and to study their mechanism(s) of action in detail. Therefore, in the near future, more extensive screening and more in-depth mechanistic work are required to study the biological properties of the cytochalasans.

4 Biosynthesis of Cytochalasans Since the 1970s, the biosynthesis of cytochalasans has been studied intensively. Due to a lack of knowledge on the genetic backgrounds of organisms producing cytochalasans, only limited approaches, such as feeding experiments, could be applied to investigate biosynthesis pathways. By feeding with isotope-labeled precursors, such as acetate, malonate, methionine, and various amino acids, as well as 18 O2 gas during strain cultivation, some biosynthesis pathways were proposed initially. These suggested the formation of cytochalasan skeletons through mixed biosynthesis origins [239–245]. Further structural tailoring has been introduced using P450 enzymes and Baeyer–Villiger enzymes, which insert oxygen into the cytochalasan backbone [115, 240, 246, 247]. The newer approaches of genome sequencing and molecular biology techniques, such as the polymerase chain reaction (PCR) and genetic recombination, have led biosynthesis studies on cytochalasans to new levels of understanding and have revealed mechanisms at the molecular level. In the past decade, several cytochalasan biosynthesis gene clusters have been identified, of which some have been characterized functionally [2].

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4.1 Cytochalasan Gene Clusters The first cytochalasan gene cluster reported in 2004 was ACE1 identified from Magnaporthe oryzae Guy11 (named earlier Magnaporthe grisea), and since then has been further investigated [248–250]. The ACE1 gene cluster was speculated to confer a virulence factor that is recognized by rice cultivars carrying the resistance gene Pi33, enabling the rice to activate a defensive response. This cluster encodes several biosynthesis proteins, which were annotated as two PKS–NRPS (ACE1 and SYN2), two trans-ER proteins (RAP1 and RAP2), four P450 monooxygenases (CYP1-4), two FAD-dependent monooxygenases (OXR1 and OXR2), an α,β-hydrolase (ORFZ), a putative Diels–Alderase (pDA, ORF3), a transporter (MFS) and a regulator (BC2) (Fig. 21A). Investigation into the evolution of the ACE1 gene cluster by Wolfe and co-workers indicated that it is distributed sporadically and shared by only a few fungal species [250]. The evolution of the ACE1-like gene clusters is characterized by complex events including gene duplication and losses, gene recruitment, and horizontal gene transfer [250]. Phylogenetic analysis has indicated that the ACE1 cluster is related to the gene clusters in Chaetomium globosum and Aspergillus clavatus, which were later shown to encode cytochalasan molecules. It was also suggested that in Magnaporthe oryza, the physical linkage between the three core genes, PKS-NRPS (ACE1), transER (RAP1) and pDA (ORF3) e, is ancient, and can be inferred to have existed in a common ancestor [250]. Although the secondary metabolite(s) from the ACE1 gene cluster has not yet been isolated, due to their tightly regulated temporal expression, recent work by two independent groups suggests that the ACE1 gene cluster encodes the biosynthesis of two different cytochalasan compounds [251, 252]. The first cytochalasan gene cluster to be characterized was the chaetoglobosin A (208) (che) cluster (Fig. 21B) from Penicillium expansum [253]. Before this work, numerous isotope labeling experiments suggested that cytochalasan biosynthesis involves the formation of acetate and methionine-derived octa- or nonaketide chain and the attachment of an amino acid, which was referred to as the PKS-NRPS pathway in the biosynthesis of other secondary metabolites such as fusarin [254], equisetin [255], tenellin [256], and aspyridone [257]. In order to discern the molecular basis of cytochalasan biosynthesis, Hertweck and coworkers used a heterologous probe spanning the C-methyltransferase (CMeT), ketoreductase (KR), and acyl carrier protein (ACP) domain coding regions from the Fusarium venenatum fusarin C PKS–NRPS, to screen a Penicillium expansum cosmid library. As a result, the che gene cluster consisted of seven genes encoding a PKS–NRPS (CheA), a trans enoylreductase (CheB) [258], two putative P450 monooxygenases (CheD and CheG), a FAD-dependent monooxygenase (CheE), and two transcription factors (CheC and CheF). Bioinformatic analysis indicated that the gene cluster was most similar to the aspyridone cluster in A. nidulans. As all attempts to generate a targeted gene knockout in P. expansum and heterologous expression of the entire or partial gene cluster in Aspergillus failed, the cluster was confirmed putatively, using RNA-mediated gene

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Fig. 21 Comparison of characterized and cryptic cytochalasan gene clusters. Abbreviations: pDA = putative Diels-Alderase, trans-ER = trans enoyl reductase, OXR = oxidoreductase/monooxygenase, CYP = cytochrome P450, BMVO = Baeyer–Villiger monooxygenase, OMT = O-methyltransferase, OAT = O-acetyltransferase, TF = transcription, factor/transcriptional regulator, MFS = transporter

silencing of CheA, which resulted in the production of chaetoglobosin A (208) and chaetoglobosin C (210) being nearly entirely abolished. In the fungus Chaetomium globosum, the second gene cluster for chaetoglobosin A (208) biosynthesis was characterized (Fig. 21C) [259]. By means of BLASTP of the previously characterized hybrid proteins CheA in the genome of C. globosum, a homologous PKS–NRPS protein encoded by CHGG_01239 was identified, which showed an amino acid sequence identity and similarity of 34.3% and 52.2% to CheA, respectively. Neighboring this PKS-NRPS was a trans-ER encoded by the gene CHGG_01240. Similar to P. expansum, the chaetoglobosin A (208) biosynthetic gene

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cluster in C. globosum also contained several tailoring enzymes such as two P450 oxygenases (CHGG_01242-1 and CHGG_01243), a FAD-dependent monooxygenase (CHGG_1242-2), as well as a transcription factor (CHGG_01237). However, two additional genes encoding proteins of unknown function CHGG_01241 and CHGG_01244 were identified. The latter encoded a protein with sequence similarity to α,β-hydrolases. Targeted gene deletion of CHGG_01239 and CHGG_01240 led to the abrogation of chaetoglobosin A (208) production in C. globosum, confirming the function of the cluster. The biosynthesis of cytochalasin E (139) and cytochalasin K (140) in Aspergillus clavatus has been determined as being achieved by gene cluster ccs (Fig. 21D) [260]. The involvement of PKS–NRPS encoded by the gene ccsA in ccs cluster was confirmed by gene disruption. Bioinformatic analysis of this cluster revealed a transER (CcsC), two P450 monooxygenases (CcsD and CcsG), a transcriptional regulator (CcsR), two proteins of unknown function (CcsE and CcsF), as well as a unique Baeyer–Villiger monooxygenase (BVMO, CcsB), which could catalyze previously proposed reactions, so that an oxygen atom may be inserted in the backbone forming the vinyl carbonate moiety in cytochalasin E (139) [240]. CcsE shows a sequence similarity to an α,β-hydrolase and shares a 54% identity with CHGG_01244, while CcsF shares a 51% sequence identity with CHGG_01241 from C. globosum. CcsF has previously been suggested to be involved in the intramolecular [4 + 2] Diels–Alder cycloaddition reaction, so for simplicity, CcsF and its homologs in other cytochalasan gene clusters, for example CHGG_01241 shall be referred to as putative Diels– Alderases (pDA) [2]. The pyrichalasin H (164) gene cluster, pyi, was identified through genome mining in Magnaporthe grisea (Fig. 21E) [8]. The function of this cluster was confirmed by disruption of the PKS–NRPS encoding gene pyiS, which resulted in the complete abolishment of pyrichalasin H (164) production. An investigation of the cluster pyi revealed that it has a high similarity to the ACE1 gene cluster in M. oryzae Guy11 [249]. Besides the PKS–NRPS, cluster pyi contains genes encoding a trans-ER (pyiC), α,β-hydrolases (pyiE), a pDA (pyiF), two P450 monooxygenases (pyiD and pyiG), an oxidoreductase (pyiH), an O-methyltransferase (pyiA), an Oacetyltransferase (pyiB), a transporter (pyiT ) and a transcriptional regulator (pyiR), respectively. On comparing to other characterized cytochalasan gene clusters, two additional tailoring genes, pyiA and pyiB, were characterized as being involved in the formation of the para-methoxyphenyl and O-acetyl groups in pyrichalasin H (164), respectively [8]. Genome sequencing of the fungus, Hypoxylon fragiforme MUCL 51264, which produces cytochalasin H (7) and several analogs, revealed the gene cluster hff [55]. Compared to the pyi cluster in M. grisea NI980 that encodes cytochalasin H (7), homologous biosynthesis genes encoding a PKS-NRPS (hffS), a trans-ER (hffC), a putative Diels–Alderase (hffF), an α,β-hydrolase (hffE), two P450s (hffD and hffG), an oxido-reductase (hffH), an acetyltransferase (hffB), a transporter (hffT ), and a transcription factor (hffR) were identified in cluster hff (Fig. 21F). Although a systematic functional investigation of cluster hff has not yet been reported, two P450 genes (hffD and hffG) in this gene cluster were confirmed to have the catalytic abilities to restore

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the production of cytochalasin H (7) in M. grisea P450 knockout strains (ΔpyiD and ΔpyiG) in a combinatorial biosynthesis study [55]. The above information supports the evolution analysis of ACE cluster in M. oryzae conducted by Wolfe and Bradshaw’s groups, which suggested four genes, the PKSNRPS, the trans-ER, the pDA and the α,β-hydrolase gene, as the core set of genes (Fig. 21) involved in the production of a group of secondary metabolites that are modified differentially by other accessory enzymes [250, 261]. Using these core genes, several ACE1 clusters homologated cryptic gene clusters were discovered in other filamentous fungi (Fig. 21g), although the metabolites they encode remain unknown [2, 250, 261].

4.2 Investigations into the Biosynthesis of Cytochalasans Before the characterization of the che gene cluster in P. expansum, there was hardly any information available on the molecular basis of cytochalasan biosynthesis. Even at the present time, the function of hybrid PKS-NRPS in fungi [254–257], which play a core role in cytochalasan clusters, has not been known very long. Using RNA-mediated gene silencing experiments, Hertweck and coworkers characterized the functions of two core genes (cheA and cheB) in the che cluster and proposed a cytochalasan biosynthetic pathway, in that the PKS part of the hybrid synthase CheA and the enoyl reductase CheB act together to synthesize a polyketide chain. Subsequently, this is passed on to the NRPS and condensed with an activated tryptophan [253], and then catalyzed by a C-terminal reductase domain in CheA, with a reductive off-loading then taking place to release an aminoaldehyde intermediate that can further form a pyrrolinone via a Knoevenagel condensation. This pyrrolinone can function as a dienophile, and could undergo intramolecular Diels–Alder (DA) cyclization with the polyketide terminal diene to generate the isoindolone-fused macrocycle (Scheme 1). The release mechanism and the formation of the isoindolonefused macrocycle was also proposed in an alternative way that a functional domain within the PKS–NRPS acted as a Dieckmann cyclase, as observed in (aspyr-)idone and tenellin biosynthesis [256, 257], and directly releases a tetramic acid derivative [253], which further undergoes a series of subsequent reactions such as dehydration, intramolecular DA cyclization and reduction. The roles of the oxidases CheD, CheE and CheG in this cluster were proposed as modifying the cytochalasan backbones to yield chaetoglobosins A (210) and C (212) (Scheme 1). To date, several cytochalasan gene clusters have been identified, and the individual steps to form the cytochalasan backbones and structural diversity were partially characterized. As described in a previous review by Skellam [2], the biosynthesis of cytochalasans may be divided into three stages: the early stages, the formation of the linear polyketide conjunct pyrrolinone; the middle stages, the formation of the macrocycle fused isoindolone via DA cyclization, and the late stages that lead to modifications on the cytochalasan framework.

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Scheme 1 The proposed biosynthesis of chaetoglobosin A

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The Early Stages of Cytochalasan Biosynthesis

To validate the early formation mechanism of cytochalasan, an exploratory study on the biosynthesis of cytochalasin E (145) was conducted by Oikawa and co-workers. They co-expressed heterologously the genes encoding the PKS–NRPS, ccsA, and the trans-ER, ccsC from A. clavatus together in Aspergillus oryzae [262]. Unexpectedly, neither the supposed aldehyde nor pyrrolinone structure was obtained when these genes were overexpressed, instead, an octaketide alcohol A1 intermediate was isolated and identified (Scheme 2a). Another unexpected feature was the absence of an olefin between C-4 and C-5 on the polyketide chain, which is observed in cytochalasin E (145), suggesting a reduction reaction catalyzed by an enzyme within A. oryzae [2] might take place on the backbone. A similar type of investigation has been conducted by Larsen and coworkers, who used Aspergillus nidulans as the heterologous expression host to characterize the functions of ccsA and ccsC [252]. They found that the expression of ccsA alone did not yield any detectable products, and the expression of trans-ER ccsC along with ccsA was vital for the product formation. The A. nidulans strain co-expressed ccsA and ccsC and produced several products, of which a minor one was speculated to be identical to the previously discovered intermediate A1 by Oikawa’s group, as the same compound mass at 440.3188 Da was measured. Differing from A1, a linear PKS-NRPS precursor, the major product named niduclavin (A2) (Scheme 2A), possesses a phenylalanine moiety joining a decalin scaffold that originates from a highly reduced octaketide chain. In comparison to the general structure of cytochalasans such as cytochalasin E (145), niduclavin (A2) has an additional double bond between the C-2 and C-3 positions of the phenylalanine side chain. The authors speculated that cross-chemical reactions with endogenous enzymes from A. nidulans were responsible for the introduction of the double bond, which would additionally activate the dienophile in the α/β/-position (C-4 and C-5) of the C-3 carbonyl group, thereby possibly favoring decalin formation rather than the tetramic acid moiety present in the native A. clavatus molecule. Inactivation of a dioxygenase AsqJ, suspected to be responsible for the introduction of the C-2 /C-3 double bond, did not prevent the formation of niduclavin (A2). The genes CcsA and ccsC had also been expressed in an alternate heterologous host, Aspergillus niger. Co-expression of these two genes in A. niger also led to the production of niduclavin (A2) [252]. As the previous basic local alignment search tool research (BLAST) revealed a high degree of similarity between CcsA and the SYN2 in the ACE1 cluster (68% similarity and 52% identity). Larsen’s group studied the cryptic genes SYN2 and RAP2 present in the ACE1 gene cluster (Fig. 21A) from M. oryzae in the same study [252]. Analogous to the integration and expression of ccsA and ccsC, SYN2 and RAP2 were transformed into A. nidulans. The SYN2/RAP2 product was purified and a structure with a close resemblance to niduclavin A2 was determined by NMR spectroscopy and given the name niduporthin A3 (Scheme 2B). Key differences between niduporthin A3 and niduclavin A2 include incorporation of different amino acid and methyl group substitutions on the polyketide chain.

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$ SEnz 1 ×acetyl-CoA 8 ×Mal-CoA 3 ×SAM 1 ×Phe-AMP

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Scheme 2 The outcome of investigations into the early stages of cytochalasan biosynthesis (A–C)

The remaining cryptic gene pair ACE1/RAP1 in the ACE1 gene cluster from M. oryzae has been investigated by Cox and co-workers [251]. Expression of ACE1 alone in A. oryzae produced a polyenyl pyrone A4, which was believed to be a shunt product since it lacked the expected amino acid moiety (Scheme 2C). However, coexpression of ACE1 and RAP1, an amide A5, closely resembling A1 from A. oryzae co-expressed with ccsA and ccsC, was isolated (Scheme 2C).

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In conclusion, the above investigations revealed that the early stages of cytochalasan biosynthesis depend on the co-expression of both PKS–NRPS and its trans-ER. In the heterologous host A. oryzae, both of the co-expression products, amides A1 and A5, possess a hydroxy group moiety rather than the expected formyl group, and suggested that an unwanted reduction occurred during the backbone formation. It remains to be seen whether this excessive reduction was caused by native enzymes or by the alternative splicing of the PKS–NRPS functional domain in the heterologous host. It has been observed that the co-expression of ccsA and ccsC in different hosts led to two distinct intermediates, A1 and A2, respectively. A proposal made by Skellam [2] is that native accessory enzymes in A. oryzae reduce the proposed aldehyde but in A. nidulans and A. niger drive the formation of the pyrrolinone unit. These modifications by the heterologous host may prevent the formation of the required pyrrolinone and thus prevent also the [4 + 2] Diels–Alder cycloaddition reaction occurring with the terminal diene [2].

4.2.2

The Middle Stage of Cytochalasan Biosynthesis

The middle stage of cytochalasan biosynthesis is defined as the conversion from the linear PKS–NRPS intermediate to the iso-indolone fused macrocyclic structure. Although there is only one intramolecular [4 + 2] Diels–Alder (DA) cycloaddition involved, it is a critical step in the formation of the cytochalasan skeleton and a step that has intrigued researchers for many years. It is believed that the DA reaction is catalyzed by specific enzymes and several pieces of indirect evidence have been provided through well-designed experiments. Earlier evidence came from the total synthesis of cytochalasans, as reviewed by Thomas in 1991 [263]. As a key step in the synthesis of proxiphomin (102), a naturally occurring cytochalasan, the intramolecular DA reaction of B1 required relatively rigorous conditions (100 °C for 5 h) to give a mixture of B2 and B3, suggesting that spontaneous cycloaddition in fungal cells was unlikely (Scheme 3A). In 1992, Oikawa et al. demonstrated that the stereospecificity of the cytochalasan DA reaction might require enzymatic stereo-control [264]. As shown in Scheme 3B, after the pyrolysis of prochaetoglobosin I (221), the authors found that both diastereomers 221 and diastereo-221 existed in the product mixture. From these results, it could be speculated that 221 first undergoes a reverse DA reaction to form a linear intermediate B4, namely, the proposed substrate for the DA reaction. As it cyclized spontaneously under the reaction conditions used, an equal amount of diastereomers were produced. The lack of stereoselectivity in this reaction indicated that the enzyme responsible for [4 + 2] cycloaddition stabilizes the endo transition state and results exclusively in the production of prochaetoglobosin I (221). In addition to the above evidence from chemical reactions, progress made in genetic engineering on fungi in recent years has accelerated the pace of searching for DA enzymes. The formation of decalin, which involves a DA reaction similar to that involved in producing the cytochalasans, was proved to be catalyzed by the enzyme MycB by the group of Tang [265] (Scheme 3C). Highlights of this study

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Scheme 3 Previous studies on Diels–Alder (DA) cycloadditions. a A chemical synthesis procedure to achieve intramolecular DA reactions. b The retro-Diels–Alder reaction of prochaetoglobosin I (221). c MycB was proven to be a Diels–Alderase to form decalin through in vitro characterization. d A gene knockout experiment of pyiF afforded multiple linear shunt products B10–B17, giving indirect evidence that PyiF is the Diels–Alderase in the biosynthesis of pyrichalasin H (164)

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were that, in addition to the in vivo experiments proving that MycB is related to DA, in vitro enzymatic assays also gave direct evidence that MycB catalyzes the DA cycloaddition with stereospecificity. Considering the similarity of the cyclization patterns of these two types of compounds, the possibility of the existence of specific DA enzymes in the cytochalasan pathway is high. Additional evidence has come from a study of the pyrichalasin H (164) biosynthetic gene cluster conducted by Hantke and others [266] (Scheme 3D). Through the knockout of pyiF in BGC, the accumulated intermediate metabolites B10–B17 all contain linear PKS moieties with several sites hydroxylated, and it could be speculated that oxygenases downstream or outside the BGC lead to nonspecific modifications to unstable intermediates. Furthermore, when ORF3, the homologous enzyme of PyiF from another cryptic BGC, was reintroduced into the knockout strain, the cyclic pyrichalasin H (164) was again produced. Although the in vivo results obtained were quite convincing, it was unfortunate that attempts to obtain a soluble PyiF protein failed. Direct evidence of a DAase may be expected from the in vitro characterization of such an enzyme in the cytochalasan pathway, including its control over the rate of cycloaddition and stereochemistry.

4.2.3

The Late Stage of Cytochalasan Biosynthesis

After the Diels–Alder cycloaddition, the cytochalasan biosynthesis pathway enters the late stage, which involves a series of oxidative modifications. This could be explained by the abundant oxidoreductases in the BGCs, including, flavin-dependent oxidoreductase, cytochrome P450s, and other enzymes, with functions elucidated continuously by in vivo and in vitro characterizations. In recent years, some novel cytochalasans isolated by our group have been seen to undergo major changes in their skeleton, mainly due to DA and nucleophilic addition reactions between cytochalasans and a polyketide small molecule. Their proposed biosynthesis also will be covered in this Section. The flavin-dependent monooxygenase (FMO) CcsB catalyzes two consecutive oxidations to form a unique carbonate structure (Scheme 4A). Its function was verified by in vivo experiments (gene knockout) and in vitro enzymatic assays by Tang’s group [267]. Interestingly, CcsB catalyzes the first oxygen insertion through a classical Baeyer–Villiger mechanism to form the ester intermediate C2, while the second oxygen insertion can occur only when the substrate C1 has a vinylogous 1,5diketo system. Other FAD-dependent oxidoreductases (OXRs) are involved mainly in ketone-alcohol conversions. Another major group of oxidases in the cytochalasan BGCs are cytochrome P450s. These activate C–H bonds at different sites of the molecules, forming various derivatives. Wan and others elucidated the functions of two P450 enzymes in pyi (the BGC of pyrichalasin H) by gene knockout [8] (Scheme 4B, left). Realizing that these P450s had substrate promiscuity, they applied a combinatorial biosynthesis strategy to replace the pyi P450s with P450s from other homologous BGCs, thus obtaining non-natural novel cytochalasans such as C4 [55] (Scheme 4B, right).

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Scheme 4 Enzymes involved in late-stage modifications. (A) The proposed mechanism of CcsB catalyzing the formation of carbonate. (B) Left: tailoring enzymes on the pyi gene cluster. The arrows point to the modifying sites of each enzyme. Right: CYP3, a cytochrome P450 from another gene cluster, catalyzes the epoxidation to give an unnatural product C4 through combinatorial biosynthesis

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Besides the cytochalasans mentioned above, Zhang and coworkers reported a series of merocytochalasans of considerable complexity [4, 193, 196]. The representative compounds included heterotrimers (455–467) and heterotetramers (468), which are all hybrid molecules of cytochalasan and epicoccine moieties (Scheme 5). A hypothetical biosynthesis was proposed to involve a Diels–Alder reaction and [3 + 2] cycloaddition as key steps, as shown in Scheme 5. The intermolecular Diels–Alder reaction between aspochalasin D (346) and an oxidized epicoccine moiety (D1) led to the linkages of C-19/C-8 and C-20/C-1 with a β-oriented oxygen bridge, giving two products, D4 and D5, with different epicoccine orientations. After this, the trihydroxy benzene parts of D4 and D5 are oxidized to give D6 and D7, respectively. The [3 + 2] cycloaddition occurred between two merocytochalasan intermediates, which led to the new cytochalasan heterotetramer, asperflavipine A (468), bearing two moieties of cytochalasans and two moieties of epicoccines. Intermediate D6, instead of reacting with D7, also may accept an additional oxidized epicoccine molecule (D2 or D3) to give D9 or D8 through a [3 + 2] cycloaddition reaction. These heterotrimer intermediates would undergo additional nucleophilic additions to generate the carbon cage scaffolds. Finally, epicochalasines A and B (461 and 462) would be formed after keto-enol tautomerization. Asperchalasine A (455), a heterotrimer with two cytochalasan moieties and one epicoccine moiety, represents another assembly mode. Aspochalasin B (344), rather than aspochalasin D (346), participates in the DA cycloaddition to give D10. After oxidization to D11, it couples with the second molecule of 344 to furnish asperchalasine A (455). The above pathways are proposed based on the structures of related intermediates isolated from the producing strain. Several obvious uncertainties remain. One is on the origin of the polyketide compound epicoccine, inclusive of the location of its synthetic gene in the genome of the producing strain, and its relation to the cytochalasan BGC in terms of the genome position or expression time. Second, the addition of epicoccine needs to undergo oxidation and [3 + 2] cyclization. It is not known if these processes are spontaneous or enzymatic, or, if enzymes are required, whether they are pathway-specific or non-specific. Currently, the search for the genes involved in epicoccine formation is underway by the present author group, and these points remain to be answered. However, as reviewed in the total synthesis part of this chapter (Sect. 5), the cycloaddition between D11 and aspochalasin B (344) proceeds when directly exposed to the air [198]. It may be suggested that the [3 + 2] cycloaddition is a non-enzymatic reaction in the fungal cell. The assembly of various forms of merocytochalasans reveals unparalleled plasticity in their biosynthesis and may provide new insight into the broad chemical diversity of the merocytochalasan family. It is worth noting that most of these assembled molecules are biologically active (Sect. 3), implying that these are functional metabolites produced by fungi during evolution. It will be interesting to study the metabolic regulation and molecular evolution mechanisms of these compounds during biosynthesis work.

5'

OH

3'

7'

Scheme 5 Proposed biosynthesis of several merocytochalasans

O

O

OH

O

O

O

OH

O

O

OH

O

O HN

O HO

O OH

OH

O

O HO

OH

O HN

O

HO

O

OH

D2

D9

HO

HO

+

O

O

O

D1

1'

8'

O D3

O

OH

O H O OH OH

O

O

OH

O HN

O HO

OH

O

O

OH

D8

[3+2] HO cycloaddition O

HO

HO

D2 or D3

344 R = O (aspochalasin B)

462 (epicochalasine B)

O

O HO

R

19

346 R = OH (aspochalasin D)

O O 20

O O O HN O OH O

OH

HN

1 cytochalasan + 2 epicoccines heterotrimer

461 (epicochalasine A)

O

O

O

O

1'

epicoccine

HO

HO

DA cycloaddition

O

O

O

O

O HO

OH

O

O

O

HO

HN

O

D5

2 cytochalasan + 2 epicoccines heterotetramer

OH

[O ]

HO OH

O HN O O

[3+2] cycloaddition

O HN O OH O

HO

O

O NH

D6

O

HO

HO HO

HO

468 (asperflavipine A)

O

O

OH

O

HO

HO

OH

[O]

O

O HN

D4

OH HO

O

O

OHN

D7

OO

H

NH

O

H O OH

D11

OH

344 (aspochalasin B)

O

O

OH

455 (asperchalasine A)

O

O

OO O

NH

O

O

2 cytochalasan + 1 epicoccines heterotrimer

HO

HO

O

[3+2] cycloaddition

O

[O]

D10

OO OH

NH

HO HO

OO HO

NH

96 H. Zhu et al.

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4.3 Future Prospects The most important breakthrough to be anticipated in future chytochalasan biosynthesis work would be to achieve pyrrolinone formation and DA cycloaddition by heterologous expression. Since the main problems are found from the unexpected side reactions occurring in the common heterologous hosts, feasible solutions may include either finding enzymes in these heterologous hosts that catalyze the side reactions and delete these genes or finding new heterologous expression host strains that do not cause unwanted modifications. These problems may be resolved in the near future and an expression system found that can provide cytochalasin core structures. This will help to characterize more cryptic BGCs than have been discovered to date and facilitate the engineering of biosynthetic pathways to give additional new compounds. In addition, due to the presence of unsaturation on ring C of the cytochalasans, further structural diversification could be targeted. For example, chaetoglobosins generally have a double bond at C-21/C-22 conjugated with adjacent carbonyl groups. At this site, multiple heteromolecular adducts are formed (e.g., in 334–336 and 340– 342 in Fig. 14). It can be speculated that the C-21/C-22 double bond can be attacked conditionally by nucleophilic reagents such as cysteine, or can readily participate in [3 + 2] or [4 + 2] cycloadditions with unsaturated small molecules (e.g., epicoccine and aureonitol). Although enzymatic catalysis cannot be ruled out at this stage, these addition reactions are likely to be spontaneous, and other hybrid molecules could be formed by feeding nucleophiles or unsaturated compounds suitable for cycloadditions. As many BGCs have been discovered, combinatorial biosynthesis will help produce further cytochalasan derivatives. As mentioned earlier, the recombination of cytochrome P450s in different BGCs has produced unnatural derivatives. Other enzymes with novel activities, including CcsB that catalyzes iterative oxidations to form carbonates, could participate in the recombination of modifying enzymes to produce novel derivatives. In addition to the enzymes in the later stages, there are also studies for engineering PKS-NRPS enzymes through domain swapping and heterologous expression [252]. Although newly assembled products were obtained, the cytochalasan backbone was not formed due to the heterologous expression problems mentioned above. Thus, if the side reaction issue in the heterologous hosts could be solved, it will be of great interest to engineer the early stage biosynthesis of cytochalasans. The three-stage biosynthesis of cytochalasans demonstrates the considerable ability of microorganisms to produce highly diversified metabolites. Studying biosynthesis not only will reveal interesting metabolic reactions, but also will inspire bioengineering and biomimetic synthesis work to obtain further interesting cytochalasan derivatives.

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5 Chemical Syntheses of Cytochalasans The typical structures of cytochalasans feature a highly substituted perhydroisoindolone moiety fused by a macrocyclic ring—either a carbocycle (periconiasin G (412) and periconiasin A (404)), a lactone (cytochalasin B (110)) or a cyclic carbonate (phenochalasin A (202)), as shown in Fig. 22A. The macrocycles presented in these structures are normally 11- to 14-membered rings. Periconiasin G (412), bearing a unique 5/6/7-fused ring system, is the smallest member among the cytochalasans [15]. In 2013, cytochalasans bearing a medium-sized ring were identified by Dai and associates, as represented by periconiasin A (404) from the endophytic fungus Periconia sp. [182]. This constituted a new subclass of cytochalasans, featuring an unprecedented 5/6/9 tricyclic framework. In addition, several structurally relevant congeners were also reported by the same laboratory in 2015, as represented by periconiasins D–F (408–410, Fig. 22B) [184]. Differing from previously identified cytochalasans, periconiasins D–F (408–410) possess highly complex polycyclic architectures containing multiple chiral carbon centers. Aspergillin PZ (398) also has a polycyclic architecture and its intricate pentacyclic skeleton features one quaternary carbon and ten contiguous stereocenters, of which five reside on an oxabicyclo[3.2.1]octane subunit [176]. In 2015, Zhang and coworkers isolated and determined a series of novel cytochalasans represented by asperchalasines A–D (455 and 448–450) from a culture broth of Aspergillus flavipes, which was formed by a Diels–Alder reaction between aspochalasin B (344) and epicoccine (Fig. 22C) [193]. Moreover, asperchalasine A (455), the first cytochalasan heterotrimer, features a unique decacyclic 5/6/11/5/5/6/5/11/6/5 ring system consisting of as many as 20 chiral centers. It was the first example of a dimeric cytochalasan alkaloid and later classified as a “merocytochalasan” (Fig. 22C). From this point on, a series of merocytochalasans was discovered by the same group, as exemplified by epicochalasines A (461) and B (462) and asperflavipine A (468), with high degrees of functionalization and intricate polycyclic structures, which greatly enriched the chemical diversity of the cytochalasan family [4, 196]. Equally noteworthy is the fact that asperchalasine A (455) induced significant G1-phase cell cycle arrest by selectively inhibiting cyclin A, CDK2, and CDK6 in cancerous, but not normal cells. On the other hand, epicochalasines A (461) and B (462) induced significant G2/M-phase cell-cycle arrest and apoptosis in leukemia cells through the activation of caspase-3 and the degradation of PARP, thus potentially representing excellent lead compounds for antineoplastic drug development. The complex and densely functionalized structures of cytochalasans combined with their promising biological profiles render them attractive synthetic targets and considerable efforts in the past have been devoted to the total synthesis of cytochalasans. As shown in Fig. 22, the most representative structure of a cytochalasan is a tricyclic hydroisoindolone moiety fused by a macrocyclic ring. Cytochalasans with more complex structures, such as periconiasins D–E (408–410), aspergillin PZ (398) and the merocytochalasans, as shown in Fig. 22B and 22C, can be furnished from their related tricyclic precursor through biomimetic synthesis pathways. Therefore,

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the key to success in the syntheses of cytochalasans is to construct these tricyclic ring systems, which can be disconnected in various ways and lead to two types of synthetic design. Thus, on the one hand, the isoindolone core and the macrocyclic fused ring may be constructed simultaneously through a late-stage intramolecular Diels–Alder condensation, as represented by the total synthesis of cytochalasin B A–tricyclic scaffolds of cytochalasans OH OH O

HN HN O OAc

O

HN

HO

OO

O

O

OH

O O

HO

26 (L-696,474)

110 (cytochalasin B)

202 (phenochalasin A)

HN

HN

OO

OO

HO

412 (periconiasin G)

404 (periconiasin A)

B–polycyclic architectures

O

NH O O

OH

OH

OH

HN O

O

398 (aspergillin PZ)

OH

HN O

HN O

HO

408 (periconiasin D)

HO

409 (periconiasin E)

410 (periconiasin F)

C–merocytochalasans

NH

NH

NH O O O

O

OH

O O

O

O

HO

OH

O O O

O

OH O

OH

448 (asperchalasine B)

HO

O

449 (asperchalasine C)

Fig. 22 Typical structures of cytochalasans

OH

OH HO

O

450 (asperchalasine D)

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HN

O OO O

O

OH O

HO

OH O

O

HO

OO O

O HN

NH

O

OH

O

O HN

O

O

O O

O OH

455 (asperchalasine A)

O

O

O OH

O OH

461 (epicochalasine A)

OH

O OH

OH

462 (epicochalasine B)

Fig. 22 (continued)

(110) by Stork, periconiasin G (412) by Nay, and periconiasins A–E (404–406, 408, and 409) by Tang. On the other hand, the isoindolone core may be formed initially through intramolecular or intermolecular Diels–Alder condensation, with the macrocyclic ring assembled subsequently by the following synthetic steps. In the case of the synthesis of cytochalasin B (110) by Myers, the isoindolone core was synthesized initially by an intramolecular Diels–Alder reaction in an enantioselective and convergent way. The macrocyclic appendage was introduced at a late stage in the synthetic sequence through an intramolecular Horner–Wadsworth–Emmons olefination ring closure. It is also possible to synthesize cytochalasin L-696,474 (26) employing this strategy using common precursors. The construction of the isoindolone core by an intermolecular Diels–Alder reaction has been used in the syntheses of aspergillin PZ (398) by Trauner, and of asperchalasine A (455) by Tang and Deng, respectively. Similarly, intramolecular Horner–Wadsworth–Emmons olefination ring closure was applied to generate the macrocycles by Trauner and Deng. In a different manner, the subsequent introduction of the macrocycle was achieved through an intramolecular ring-closing metathesis in the case of Tang’s synthesis of asperchalasine A (455). The efforts stimulated by total syntheses have demonstrated the diversity and creativity of organic chemistry as applied to cytochalasans. Such results up to 2010 have been summarized in two major comprehensive reviews by Bräse and Hertweck. Since then, significant progress has been made in this area and more efficient syntheses of cytochalasans have been achieved. Meanwhile, merocytochalsans with highly complex structures were also prepared by chemical synthesis. As such, recent examples in the synthesis of cytochalasans will be described below, emphasizing the current advanced nature and state-of-the-art developments in the application of total syntheses to cytochalasans.

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5.1 Synthesis of Periconiasin G Periconiasin G (412) was isolated in 2016 from the endophytic fungus Periconia sp. F-31 was incubated in PDA medium by Dai and coworkers [15]. It has a unique 5/6/7fused ring system and is the smallest member so far discovered among the cytochalasans, showing weak anti-HIV activities. Nay and coworkers reported the total syntheses of four stereoisomers of this seven-membered cytochalasan by biomimic endo and exo IMDA cycloadditions [186]. After careful comparison, a revision of the natural product structure (+)-periconiasin G (462) was achieved, and the stereocenter of C-14 was revised as R. The synthesis of periconiasin G (412) commenced from the protection of (R)citronellal with a 1,3-diol (Scheme 6) followed by dihydroxylation and oxidative cleavage of the diol to deliver carboxylic acid E1. After esterification, the aldehyde E2 was readily obtained under the hydrolysis of the acetal. It was submitted to a modified Takai olefination in the presence of anhydrous CrCl2 and CHI3 to generate the iodoalkene E3, which was coupled with boronic acid E4 through a Suzuki–Miyaura cross-coupling to afford diene E5. After hydrolysis, the carboxylic acid was transformed into an acylimidazole. Then, the pre-lithiated N-benzoyl-5-isobutyl-pyrrolidin-2-one was added to acylimidazole E6 to provide the branched pyrrolidinone E7 in 65% yield. After deprotonation by LiHMDS, phenylselenation and selenide oxidation afforded a highly sensitive linear IMDA substrate E8, which was immediately engaged in a Diels–Alder reaction. The IMDA reaction was performed at 100 °C in CHCl3 and an inseparable mixture of endo and exo cycloadducts was obtained in a 3:1 ratio. The removal of the protective benzoyl amide groups produced a chromatographically separable mixture of periconiasin G (412) and E9. Supported by X-ray crystallography, the proton NMR spectra and optical rotation of synthetic periconiasin G (412) were in full accordance with those of the reported natural product.

5.2 Syntheses of Cytochalasin B and L-696,474 Cytochalasin B (110, Fig. 23), together with cytochalasin A (109), was the first isolated cytochalasan from Helminthosporium dematioideum [15]. In 1978, Stork and coworkers reported the first total synthesis of cytochalasin B (110). The formation of the tricyclic ring was realized by a late-stage intramolecular Diels–Alder condensation [268]. In 2004, Myers and coworkers reported an convergent synthesis route in which the isoindolone core was constructed by a Diels–Alder reaction and the macrocyclic ring was formed by an intramolecular Horner–Wadsworth–Emmons (HWE) olefination ring closure with the macrocyclic appendage introduced at a latestage in the synthetic sequence [269]. Furthermore, using the same strategy, it is also possible to synthesize cytochalasin L-696,474 (26, Fig. 23).

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H. Zhu et al. 1) (CH2OH)2, (±)-CSA, HC(OEt )3 (99 %) 1) BnBr, K2CO3, acetone, 130°C (Mw), 8 min (86 %) 2) AcOH (80% in H2O), 10°C (Mw), 6 min (quant.)

2) OsO4 (1.2 mol%), 2,4,6-collidine, H 2O2, KMnO4 3) NaIO4, H2O/MeOH (10:1), (44% over two steps) O

O

CO2H

O

CO2Bn

O E2

E1 (R)-citronellal

CrCl2, CHI3,

1) aq. NaOH (90%)

2) CDI, Et3N, (96%)

O

CO2Bn

E6

(HO)2B A4 PdCl2(dppf).CH2Cl2, KOH, (4.7:1) (75%, (E)/(Z) = 8:1)

N N

(77%, (E)/(Z) = 11:1)

CO2Bn I

E5 O

E3

Bz N OO

LiHMDS, THF (65%)

1) LiHMDS, PhSeBr (93%) 2) m-CPBA, H 2O2 , CHCl 3

O

1) sealed tube, 100°C (44% over two steps, endo/exo: 3:1 ) 2) aq. NaOH, MeOH, (64% of and 32% of 20)

O

O

NH

(14R)

412 (periconiasin G)

O

NBz

OO

NBz

E7

NH

E8 E9

Scheme 6 Synthesis of periconiasin G OH

OH

HN

O OO

HN O OAc

HO

103 (cytochalasin B)

Fig. 23 Structures of cytochalasin B and L-696,474

26 (L-696,474)

Progress in the Chemistry of Cytochalasans

5.2.1

103

Total Synthesis of Cytochalasin B

The synthesis of cytochalasin B (110) (Schemes 7 and 8) commenced from the intermolecular HWE condensation of diethyl 3-oxo-2-butylphosphonate [270] with N,N-dibenzyl phenylalanal F1 [271] followed by selective mono-N-debenzylation, enolization and silylation to provide the expected diene F2 [272] Of particular mention, it is crucial to choose the N,N-dibenzyl protecting system to avoid epimerization in the HWE reaction [273]. The Diels–Alder substrate F4 was obtained by an addition-elimination sequence by mixing of the silyl enol ether F2 [274] (1.0 eq) with the known exo-methylene lactone F3 (racemic, 1.1 eq) in methanol at 23°C. Under high temperature, intramolecular Diels–Alder smoothly proceeded to generate two diastereomeric products with the desired endo diastereomer F5 in 77% yield. After the deprotection of the N-benzyl protecting group, the tricyclic compound F5 was protected with a Boc group to afford F6 in one pot. After cleavage of 1) diethyl 3-oxo-2-butylphosphonate, Ba(OH) 2, 87% 2) 2,3-dichloro-5,6-dicyanobenzoquinone, 86%

O NBn2

O

OTBS

O

3) TBSOTf, 2,6-lutidine, 99%

B3

AcO

NHBn

N

CH3OH, 98% Cl

F1 Tf2N

O

OTBS O Bn

F4

F2

N B7

OTBS

O

m-xylene, 150°C, 77%.

N

1) TBAF, AcOH 2) KHMDS, THF, B7, 93% (two steps) 3) (CH3)2CuLi, 95% N O

1) H2, 10% Pd/C, BOC2O, Et3N,

O

N

O

O

Ot-Bu

F8

O N Bn

O

Ot-Bu

F5

F6

1) dimethyldioxirane, 100%. 2) trifuoroacetic acid 3) PhI(OAc)2, 4 Å MS, 92% (two steps)

O

O

ethylenediamine, tert-amyl alcohol, 96%

OH N

O O

F9

Scheme 7 Synthesis of intermediate F10

OTBS 96%

TBSO

N H F10

O

O O

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H. Zhu et al. O

O

O

A

B23, KHMDS,

Dess-Martin, OH N H

NaHCO3 N H

O

F10

60% (two steps)

O

N H

O

F11

O TBSO F12

OTBS

1) LDA, Boc2O, 80%; 2)KHMDS, trans-(sulfonyl)-

phenyloxaziridine, 85% O

BocN

O

O

NaOCH2CF3, CF3CH2OH, 65% (two steps)

O 1) diethylphosphonoacetic acid, DCC, 81%

O O O (EtO)2OP F14

2) HF•pyridine, 69% 3) Dess-Martin, NaHCO3

BocN

O OTBS

O F15

OH BocN

OTBS O

O

TBSO

OTBS

F13

O 1) Mg(OCH3)2, CH3OH, 95%

MgSO4, 66%

2) TBAF, THF, 96%

BocN

O

O

B OBn

O

O

OTBS

110 (cytochalasin B)

OBn

TBSO

1) Dess-Martin periodinane 2) TsNHNH2, THF, 94% (two steps) 3) TBSOTf, Et3N

79% (twosteps)

S O2

TBS N N H

F20

I

TBSO

2) PPh3, I2, imidazole, 86%

OTBS F18

2) TBSOTf, Et3N

OBn

HO

O

1) H2, Pd/C, EtOAc,

1) (1S,2S )-1,2-Salen-Co(II), AcOH, H2O (0.45 eq), 41%

O F17

N H

OTBS

O F16

OTBS F19

tert-BuLi, Et2O,

90% (two steps).

OBn

F21

O TBSO OTBS F23

S

O

N Ph N N N

1) H2, Pd/C, EtOAc, 87% 2) 1-phenyl-1H-tetrazole-5-thiol, PPh3, DIAD, 90%

OBn

TBSO OTBS

3) m -CPBA, NaHCO3, 84%

F22

Scheme 8 Synthesis of cytochalasin B and intermediate F23

the silyl enol ether group of the resulting carbamate, region-selective deprotonation of the ketone was accomplished by use of potassium bis(trimethylsilyl)amide (KHMDS) followed by addition of 2-[N,N-bis(trifluoromethylsulfonyl)amino]-5chloropyridine to give the corresponding enol triflate. It then reacted with lithium dimethylcuprate and afforded the trisubstituted alkene F8. The key intermediate F8 was then oxidized followed by the removal of the N-Boc group and oxidation of the resultant amine to deliver the imine F9. An intramolecular transamination and deformylation were achieved in one pot with 1,2-diaminoethane in tert-amyl alcohol, affording the hydroxylactam F10 in 96% yield. Compound F10 was then oxidized to aldehyde F11 and coupled with Nphenyltetrazole sulfone F22, which was synthesized as shown in Scheme 8B. The resulting compound F12 was protected with a Boc group followed by lithiation with KHMDS, which was trapped with trans-2-(phenylsulfonyl)-3-phenyloxyaziridine to afford the tertiary alcohol F13 in 85% yield. After esterification with diethylphosphonoacetic acid [275], tert-butyldimethylsilyl ether on the primary alcohol on the amide F13 was selectively deprotected in the presence of HF/pyridine. The primary alcohol

Progress in the Chemistry of Cytochalasans

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obtained was oxidized to aldehyde F14 and ready for HWE macro-ring-closure. The macrolactone F15 was generated in 65% yield by the use of NaOCH2 CF3 as a base in CF3 CH2 OH and DME. Deprotection of N-Boc followed by hydrolysis of the tert-butyldimethylsilyl ether afforded compound F16, in which the epoxide was transformed into allylic alcohol to deliver the final product cytochalasin B (110).

5.2.2

Total Synthesis of L-696,474

Starting with the same intermediate F8 mentioned above, it was possible to synthesize the HIV-1 protease inhibitor L-696,474 (26) [35, 276] (Scheme 9). After the removal of the N-Boc group, it was then oxidized to the imine F24 by use of [bis(trifluoro-acetoxy)iodo]benzene. Then, this was converted to an aldehyde by a non-hydrolytic opening strategy followed by spontaneous intramolecular transamination and hydrolysis. The crude aldehyde was directly converted to the methyl ester F25 by treatment with potassium hydroxide and iodine in methanol. After epoxidation and the oxidation of the primary alcohol, the resultant aldehyde F26 was reacted with another precursor F33, which was synthesized as shown in Scheme 9B. The ester obtained was treated with lithium dimethyl methylphosphonate followed by cleavage of the TBS group and oxidation of the resultant primary alcohol to afford the aldehyde F27, which was ready for the stage of macrocyclization through an intramolecular HWE reaction. The macrocyclization proceeded smoothly from the treatment of NaOCH2 CF3 as a base in hot dimethoxyethane (80°C) containing 2,2,2trifluroroethanol to afford the tricyclic ketone F28. [274] Stereoselective reduction of the ketone followed by acetylation then afforded the intermediate F29, which was converted to L-696,474 (26) after exposure to magnesium sulfate in warm benzene.

5.3 Total Syntheses of Periconiasins A–E Periconiasins A–C (404–406, Fig. 24), bearing an unprecedented 5/6/9 tricyclic medium-size ring, were reported by Dai and coworkers from the endophytic fungus Periconia sp. F-31 in 2013 [182]. More intriguingly, several structurally relevant congeners were also identified by the same laboratory in 2015, as exemplified by periconiasins D–F (408–410, Fig. 24) [183, 184]. Different from previously identified cytochalasans, periconiasins D–F (408–410) possess highly complex polycyclic architectures bearing multiple stereogenic centers. Biological studies have shown that some periconiasin derivatives display selective cytotoxicity against the HCT-8 and BGC-823 cell lines, thus being of some interest in antineoplastic drug development. The collective total syntheses of periconiasins A–E (408–410, 408, and 409) have been achieved by Tang and coworkers through a highly efficient and biomimetic strategy [277]. The synthesis (Scheme 10) commenced from the alkylation 2-methylcyclohexane1,3-dione G1 followed by partial reduction and mesylation to afford compound G3

Scheme 9 Synthesis of L-696,474 and intermediate F33

B

A

O OAc

N Boc O

OH

OH

F30

N H

O

26 (L-696,474)

HN

F8

O

2) LiH2NBH3, 88% 3) PPh3, I2, imidazole, 89%

1) LDA, LiCl, 2-benzyloxy-1-iodoethane, 94%

MgSO4, 77%

I

2) PhI(OOCCF3)2, 2,6-lutidine, 4 Å MS, 90% (two steps)

1) TFA

F31

HN

OBn

F29

O OAc

O

3) TBDPSCl, Et3N, dimethylaminopyridine, 89%

TBDPSO

2) Ac2O, pyridine, 86% (two steps)

1) CeCl3, NaBH4

2) KOH, I2, 96% (two steps)

1) LDA. LiCl; 2) LiH2NBH3, 91% (two steps)

O F24

N

O

1) 1,3-diaminopropane, CF3CH2OH, Et2O

F32

HN

OBn

O O F28

O

NaOCH2CF3, CF3CH2OH, 52% (two steps, 5:1 mixture of diastereomers)

2) Dess-Martin, NaHCO3.

3) m-CPBA, NaHCO3, 97%

1) H2, Pd(OH)2/C, 2) 1-phenyl-1H-tetrazole-5-thiol, DEAD, 86% (two steps)

CO2Me N H O F25

OH

1) dimethyldioxirane, 95%

TBDPSO

N H

O CO2Me

O

F33

F27

O

O

O

Ph

S

N N N N

O

P(O)(OEt)2

1) B33, KHMDS, 86% (two steps) 2) (CH3O)2POCH2Li 3) TBAF, AcOH, 81% (two steps) 4) Dess-Martin, NaHCO3

N H O F26

O

CHO

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HN

HN

HN

O

HO

HO

405 (periconiasin B)

404 (periconiasin A)

HN O

OO

OO

OO

406 (periconiasin C)

OH

OH

OH

HN O

O

HN O

HO

408 (periconiasin D)

HO

409 (periconiasin E)

410 (periconiasin F)

Fig. 24 Structures of periconiasins A–F

1) NaH, G2, 88% 2) LTBA, 90% 3) MsCl, TEA, 92%

O

1) G4, KHMDS, 70% 2) LiHMDS, PhSeCl 83% 3) H2O2

O

O O

NR

OMs

O G1

G3 G5: R = o-Me-Bz O Br

R N

CHCl3, 90°C, 50%, 2 steps

G2 G4: R = o-Me-Bz

R O N

R O N O

O

+ G6

G7

Scheme 10 Synthesis of the key intermediate G7

[278]. The linear polyketide-amino acid hybrid precursor G3 was assembled by a tandem aldol condensation/Grob fragmentation after an extensive survey of reaction conditions. Deprotonation of G4 with KHMDS followed by quenching the resulting enolate with G3 resulted in spontaneous Grob fragmentation followed by sequential selenylation and oxidative elimination to provide enones G5 [279]. The corresponding products G6 and G7 were obtained in notably improved yields after heating G5 in CHCl3 through the Diels–Alder reaction. Notably, the N-protecting group MeO-Bz is crucial for the improved yield to obtain the crucial tricyclic intermediate G7 compared with Bz [277].

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After sequential selenylation, deprotection, and oxidative elimination, G7 was converted to G8 as a single (Z)-configured isomer. Next, the introduction of the hydroxy group was achieved by 1,4-conjugate addition of alcohol, delivering periconiasin A (404) as a single diastereoisomer after removal of the PMB group with DDQ [280]. The proposed biomimetic transannular carbonyl–ene reaction of periconiasin A (404) was effected under thermal conditions (MeOH, sealed tube, 150°C), to furnish periconiasin E (409) in 36% yield [281]. Furthermore, periconiasin A (404) was oxidized to periconiasin C (406) by Dess–Martin oxidation, and it was then converted into periconiasin B (405) by regio- and diastereoselective reduction of the C-17 carbonyl group. Periconiasin B (405) was submitted to the reaction conditions employed for the synthesis of periconiasin E (409), and this resulted in the direct formation of periconiasin D (408) by tandem transannular carbonyl–ene reaction/etherification (Scheme 11). Meanwhile, an alternative approach was also described for periconiasin D (408) in a diastereoselective manner. Initial oxidation of periconiasin E (409) with DMP produced the corresponding β-hydroxyketone in 83% yield followed by a 1,3-directed reduction in the presence of Me4 NBH(OAc)3 to yield 17-epi-periconiasin E as a single diastereoisomer. Upon treatment with TsOH, it was readily converted into periconiasin D (408) by etherification in 80% yield.

5.4 Total Synthesis of Aspochalasins D and B and (+)-Aspergillin PZ Aspergillin PZ (398) was first isolated in 2002 by Pei and coworkers from Aspergillus awamori and has since been re-isolated from several species of Aspergillus as well as Trichoderma gamsii [171, 176]. In a phenotype assay, aspergillin PZ (398) was reported to induce morphological deformation of the conidia of P. oryzae at 0.089 mM [176]. Structurally, it contains a unique pentacyclic skeleton, which features one quaternary carbon and ten contiguous stereocenters. Its interesting biological activity combined with the synthesis challenge have inspired great efforts of synthesis chemists. In 2011, Overman and coworkers reported the first total synthesis of aspergillin PZ (398) in 28 steps, featuring an unexpected 2-oxonia[3,3]sigmatropic rearrangement/aldol pathway as shown in Scheme 12 [282]. Quite recently, Trauner and coworkers (Scheme 13) have devised an elegant biomimetic synthesis of aspergillin PZ (398) in 13 steps [283]. Since it was believed that aspochalasin D (346) is the likely biogenetic precursor of aspergillin PZ (398) via a “vinylogous Prins reaction”. The synthesis commenced with the ring-opening of epoxy alcohol H1 with propargyl magnesium bromide to generate a diol in excellent yield (Scheme 13). Then, a one-pot ozonolysis-reduction sequence followed by TBS protection provided alkyne H2 in excellent yield over two steps. Silylcupration, alkylative quenching of the in situ generated vinyl cuprate, and treatment of the resulting vinyl silane with NIS generated vinyl iodide H3. It was then coupled with H11 by Suzuki coupling to deliver triene H4 [284]. The Diels–Alder reaction

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G7

1) LiHMDS, PhSeCl; 2) NaOH 3) H2O2 61%, 3 steps

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O HN

1) 1,3-dimesityl-1Himidazol-3-ium chloride, n-BuLi, PMBOH, 70%

O

2) DDQ, 82% G8

MeOH,

OH HN O

150°C, 6%

HN OO HO 404 (periconiasin A)

HO 409 (periconiasin E) 1) DMP 2) Me4NBH(OAc)3 62%, 2 steps

DMP, 85%

OH HN O

HN OO O

17

HO 17-epi-periconiasin E

406 (periconiasin C)

TsOH, 80%

L-selectride,

82%

MeOH, 150°C, 59% HN O

HN OO HO

OH

O

405 (periconiasin B)

408 (periconiasin D)

Scheme 11 Collective synthesis of periconiasins A–E OTBDPS O SnCl4 Prins/pinacol

O

OTBS TBSO X

O

NH

O TBDPSO

O O 398 (aspergillin PZ)

Scheme 12 Overman’s synthesis of aspergillin PZ

OH

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OH

OTBS TBSO

O

1) (PhMe2Si)2Cu(CN)Li2, then MeI 2) NIS, 78% two steps

OTBS TBSO

OTBS

H1

Br

I

H11, Pd(PPh3)4

OTBS

H2

OTBS

H4

H3

TBSO O

OTBS

O N Bz

MeO

10 kbar, 41%

H5

(MeO)2OP O H TBSO

O

H N

O

(MeO)2OP TBSO

1) Et3N•HF 2) DMP

H N

O

Li

O

TBSO H8

TBSO

75%

O

O

O TBSO TBSO

H6

H7

H N

O

TBAF, 53%

O

H N

O

H N

O HO

HO H9

O

DDQ, 45%

HO

TBSO

Bz N

MeO2C

TBSO

TBSO

LiCl, DIPEA, 46% three steps

TBSO

O OMe P OMe

346 (aspochalasin D)

344 (aspochalasin B)

HF/MeCN 89% O

NH O O

OH

398 (aspergillin PZ)

1) CBr4, PPh3, Et3N 2) n-BuLi, then TMSCl, 60%, two steps

O

TMS

1) LiOH 2) (Cat)BH Cy2BH

H H1 0

O B

O

H11

Scheme 13 Total syntheses of aspergillin PZ

smoothly proceeded between H4 and H5 [285] under 10 kbar to furnish H6 with an isomeric ratio of 13:1 [286]. After N-debenzoylation and followed by β-keto phosphonate formation, it was converted to H7. On selective removal of the TBS group, the thus obtained primary alcohol was oxidized to aldehyde H8, which underwent effective macrocyclization under Masamune–Roush conditions (LiCl, i-Pr2 NEt) to provide enone H9 in 46% yield over three steps [287]. Double desilylation of H9 with excess TBAF completed the total synthesis of aspochalasin D (344), which was transformed into aspochalasin B (344) by mild oxidation with DDQ. After a careful screen of a variety of Lewis acids and Brønsted acids, H9 was effectively converted to aspergillin PZ (398) using HF/CH3 CN.

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5.5 Total Synthesis of Asperchalasine A and Related Derivatives Recently, several unprecedented merocytochalasans were identified from the fermentation broth of Aspergillus flavipes by Zhang and coworkers, including asperchalasines A–D (455, 448–450) and epicochalasines A (461) and B (462) [193, 196]. Asperchalasine A (455), which is the first example of a cytochalasan heterotrimer generated by the fusion of two cytochalasan molecules to an epicoccine, possesses an unprecedented 13-oxatetracyclo[7.2.1.12,5 .01,6 ]tridec-8,12-dione core containing as many as 20 chiral centers. Structurally, the dimeric and polymeric features of asperchalasine A (455) are sufficient to make it stand out from the large family of cytochalasans. More importantly, asperchalasine A (455) induced significant G1-phase cell cycle arrest by selectively inhibiting cyclin A, CDK2, and CDK6 in cancerous, but not normal cells, highlighting it as a potentially selective cell cycle regulator against cancer cells. Recently, the synthesis of asperchalasine A (455) and its related derivatives has become a topic of great interest in organic chemistry.

5.5.1

Total Synthesis of Asperchalasines A–E by Tang’s Group

The first total syntheses of asperchalasines A–E (455 and 448–451), a suite of unprecedented merocytochalasans, was achieved by Tang and coworkers [198]. Key to the success of a chemical synthesis approach that would deliver all these merocytochalasans was a convergent and scalable route to the key tricyclic monomer, aspochalasin B (344). Aspochalasin B (344), was synthesized initially through a unified approach that hinges on a Diels–Alder reaction and a ring-closing metathesis reaction as shown in Scheme 14. The synthesis commenced with the preparation of the fragment I5 through sequential acylation of the lactam G4 followed by selenation and oxidative elimination, which was found to be quite unstable [277]. Thus, it had to be promptly submitted in the subsequent Diels–Alder reaction with the diene I8 [288], which was prepared in one step from the known aldehyde I6 through Julia olefination. (CuOTf)2 ·PhMe was found to the best catalyst, and yielded I9 as a single adduct in 63% yield [289, 290]. The 11-membered macrocycle was formed through the RCM reaction using the Grubb’s second-generation catalyst, to afford tricyclic intermediate I10 as a single (E)-isomer in 85% yield [291]. Dihydroxylation of the double bond followed by removal of the N-benzoyl group afforded the trans-diol I11. Then, selective protection of OH-17, oxidation of OH-18, and removal of the protecting group converted I11 into I12, which was transformed into aspochalasin P (358) by removal of the acetyl group. Finally, selenylation followed by oxidative elimination afforded aspochalasin B (344) in 40% yield. The bioinspired Diels–Alder reactions of aspochalasin B (344) with different epicoccine precursors were then studied, which enabled the divergent synthesis of the heterodimers asperchalasines B–E (448–451) and its related congeners (Scheme 15). Treatment of aspochalasin B (344) with I14 [292] using CSA as acid smoothly

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O

O

I1, t-BuLi, then E2, 60%;

H

O

O

O LiHMDS, PhSeCl, 78 %

N R

N R PhSe I4

I3

Br R = o-Me-Bz G4

Cl I1

I2

H2O2

O

O

O E7, LiHMDS, 72% ((E)/(Z) > 10/1)

N R

+

O S

I6

N

O HO HO

Boc N

O

O S O I7

I8

I5 (CuOTf)2•PhMe, 63 % over 2 steps

1) Grubb's catalyst (II), 85 % 2) OsO4, NMO, 45 %;

O

Boc N

O

Grubb's catalyst (II), 85 %

O

H N

O

18

17

I11

I10

I9

1) NaOH, 90%; 2) BzCl, Et3N, DMAP, 85%; 3) DMP, 83%

O

H N

O

NaSMe, 75%

O

O

O

1) LiHMDS, PhSeCl; 2) H2O2, 40 % over two steps

H N

O HO

BzO I12

O

H N

O

O

HO 358 (aspochalasin P)

344 (aspochalasin B)

Scheme 14 Total syntheses of aspochalasin B

HN OO HO

O

HN OO O

OH

HN

OAc

OO

O

HO OH

HO

O

I14

O

OH O

HO OH

O

OH

O

451 (asperchalasine E)

448 (asperchalasine B)

O

+

CSA, BHT, toluene, 60°C 80% (373:407:374:375 = 10:2:1:1)

344 (aspochalasin B ) HN OO

O

OH O

HN OO

O

OH O OH

HO HO

O

449 (asperchalasine C)

Scheme 15 One-pot syntheses of asperchalasines B–E

HO

O

450 (asperchalasine D)

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generated asperchalasines B–E (448–451) (10:2:1:1) in a combined yield (80%). Interestingly, if the fully protected hemiacetal I15 was used in the reaction, only two endo-type products, asperchalasines E and D (451 and 450) (57%, 3:2), were detected after debenzylation (Scheme 15). A similar outcome was obtained with I16, affording two endo-adducts I19 and I20 in excellent combined yield (80%, 3:2). Finally, the biomimetic oxidative formal [5 + 2]-cycloaddition [293, 294] was achieved to deliver asperchalasine A (455) (Scheme 16) after debenzylation of I19 with Raney Ni/H2 followed by directly exposed to air in the presence of aspochalasin B (344).

5.5.2

Total Synthesis of Asperchalasines A, D, E, and H by Deng’s Group

Concurrent with the work of Tang and associates, the group of Deng, from the Kunming Institute of Botany, also reported independently the total syntheses of asperchalasines A (455), D (450), E (451), and H (454). Deng’s synthesis started with the stereospecific preparation of the triene segment J4 (Scheme 17). Initially, the l-arabinose was converted to hemiacetal J1 through a known three-step sequence [295]. After sequential Wittig olefination, silylation protection, and hydrogenation of the resulting double bond, the methyl ketone J2 was obtained in 69% yield. This was then coupled with the dienyl phosphonate J3 to afford the conjugated triene J4 (82%, (E)/(Z) = 7:1). The known lactam J5 [186], prepared from N-Boc-l-leucine in five steps, was reacted with methyl chloroformate, affording the methyl ester J6. After sequential selenylation and oxidative elimination, it was transformed into the activated dienophile J7. A mixture of J7 and J4 was heated to 100 °C to furnish the Diels–Alder products J8 and its isomer (85%, ((E)/(Z) = 2:1) [296]. The addition of lithium dimethyl methylphosphonate followed by selective deprotection and oxidation of the resultant primary alcohol yielded the aldehyde J9. The HWE macrocyclization was accomplished by the use of Zn(OTf)2 [297]. After deprotection of the TBS group followed by selective oxidation, aspochalasin D (346) and aspochalasin B (344) were obtained in overall 79% and 72% yields, respectively. Then, aspochalasin B (344) was added to the active diene, which was generated in situ by the acidic treatment with J10, affording the endo-Diels–Alder adducts J12 and its regioisomer J13 in 78% yield (Scheme 18) [298, 299]. Both adducts were subjected to deallylation to furnish J14 and asperchalasine H (454). Similarly, asperchalasines D (450) and E (451) also could be obtained using the same sequence for the synthesis of asperchalasine H (454) in 73% overall yield as shown in Scheme 18B. Treatment of J14 with potassium ferricyanide led to facile oxidation of the electron-rich aromatic ring to yield the corresponding o-quinone, which was unstable and trapped by another molecule of aspochalasin B (344) in the presence of sodium bicarbonate to furnish the formal the [5 + 2] adduct asperchalasine A (455) in 49% yield [300].

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OH OH

HO

I13

HN OO

O

1) CSA, toluene, 80°C 2) Ac2O, DMAP, DCM 30% (5:1)

O

HN

OAc

OO

O

OH O OAc

AcO OAc AcO I17

HO

O

AcO I18

OAc

O OH

HO

HN

I14

OO

O

CSA, toluene, 60°C 63% (10:1)

O

HN

OH

OO

O

OH O

HO

HO

OH O 451 (asperchalasine E)

OH O 448 (asperchalasine B)

HN

O

OO

OH O 344 (aspochalasin B)

BnO

O OBn

HO

O

I15

HN OO

1) CSA, toluene, 60°C 2) Raney Ni, H2, EtOH 57% (3:2)

O

HN

OH

OO

O

OH O OH

HO

O HO 350 (asperchalasine D)

OH O 451 (asperchalasine E) BnO

O

OBn OBn

HO

O

I16

HN

HN OO

CSA, toluene, 60°C 80% (3:2)

O

OO

OH

O

O

OH O OBn

BnO BnO

OBn

O OO O

OH O

HO

aspochalasin B [5+2]

O

HO

OO O

I20

Raney Ni H2, EtOH

I19

HN

OBn

BnO

NH

57% from I19

HN OO

O

OH O

Scheme 16 Total syntheses of asperchalasines A–E

HN OO

O

OH O

HO

HO

455 (asperchalasine A)

Air

O I22

O

HO I21

OH

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O OH

115

1) MeCOCHPPh3 2) TBSOTf, 2,6-lutidine 3) H2, Pd/C 66%, three steps

O

F3, KHDMS, 82%, (d.r. 7:1)

OTBS

TBSO

HO

OTBS OTBS

OH OTBS

TBSO

J1

J2

5 steps, 57%.

O

O O NHBoc

J4

PO(OEt)2 J3

CO2Me

LiHDMS, ClCO2Me

LiHDMS, PhSeCl, then H2O2, 88% two steps.

CO2Me O N Bz J7

O

N Bz J5

N Bz J6

F4, neat, 100°C, (E)/(Z) = 2:1 1) Zn(OTf)2, Et3N, TMEDA; 2) TBAF, 79% two steps.

1) BuLi, MePO(OMe)2 2) HF•pyridine 3) DMP, 72% three steps

BzN O O

HN O O

HO 346 (aspochalasin D)

OH

OTBS

(MeO)2OP TBSO

O

BzN O CO Me 2 OTBS J8

TBSO

OTBS

J9

TsOH•H2O, TEMPO 92% HN O O OH O 344 (aspochalasin B)

Scheme 17 Total synthesis of aspochalasin B

5.6 Future Prospects The fascinating structures of the cytochalasans have motivated relevant investigations by several synthesis chemists thus far. Diversified strategies and creative solutions have been realized in the pursuit of these complex molecules, ultimately resulting in an inspiring series of successful total syntheses, as documented above. However, there is the potential for new approaches for the advancement of this type of scientific work on the cytochalasans. The complexity of isolated cytochalasans is ever increasing as unprecedented structures are continuing to be disclosed. For example, asperchalasine A (455), with both a high degree of functionalization and an intricate polycyclic structure has enriched the known cytochalasan chemical diversity, and it was synthesized chemically through a biomimetic Diels–Alder reaction followed by an oxidative [5 + 2]cyclo-addition. Different from compound 455 in their fusion patterns, the oxidized epicoccine moiety in epicochalasines A (461) and B (462) may be rotated when at 180 °C, thus resulting in different carbon skeletons. These two natural products remain yet unconquered by total synthesis and should stimulate the development of novel synthesis strategies in order for them to be produced. In terms of future prospects for synthesis efficiency, it seems that these natural products are too complex to allow truly concise and scalable routes. Moreover, the

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A OAllyl

OAllyl

AllylO

HOAc

O AllylO

then aspochalasin B

AllylO

HN

O

OO

AllylO

HN

+

OH

O

OO

O

O

OH O

OH J10

J11

OAllyl

AllylO OAllyl

AllylO J12

Pd/C, HCO2NH4, 70%

Pd/C, HCO2NH4, 75%

then aspochalasin B

HN OO

49%

455 (asperchalasine A)

O

K3Fe(CN)6

OH

HN OO

O

HN

OH

OO

O

O

O

OH O OH

HO

HO O J15

O

OAllyl

AllylO J13

HO

OH

HO

J14

OH

454 (asperchalasine H)

B 1) HOAc, then aspochalasin B 2) Ph(PPh3)4, Et3SiH, 72%, two steps

OAllyl O

HN

O

OO

AllylO

O

OH

HN

+

OO

O

OH

O

OH O

12 : 1

J16

OH

HO

O HO 350 (asperchalasine D)

OH O 451 (asperchalasine E)

C 65% over 3 steps

O O

1) BBr3 2) K2CO3 allyl bromide 83%

O O

OAllyl AllylO

O

O

O

COOH

O AllylO

O

O

1) BBr3 2) NaHCO3, KI, 75% 1) K2CO3 allyl bromide 2) DIBAL-H 72%

OAllyl O

OAllyl

OH AllylO

O

O AllylO

O

O AllylO

HO O

DIBAL-H

OH

O

J16

J10

Scheme 18 Total syntheses of asperchalasines A, D, E, and H

medium to large rings with their tricyclic structures usually require highly dilute concentrations, which is a major limitation on the amount of a synthetic product finally obtained. Creative synthesis strategies and the development of highly efficient cascade reactions could provide solutions to this problem, as exemplified by the syntheses of periconiasin G (412) through an IMDA reaction by Nay and coworkers and the concise total syntheses of periconiasins A–E (404–406, 408, and 409) by Tang that enabled the creative use of the Grob reaction during the process used. The development of new synthesis methodologies will also be fundamentally important

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for chemists to deliver ample amounts of compounds in order to probe the bioactivity of these natural products and study their structure-activity relationships through chemical derivatization. Last, but not least, their highly complex structures combined with the noncrystalline properties of some of these natural products makes their structural characterization a great challenge, as illustrated by the case of periconiasin G (412). Additional work must be conducted on the syntheses of these natural products in order to correctly assign structures from an analytical point of view, since chemical synthesis still has important roles to play in the process of solving some of Nature’s most intriguing molecular puzzles. The cytochalasan family of compounds has attracted considerable attention over several decades, and many achievements in their total syntheses have been realized. In light of the increasing structural complexity of newly found derivatives, cytochalasans will undoubtedly continue to serve as a compound class of major interest for organic synthesis.

6 Conclusions Cytochalasans are a class of structurally complex and biologically active alkaloids found predominantly from various fungi, including from the genera Chaetomium, Aspergillus, Phoma, and Phomopsis, which have attracted substantial attention from chemists and biologists all over the world. Several reviews have been published previously on cytochalasans, focusing on their isolation and characterization, biosynthesis, and organic synthesis. However, laboratory investigations on the cytochalasans have grown rapidly in recent years, so it was considered necessary by the present authors to write the present updated chapter. This contribution covers 477 cytochalasans reported between the years 1966 through 2020, and provides an overview of research on these compounds in terms of their isolation, structural determination, biological activities, biosynthesis, and total synthesis. It in order to interest further pharmaceutical companies on the biological potential of cytochalasans, new assays on inactive or untested cytochalasans should be carried out, with the aim of developing structure-activity relationship information to discern the functionalities responsible for the activities of members of this compound class. The future contributions of natural product chemists will promote research on the total synthesis and biosynthesis of cytochalasans, which in turn will promote investigations on their more advanced biological testing. As a result of additional interdisciplinary efforts from members of the chemical and biomedical scientific communities, the present authors believe that cytochalasans will play an important role in the future development and utilization of natural products as a whole.

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Huchen Zhu obtained his Ph.D. degree in pharmaceutical sciences in 2014 from Huazhong University of Science and Technology (HUST), under the direction of Prof. Yonghui Zhang. After two years of postdoctoral research, again with Prof. Zhang (2014–2016), he was appointed as Associate Professor and has continued to work with the group of Prof. Zhang. He is the recipient of an Excellent Young Scholars award of the National Natural Science Foundation of China and was nominated as Professor in 2019. The main topics of his research program are on structurally unique and pharmacological active secondary metabolites from both medicinal plants and fungi.

Chunmei Chen obtained her Ph.D. degree in pharmaceutical sciences in 2015 from Huazhong University of Science and Technology (HUST), under the direction of Prof. Yonghui Zhang. She worked as a postdoctoral research assistant from 2015 to 2017, again in the group of Prof. Y. Zhang, and then became a Lecturer at the School of Pharmacy, HUST in 2017. She was appointed as an Assistant Professor in 2019. Her main research focus is on the isolation and structural characterization of compounds of interest from natural sources with the potential for drug development.

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Qingyi Tong graduated with a bachelor’s degree in bioengineering from Nanyang Institute of Technology in 2009. He then obtained his Ph.D. degree in 2014 in the Biotherapy of Major Human Diseases from Sichuan University, under the direction of Professor Xiaohua Wu. During his doctoral studies, he won a scholarship from the Japan Student Services Organization (JASSO), and went to the Medical School of Kyoto University to study in 2013. After almost four years of postdoctoral research in pharmacology with Professor Yonghui Zhang at Huazhong University of Science and Technology (HUST), in 2018 he took up a Lecturer position at the School of Pharmacy, Tongji Medical College, HUST, and was promoted to Associate Professor in 2020. He has studied the pharmacological activities of many natural products and their derivatives, focusing on cancer-related and anti-inflammatory activities and effects on metabolic diseases, and also conducts mechanistic work on active compounds. He has authored or coauthored more than 30 international papers in pharmacology and new drug discovery. Yuan Zhou obtained his Ph.D. degree in 2011 from Peking University under the direction of Professor Pengfei Tu. After two years working in the Yangtze River Pharmaceutical Group (2011–2013), he moved to the Institute of Vegetables and Flowers, Chinese Academy of Agricultural Sciences, as a Postdoctoral Fellow, and began to investigate the biosynthetic pathways of natural products from plant sources with Professor Sanwen Huang (2013–2016). Currently, he is an Associate Professor at the School of Pharmacy, Tongji Medical College, Huazhong University of Science and Technology (2016–). His current research is focused on the isolation of bioactive compounds from natural sources such as plants and fungi, as well as studying the biosynthesis of these bioactive natural products. Ying Ye received her Ph.D. degree in 2017 from Hokkaido University in Japan, where she studied the biosynthesis of fungal secondary metabolites under the supervision of Professor Hideaki Oikawa. Then, she received postdoctoral training from 2017 to 2018 with Professor David Sherman at the University of Michigan. After this, she was appointed to the position of Associate Professor at Huazhong University of Science and Technology, where she is a member of Yonghui Zhang’s group. Her current research is oriented towards the biosynthesis of natural products from fungi, as well as the discovery of novel metabolites and enzymes through biosynthesis investigations.

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H. Zhu et al. Lianghu Gu obtained his Ph.D. degree on chemistry in 2016 from Max-Planck-Institut für Kohlenforschung in Germany under the co-direction of Prof. Manuel Alcarazo and Prof. Alois Fürstner. After two years of working at WuXi AppTech Wuhan as a Senior Research Scientist (2016–2018), he then commenced a position as Associate Professor at Huazhong University of Science and Technology, where he is a member of the group of Prof. Yonghui Zhang. His current research is based on the synthesis of bioactive natural products for use as lead compounds in drug discovery.

Yonghui Zhang obtained a Ph.D. degree in pharmaceutical sciences in 2004 from Huazhong University of Science and Technology (HUST). Then, he was appointed in turn as a Lecturer (2005), Assistant Professor (2005–2008), and Professor (2008–present) at the School of Pharmacy, Tongji Medical College, HUST. He is currently Dean of the School of Pharmacy (2014), Distinguished Professor of the Program for Changjiang Scholars of the Ministry of Education of China (2016), and Distinguished Young Scholar of the National Natural Science Foundation of China (2017). He is a medicinal chemist focusing on natural compounds, and engages in research on structurally unique and pharmacologically active secondary metabolites from natural sources that have the potential for development as new drugs.

Bioactive Compounds from Medicinal Plants in Myanmar Nwet Nwet Win and Hiroyuki Morita

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Phytochemicals and Pharmacological Activities of Medicinal Plants from Myanmar . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Acacia concinna Wall. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Orthosiphon stamineus Benth. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Panax zingiberensis C.Y. Wu & Feng (Myanmar wild ginseng) . . . . . . . . . . . . . . 2.4 Caesalpinia crista L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Elephantopus scaber L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Phyllanthus niruri L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Streptocaulon tomentosum Wight & Arn. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Curcuma comosa Roxb. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Vitis repens Wight & Arn. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Boesenbergia pandurata (Roxb.) Schltr. (Boesenbergia rotunda (L.) Mansf.) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11 Kayea assamica King & Prain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.12 Cordia fragrantissima Kurz. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13 Thanakha plants: Hesperethusa crenulata L. and Limonia acidissima L. . . . . . . 2.14 Soymida febrifuga (Roxb.) A. Juss. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.15 Vitex negundo L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.16 Diospyros burmanica Kurz. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.17 Cinnamomum inunctum Kurz. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.18 Kaempferia pulchra Ridl. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.19 Picrasma javanica Blume . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.20 Curcuma amada Roxb. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.21 Vitex trifolia L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.22 Mansonia gagei Drumm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.23 Premna integrifolia L. (syn.: P. serratifolia L.) . . . . . . . . . . . . . . . . . . . . . . . . . . .

136 137 137 143 145 148 152 155 158 162 166 168 174 178 180 185 189 192 192 194 198 201 204 206 208

N. N. Win · H. Morita (B) Institute of Natural Medicine, University of Toyama, 2630-Sugitani, Toyama 930-0194, Japan e-mail: [email protected] N. N. Win e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 114, https://doi.org/10.1007/978-3-030-59444-2_2

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2.24 Picrorhiza kurroa Royle ex Benth. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.25 Jatropha multifida L. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.26 Swertia chirata Buch.-Ham. ex Wall. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Synthesis Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 Introduction The Republic of the Union of Myanmar is a tropical country that is situated geographically in Southeast Asia and occupies a land area of 676,577 km2 and a coastline of 2,832 km. It lies between latitudes 9° 32 N to 28° 31 N and longitudes 92° 10 E to 101° 11 E and is comprised of seven states and seven regions, with the former consisting of Chin, Kachin, Kayah, Kayin, Mon, Rakhine and Shan, and the latter Ayeyarwady, Bago, Magway, Mandalay, Sagaing, Tanintharyi, and Yangon. Myanmar borders Thailand to the southeast and east, with Laos as a part of its east border following the Mekong River, as well as China (Yunnan and Tibet) to the northeast and India to the northwest. A small part of the west is bordered by Bangladesh. Generally, Myanmar has three different seasons characterized as summer (from March to May), rainy (from June to October), and winter (November to February). Different landscapes, such as mountains, lowlands, plateaus, plains, rivers, and lakes occur from the east to the west and from the north to south. The average temperature in the lowlands, plains, and coastal zones is approximately 27 °C. Fifty percent of the land area was covered by forest 50 years ago. However, the amount of forest area is decreasing gradually due to meeting a continuing demand for timber. Nevertheless, Myanmar is a country that still retains valuable natural resources that are being lost elsewhere in the world. The population of Myanmar has inherited its own traditional medicine system and has practiced this for over 2000 years. Traditional medicine in Myanmar is based on Ayurvedic concepts and is influenced by Buddhist philosophy. In the primary healthcare system of Myanmar, traditional medicine still occupies an important and integral role. The traditional medicinal formulation used in Myanmar is abbreviated as “TMF.” Altogether, 57 formulations have been documented in a manual for Myanmar traditional practitioners. The formulations generally consist of three major ingredients, which originate from medicinal plants, animals, and minerals. Medicinal plants are the major ingredients in most formulations, whereas minerals and animal products are minor ingredients. The population has utilized not only the TMF but also medicinal plants in their daily life to prevent, alleviate, and cure human disease from time immemorial. At the time of writing this chapter, the chemical constituents of more than 26 plants in Myanmar have been investigated, and the results have been reported in international and local journals. Some biological activities of the extracts and the isolated compounds from the plants from Myanmar have also been documented in these reports. This chapter covers phytochemical and

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biological studies on these 26 Myanmar plants, based both on publications in international journals and two Ph.D. dissertations that have appeared between 2000 and 2020. The species names of the plants, their families, and years of publication of the relevant literature reports are summarized in Table 1.

2 Phytochemicals and Pharmacological Activities of Medicinal Plants from Myanmar 2.1 Acacia concinna Wall. Acacia concinna, belonging to the family Leguminosae, is a medicinal plant that grows in the tropical rainforests of southern Asia (Fig. 1).In Myanmar, this plant is known locally as Kin-moon-gyin. The young leaves and ripened dried pods have been recognized as the plant parts useful in traditional medicine in Myanmar. The young leaves have a sour taste, but the soup or fried leaves, together with fish paste and prawns, are popular appetizer cuisine. A decoction of the pods has been used for Fig. 1 Acacia concinna, known as Kin-moon-gyin in Myanmar. The young leaves and ripened dried pods are the parts used in traditional medicine in Myanmar

Scientific name

Acacia concinna Wall.

Orthosiphon stamineus Benth.

Panax zingiberensis C.Y. Wu & Feng

Caesalpinia crista L.

Elephantopus scaber L.

Phyllanthus niruri L.

Streptocaulon tomentosum Wight & Arn.

Curcuma comosa Roxb.

Vitis repens Wight & Arn. (syn. Cissus repens Lan)

Boesenbergia pandurata (Roxb.) Schltr. Boesenbergia rotunda (L.) Mansf.

Kayea assamica King & Prain

Cordia fragrantissima Kurz.

Thanakha plants: Hesperethus acrenulata L. Limonia acidissima L.

Soymida febrifuga (Roxb.) A. Juss.

Vitex negundo L.

Diospyros pendula Hasselt. ex Hassk.

Cinnamomum inunctum Meisn.

Kaempferia pulchra Ridl.

No.

1

2

3

4

5

6

7

8

9

10

11

12

13

14

15

16

17

18

Shan-pan-oot

Karaway

Te

Kyaung-ban-gyi

Dan-da-kuu-ni

Thanakha Thi

Sandawa

Theraphi

Seik-phoo Seik-phoo-chin

Ta-bin-daing-mya-nan

Nanwin-yaing

Myin-sagon-ni

Taung-zi-phyu

Taw-monlar

Kalein

Myanmar Wild Ginseng

Se-cho or Myit-shwe

Kin-moon-gyin

Local name

Zingiberaceae

Lauraceae

Ebenaceae

Lamiaceae

Meliaceae

Rutaceae Rutaceae

Boraginaceae

Clusiaceae

Zingiberaceae

Vitaceae

Zingiberaceae

Asclepiadaceae

Euphorbiaceae

Asteraceae

Fabaceae

Araliaceae

Lamiaceae

Leguminosae

Family

Rhizome

Fruit

Wood

Fruit

Bark

Bark Bark

Wood

Bark Flower

Rhizome Rhizome

Rhizome

Rhizome

Root

Whole plant

Whole plant

Seed

Rhizome

Whole plant

Fruit

Part used

(continued)

[356–361, 367]

[350]

[335]

[331]

[311]

[274] [277–279]

[258]

[23] [238, 239]

[211, 212] [226]

[146, 186, 187]

[146]

[141, 145]

[132]

[87]

[61–63, 73]

[44]

[27, 28, 34]

[3]

References

Table 1 List of phytochemical and biological investigations of medicinal plants from Myanmar that appeared in international publications from 2000 to 2020

138 N. N. Win and H. Morita

Scientific name

Picrasma javanica Blume.

Curcuma amada Roxb.

Vitex trifolia L.

Mansonia gagei Drumm.

Premna integrifolia L (syn.: P. serratifolia L.)

Picrorhiza kurroa Royle ex Benth.

Jatropha multifida L.

Swertia chirata Buch.-Ham. ex Wall.

No.

19

20

21

22

23

24

25

26

Table 1 (continued) Local name

Pan-khar

Say-ma-khan

Saung-may-ga

Taung-tan-gyi

Karamat

Kyaung-ban-lay

Thayetkin

Nan-paw-kyawt

Family

Gentianaceae

Euphorbiaceae

Scrophulariaceae

Verbenaceae

Malvaceae

Lamiaceae

Zingiberaceae

Simaroubaceae

Part used

Whole plant

Stem

Stem

Wood

Bark

Whole plant

Rhizome

Bark Wood

[537]

[525, 526]

[509]

[476, 482, 483, 490]

[467]

[451, 455]

[429]

[367, 395–398] [399]

References

Bioactive Compounds from Medicinal Plants in Myanmar 139

140

N. N. Win and H. Morita

washing hair to promote hair growth, and as an expectorant, emetic, and purgative [1]. In addition, the dried powder of the fruits is utilized as a body scrub in Myanmar. In Thai folk medicine, the seeds of A. concinna have been used as a therapy for certain skin diseases, and flavonoids and monoterpenoids have been reported as constituents of a sample collected in Thailand [2]. Phytochemical studies of the pods of A. concinna in Myanmar were reported in 2000 [3], and the potential of the pods as a natural source of the discovery of cytotoxic compounds was shown for the first time. In addition, this was the first report in an international journal on the investigation of the chemical constituents of this species when collected in Myanmar. Three unprecedented new saponins, named Fig. 2 Structures of 1–4 isolated from the MeOH extract of Acacia concinna pods

O

6'

O O CH OH 2 O

OH HO O

OH

O

O

OH OH

OH O OH

O

OH HO

O OH

O

OH

O R

O

O

O O

HO OH

OH

OH OH

O

HO

OH

O OH OH

OH O OH

1 (kinmoonoside A) R = X, 6' = R 2 (kinmoonoside B) R = X, 6' = S 3 (kinmoonoside C) R = H

X= O O HOH2C

O OH O O HOH2C

OH OH OH

4 (4-O-[(2E)-6-hydroxy-2-hydroxymethyl-6-methyl2,7-octadienoyl]-D-quinovopyranose)

OH OH

Bioactive Compounds from Medicinal Plants in Myanmar

141

kinmoonosides A–C (1–3), and the new monoterpenoid 4-O-[(2E)-6-hydroxy2-hydroxymethyl-6-methyl-2,7-octadienoyl]-d-quinovopyranose (4) were isolated from a cytotoxic MeOH extract of the fruits of A. concinna, which showed cytotoxic effects against HT-1080 human fibrosarcoma cells [3] (Fig. 2). Kinmoonosides A–C (1–3) have a conserved trisaccharide moiety at C-3 and the same tetrasaccharide at C-28 of the acacic acid moiety, and kinmoonosides A and B (1 and 2) showed a diastereomeric relationship at C-6 . On the other hand, kinmoonoside C (3) displayed structural differences compared with 1 and 2 at C-6 , since compound 3 lacks a quinovopyranosyl-monoterpenyl moiety. At present, only kinmoonosides A–C (1–3) have been reported from A. concinna in Myanmar, suggesting that these compounds may be utilized as chemical markers for this plant source. In addition to these naturally occurring compounds, four monoterpenoids (5–8) (menthiafolic acid (5) [4], a 1:1 mixture of (6S)- and (6R)-menthiafolic acid 6-O-β-d-quinovopyranoside (6) [4], a 2:1 mixture of two diastereomers of 3-hydroxy-2-(5-methyl-5-vinyltetrahydrofuran-2-yl) propanoic acid (7) [5], a 1:1 mixture of (6S)- and (6R)-(2E)-6-hydroxy-2-hydroxymethyl-6-methyl-2,7octadienoic acid 6-O-β-d-quinovopyranoside (8)) (Fig. 3), five prosapogenins (9–13) (acacic acid 3-O-α-l-arabinopyranosyl (1→6)-[β-d-glucopyranosyl(1→2)]-β-dglucopyranoside (9), acacic acid 3-O-α-l-arabinopyranosyl(1→6)-2-acetamido-2deoxy-β-d-glucopyranoside (10) [3], acacic acid 3-O-β-d-xylopyranosyl(1→2)-αl-fucopyranosyl(1→6)-[β-d-glucopyranosyl(1→2)]-β-d-glucopyranoside (11) [6], acacic acid 3-O-β-d-xylopyranosyl(1→2)-α-l-arabinopyranosyl(1→6)-2acetamido-2-deoxy-β-d-glucopyranoside (12) [7], and acacic acid lactone 3-O-β-dxylo-pyranosyl(1→2)-α-l-arabinopyranosyl(1→6)-[β-d-glucopyranosyl(1→2)]β-d-glucopyranoside (13, concinnoside F = albiziasaponin C) [4, 8])), and an acacic acid lactone (14) [9] (Fig. 4) were obtained from the crude saponin fraction of A. Fig. 3 Structures of monoterpenoids 5–8 obtained from the crude saponin fraction of A. concinna by alkaline and acid hydrolysis

COOH

COOH O OH

Qui = OH

O

OH

Qui 5 (menthiafolic acid)

HOH2C

COOH

OH OH

6 ((6S)- and (6R)-menthiafolic acid 6-O-β-D-quinovopyranoside) HOH2C

COOH

O O Qui 7 (3-hydroxy-2-(5-methyl-5-vinyltetrahydrofuran-2-yl)propanoic acid)

8 ((2E)-6-hydroxy-2-hydroxymethyl6-methyl-2,7-octadienoic acid 6-O-β-D-quinovopyranoside)

142

N. N. Win and H. Morita OH

COOH OH RO

9 (acacia acid 3-O-α-L-arabinopyranosyl(1 6)β-D-glucopyranosyl(1 2)]-β-D-glucopyranoside)

R = -Glc 2 Glc

6

Ara

10 (acacic acid 3-O-α-L-arabinopyranosyl(1 6)2-acetamido-2-deoxy-β-D-glucopyranoside)

R = -GlcNAc

11 (acacic acid 3-O-α -D-xylopyranosyl(1 2)-α -L -fucopyranosyl(1 6)-[β-D-glucopyranosyl(1 2)] -β-D-glucopyranoside

R = Glc Fuc Xyl

12 (acacic acid 3-O-β-D-xylopyranosyl(1 2)-α -L -arabinopyranosyl(1 6)-2-acetamido-2-deoxy -β-D-glucopyranoside)

R = -GlcNAc

6

6

Ara

2

2

Glc 6

2

Ara Xyl

O O OH RO

13 (acacic acid lactone 3-O-β-D-xylopyranosyl(1 2)α -L-arabinopyranosyl(1 6)-[β-D-glucopyranosyl(1 β-D-glucopyranoside) 14 (acacic acid lactone) R = H

2)]- R =

Glc 2

6

2

Ara Xyl

Glc

Fig. 4 Structures of prosapogenins 9–13 and acacic acid lactone 14 obtained from the crude saponin fraction of A. concinna by alkaline and acid hydrolysis

acacia as alkaline- and acid-hydrolyzed products during the structure elucidation of the kinmoonosides. Among the hydrolyzed products, compounds 8–10 were determined as being new compounds. Furthermore, it was demonstrated that kinmoonosides A–C from the pods of A. concinna grown in Myanmar are chemical components with substantial cytotoxicity, and they exhibited ED50 values of 0.70, 0.91, and 2.83 μM, respectively, against human HT-1080 fibrosarcoma cells and were more potent than 5-fluorouracil (ED50 8.0 μM) used as positive control [10]. In contrast, the monoterpenoids 4–8 and prosapogenins 9–13 were inactive (ED50 > 100 μg/cm3 ) in the in vitro assay used. Earlier, the presence of the tetrasaccharide moiety at C-28 in acacic acid aglycone has been demonstrated as being important for mediating cytotoxicity, as reported in the case of julibrosides I–III [7].

Bioactive Compounds from Medicinal Plants in Myanmar

143

2.2 Orthosiphon stamineus Benth. Orthosiphon stamineus (syn.: O. grandiflorus, O. spicatus, O. aristatus; Lamiaceae) is a medicinal herb grown in Southeast Asia [11] (Fig. 5). In Myanmar, this plant is known locally as Se-cho or Myint-shwe, and the leaves are well known for their antidiabetic effects. A decoction of the leaves is effective in lowering blood glucose levels and alleviating urinary tract and renal diseases [12, 13]. The plant is also popular in Indonesia and Vietnam and has been used as a diuretic and to treat rheumatism, diabetes, hypertension, tonsillitis, epilepsy, menstrual disorders, gonorrhea, syphilis, renal calculus, gallstones, urinary lithiasis, edema, eruptive fever, influenza, hepatitis, jaundice, and biliary lithiasis [11, 14]. The highly oxygenated isopimarane-type diterpenoids, orthosiphols A–E, together with various monoterpenoids, triterpenoids, saponins, flavonoids, hexoses, organic acids, rosmarinic acid, chromene, and myo-inositol, have been reported from this plant when sourced from Indonesia and Okinawa, Japan [15–20]. In addition, two migrated pimarane-type and two isopimarane-type diterpenoids and a benzochromene have been isolated from the leaves of Indonesian O. aristatus (syn. O. stamineus) [21–23]. Two diterpenes with a novel carbon framework named staminane Fig. 5 Orthosiphon stamineus, known as Se-cho or Myint-shwe. The leaves are used as antidiabetic herbs in Myanmar

144

N. N. Win and H. Morita

(staminols A and B), two seco-staminanes (staminolactones A and B), a norstaminane (norstaminolactone), and five isopimarane-type diterpenes (orthosiphols F–J) have also been isolated from Vietnamese O. stamineus [24–26]. Phytochemical constituents of O. stamineus native to Myanmar were first reported in 2001 [27] and 2002 [28]. Thus, nine new highly oxygenated diterpenoids, orthosiphols K–Q (15–21), nororthosiphonolide A (22), and norstaminone A (23) (Fig. 6), six known diterpenoids, orthosiphols A (24) [16], B (25) [16], D (26) [15], and E (27) [15], orthosiphonone A (28) [21] and neoorthosiphol A (29) [22] (Fig. 6), four known flavonoids, tetramethylscutellarein (30) [29], eupatorin (31) [30], 5,6dihydroxy-7,4 -dimethoxyflavone (32) [31], and 6,7,8,3 ,4 -pentamethoxyflavone (33) [32]; and a ferulate derivative, (4-hydroxyphenyl)ethyl (E)-ferulate (34) [33] (Fig. 7), have been shown to be constituents of O. stamineus in Myanmar. Based on Fig. 6 Structures of isopimarane-type diterpenes 15–22 and 24–28 and staminane-type diterpenes 23 and 29 isolated from the chloroform-soluble fraction of the aerial parts of O. stamineus

R5 R4O OBz R1O

O OH

R2O

OAc R3

15 (orthosiphol K) R1 = R2 = R3 = R5= H, R4 = Bz 16 (orthosiphol L) R1 = Ac, R2 = R3 =H, R4 = Bz, R5= OH 19 (orthosiphol O) R1 = Ac, R2 = Bz, R3 = R4 = R5= H 20 (orthosiphol P) R1 = Ac, R2 = R5= H, R3 = OH, R4 = Bz 24 (orthosiphol A) R1 = Ac, R2 = R3 = R5 = H, R4= Bz 25 (orthosiphol B) R1 = R3 = R5 = H, R2 = Ac, R4 = Bz O

O OBz AcO

O OH

1

R O

OR

2

AcO

O OH OAc

O

21 (orthosiphol Q)

17 (orthosiphol M) R = H, R = Ac 18 (orthosiphol N) R1 = Bz, R2 = H 28 (orthosiphonone A) R1 = Bz, R2 = Ac 1

2

O BzO OH O

OBz O OH

O

OAc

22 (nororthosiphonolide A)

AcO

O

OAc

HO

23 (norstaminone A)

BzO RO

O

BzO OBz O OH

AcO

OAc

HO

26 (orthosiphol D) R = Ac 27 (orthosiphol E) R = H

OH O OH OAc

29 (neoorthosiphol A)

Bioactive Compounds from Medicinal Plants in Myanmar

145

R4 O

O

R1

O

O

O

OH

2

R O R3

O

30 (tetramethylscutellarein) R1 = R4 = H, R2 = Me, R3 = OMe 31 (eupatorin) R1 = H, R2 = Me, R3 = R4 = OH 32 (5,6-dihydroxy-7,4'-dimethoxyflavone) R1 = R2 = R4 = H, R3 = OH 33 (6,7,8,3',4'-pentamethoxyflavone) R1 = R4 = OMe, R2 = Me, R3 = H

O OH

34 (4-hydroxyphenyl)ethyl (E)-ferulate)

Fig. 7 Structures of flavonoids 30–33 and ferulate derivative 34 isolated from a chloroform-soluble fraction of the aerial parts of O. stamineus

the coexistence of isopimarane-type diterpenes 15–22 and 24–28 and staminane-type diterpenes 23 and 29, Awale et al. and Shibuya et al. proposed that isopimarane-type diterpenes are biosynthesized from staminane-type diterpenes via migration of the vinylic group from C-13 to C-12 [21, 24]. Involvement of a Baeyer-Villiger-type oxidation in the biosynthesis of norstaminone A (22) was also expected [24]. The antiproliferative activity against highly metastatic liver murine colon 26L5 carcinoma and human HT-1080 fibrosarcoma cells has been assessed for 15–29 [27, 28]. However, the assays revealed that diterpenes 15–29 displayed generally weak activity against both cell lines. Orthosiphol K (15), norstaminone A (23), and orthosiphol D (26) were found to be the most potent inhibitors, with ED50 values of 13.8, 12.8, and 16.0 μg/cm3 for the former cell line and 21.8, 23.2, and 23.0 μg/cm3 for the latter cell line, respectively. In contrast, flavones 30–32, but not 33 and 34, were shown to exhibit more potent antiproliferative activity than the diterpenes tested. The highest activity was observed for 5,6-dihydroxy-7,4 -dimethoxyflavone (32), with ED50 values of 2.3 and 3.0 μg/cm3 against murine colon 26-L5 carcinoma and HT-1080 fibrosarcoma cell lines, respectively. Inhibition of nitric oxide (NO) production by diterpenes 15, 16, 18–22 and 24–29 in lipopolysaccharide (LPS)-activated J774.1 cells also has been documented. More potent activities than the positive controls (N G -monomethyl-l-arginine (l-NMMA) (IC 50 26.0 μM)) and polymyxin B (IC 50 27.8 μM) were reported for 16 (IC 50 25.1 μM), 20 (IC 50 22.8 μM), 24 (IC 50 10.5 μM), 25 (IC 50 20.5 μM), and 26 (IC 50 14.4 μM) [34].

2.3 Panax zingiberensis C.Y. Wu & Feng (Myanmar wild ginseng) Ginseng is the underground rhizomatous part of plants belonging to the genus Panax in the family Araliaceae. Panax is one of the most medicinally important genera in the Orient, where almost every species of the genus has been used as a source of medicine

146

N. N. Win and H. Morita

[35]. Panax species are distributed naturally in the Northern Hemisphere from the central Himalayas onward through China, Korea, and Japan to North America [36]. Twelve Panax species, including P. ginseng C.A. Meyer, P. japonicus C.A. Meyer, P. major Ting, P. notoginseng (Burkill) F.H. Chen, P. omeiensis J. Wen, P. pseudoginseng Wall., P. quinquefolius L., P. sinensis J. Wen, P. stipuleanatus H.T. Tsai & K.M. Feng, P. trifolius L., P. wangianus Sun, and P. zingiberensis C.Y. Wu & K.M. Feng, are included in this genus [35]. On the market, three types of ginseng, fresh, white, and red, are available. White ginseng is made by peeling fresh ginseng roots and drying them without steaming. In contrast, red ginseng is made by steaming and drying fresh ginseng, suggesting a chemical transformation by heat to preserve the ginseng for an extended period of time [37, 38]. A decoction of the root of P. ginseng has been used traditionally both as a tonic for the restoration of strength and a panacea (hence the genus name Panax, meaning all-healing) [39]. It has also been suggested that the life-prolonging effects of ginseng may be due to the preventive activity of ginseng against the development of cancer. Administration of ginseng tablets has reportedly increased resistance to the adverse effects of antineoplastic agents and protects blood-forming organs [40]. There is a long-standing perception in Korea and China that wild P. ginseng has greater pharmacological activity than cultured P. ginseng. In fact, wild P. ginseng is sold at higher prices than cultured P. ginseng in these two countries. However, because of continual harvesting over thousands of years, the natural sources of this species have become almost exhausted, and wild plants are rarely available [41]. The chemical constituents of plants of Panax species are related to their morphology, which can be divided tentatively into two groups [42]. The first group contains P. ginseng and P. quinquefolius, and their underground parts are carrot-like roots with small rhizomes, containing dammarane triterpene saponins with minor amounts of oleanolic acid saponins. The second group includes P. japonicus, P. japonicas var. major, P. pseudo-ginseng subsp. himalaicus and P. zingiberensis. These species are grown in the eastern Himalayas and in southwestern China. The underground parts are large rhizomes with small roots that contain not only dammarane triterpene saponins but also a high content of a variety of oleanolic acid saponins [42, 43]. In 2001, a sample of wild ginseng from Myanmar (Fig. 8) was collected at the Par Moe Ne Water Spring area in Taunggyi, Shan State, at an altitude of 1500 m above sea level [44]. It is a perennial herb with an erect stem 30–50 cm in height, palmate leaves, and a large horizontal rhizome with small roots. The morphological characteristics are similar to those of the Panax species that grow in eastern Himalaya and southwestern China, such as P. japonicus, P. japonicus var. major, and P. pseudoginseng subsp. himalaicus. The isolation of the chemical constituents of a 60% hot EtOH extract by various chromatographic techniques afforded seven saponins, including three dammarane triterpene saponins, namely, ginsenosides Rg1 (35) [45] Rh1 (36) [46], and Rb1 (37) [47], and four oleanolic acid saponins, ginsenoside Ro (38) [47], chikusetsusaponins IV (39) [48] and IVa (40) [49], and zingibroside R1 (41) [50] (Fig. 9). The occurrence of both dammarane triterpene saponins and oleanolic acid saponins is supportive of Myanmar wild ginseng as a member of the

Bioactive Compounds from Medicinal Plants in Myanmar

147

Fig. 8 Fresh rhizome of Panax zingiberensis R3 O OH COOR2

R1 O

R1O R2

35 (ginsenoside Rg1) R1 = H, R2 = O-glc, R3 = glc 36 (ginsenoside Rh1) R1 = H, R2 = O-glc, R3 = H 37 (ginsenoside Rb1) R1 = glc2-glc, R2 = H, R3 = glc6- glc

38 (ginsenoside Ro) R1 = glcUA2-glc, R2 = glc 39 (chikusetsusaponin IV) R1 = glcUA4-ara(f), R2 = glc 40 (chikusetsusaponin IVa) R1 = glcUA, R2 = glc 41 (zingibroside R1) R1 = glcUA2-glc, R2 = H glc: β-D-glucopyranosyl glcUA: β-D-glucuronic acid ara(f): α-L-arabinofuranosyl

Fig. 9 Structures of dammarane triterpene saponins 35–37 and four oleanolic acid saponins 38–41 isolated from a 60% hot ethanol extract of P. zingiberensis rhizomes

second group of Panax species that includes P. japonicus, P. japonicus var. major, P. pseudo-ginseng subsp. himalaicus, and P. zingiberensis. The isolation of zingibroside R1 (41), which was reported only from P. zingiberensis [50], pointed to the identity of Myanmar wild ginseng as being P. zingiberensis. Further analysis using its 18S rRNA gene and matK gene sequences additionally confirmed the scientific name of Myanmar wild ginseng as P. zingiberensis [44]. Since P. zingiberensis was listed in

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the 1997 IUCN Red List of Threatened Plants as having an endangered status [51], full-scale inventory and resource assessment for this species is highly recommended.

2.4 Caesalpinia crista L. Caesalpinia crista belongs to the family Caesalpiniaceae, which was formerly part of the family Leguminosae (or Fabaceae). It is a well-known medicinal plant that is distributed widely in tropical and subtropical regions of Southeast Asia. The plant is known locally as Ka-lain in Myanmar (Fig. 10), and its seeds are used as an anthelmintic, antipyretic, anti-inflammatory, and antimalarial agent [52]. Several parts of this species, such as the seeds, nuts and root bark, have been used in folk medicine to treat intermittent fevers, asthma, and colic. The tender leaves are considered to be efficacious against disorders of the liver, and the oil expressed from them is used to treat convulsions, palsy, and similar nervous complaints. A decoction of the seeds is used effectively for malaria, edema, fever, and inflammation [53]. In Indonesia, it is known as Bagore, and a decoction of the roots has been used as a tonic and for the treatment of rheumatism and backache [54]. Analgesic [55], adaptogenic, antiulcer, anthelmintic, antibacterial, insecticidal,

Fig. 10 Caesalpinia crista, known as Ka-lain in Myanmar. Various parts are used in folk medicines in Myanmar

Bioactive Compounds from Medicinal Plants in Myanmar

149

antifungal, anti-inflammatory, antipyretic, antioxidant, antiproliferative, antiviral, immunomodulatory, and immunosuppressive activities have been reported from plants in the Caesalpinia genus [56]. Several classes of phytoconstituents, such as flavonoids, diterpenes, and steroids, have been reported from plants in this genus [56]. Caesalpinia spp. are rich sources of cassane-type diterpenes, inclusive of those without a C-5 hydroxy group substituent or furan ring, cassane-type furanoditerpenes, and norcassane-type diterpenes. Antimalarial [57, 58], antiviral [59], and cytotoxic activities of cassane-type furanoditerpenes, isolated from Indonesian C. crista, have been reported [60]. The seed kernels of Caesalpinia crista from Myanmar were studied initially in 2004 [61–63]. As in the case of other members of the genus Caesalpinia, the seed kernels of C. crista from Myanmar have proved to be a good source of cassanetype diterpenes. Investigation of the chemical constituents of the CH2 Cl2 extract prepared from the seed kernels of C. crista in Myanmar revealed the occurrence of O

O R

1

OAc

R5 R6

R

2

R4

OH R3

44 (caesalpinin MC) AcO

OH

42 (caesalpinin MA) R1 = R2 = OAc, R3 = R4 = R5 = H, R6 = Me 43 (caesalpinin MB) R1 = OAc, R2 = R3 = R4 = R6 = H, R5 = COOMe 46 (caesalpinin MF) R1= R2 = OAc, R3 = R4 = R6 = H, R5 = COOMe 47 (caesalpinin MG) R1 = R3 = R4 = OAc, R2 = R6= H, R5 = COOMe 48 (caesalpinin MH) R1 = R3 = OAc, R2 = R6 = H, R4 = OH, R5 = COOH 49 (caesalpinin MI) R1 = R2 = R3 = R5 = H, R4 = OH, R6 = Me O

O

O

OAc R1

R2

OH 3 R

OH OAc

45 (caesalpinin MD) R1 = R3 = OAc, R2 = H 55 (caesalpinin MP) R1 = R2 = R3 = H

OAc

54 (caesalpinin MO)

O

O OAc

OAc

R1 R2

OH R1

R2

OAc O OAc

50 (caesalpinin MJ) R1 = H, R2= OAc 52 (caesalpinin MM) R1 = Me, R2 = OH 51 (caesalpinin MK) R1 = OAc, R2 = H 53 (caesalpinin MN) R1 = OH, R2 = Me

Fig. 11 Structures of new cassane-type furanoditerpenes 42–55 from the seed kernels of C. crista

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N. N. Win and H. Morita

O O

AcO

OH

HO

OAc

56 (caesalpinin ME)

OH

57 (caesalpinin ML)

O

O

OAc

R

R

AcO

O

OH

58 (norcaesalpinin MA) R = H 59 (norcaesalpinin MB) R = OAc

OH OAc

OAc

60 (norcaesalpinin MC) R = α-OAc, β-H, 61 (norcaesalpinin MD) R = O

Fig. 12 Structures of new cassane-type diterpenes 56 and 57 without a C-5 hydroxy substituent and furan ring, and of 16-norcassane-type diterpenes 58 and 59 and 17-norcassane-type diterpenes 60 and 61 from the seed kernels of C. crista

20 unprecedented cassane-type diterpenes (42–53), including 14 cassane-type furanoditerpenes, named caesalpinins MA–MD (42–45), MF–MK (46–51), and MM– MP (52–55) (Fig. 11); two cassane-type diterpenes without any C-5 hydroxy group substituent or furan ring (caesalpinins ME (56) and ML (57)); two 16-norcassanetype diterpenes (norcaesalpinins MA (58) and MB (59)), and two 17-norcassane-type diterpenes (norcaesalpinins MC (60) and MD (61)) (Fig. 12). Also identified were 27 known cassane-type furanoditerpenes, caesalmin C (62) [59], 1-deacetoxy-1oxocaesalmin C (63), 1-deacetylcaesalmin C (64) [64], caesalpinins C (65) and F (66) [58], 14(17)-dehydrocaesalmin F (67) [65], caesalpinin J (68) [66], ζ-caesalpin (69) [67], caesalmin E (70) [59], caesalpinin E (71) [58], caesalpinins K (72), M (73), and N (74) [66], bonducellpin C (75) [68], 7-acetoxybonducellpin C (76) [68] (Fig. 13), caesalmins B (77) [69] and G (78) [59], caesalpinins D (79) [58], H (80), I (81), and O (82) [66], norcaesalpinins B (83) [57] and E (84) [58], caesaldekarin e (85) [70], 2-acetoxycaesaldekarin E (86) [65], 2-acetoxy-3-deacetoxycaesaldekarin e (87) [71], and 6-acetoxy-3-deacetoxycaesaldekarin e (88) [72] (Fig. 14). Caesalpinin MC (44) was found to be the first example of a cassane-type furanoditerpene with a dihydrofuran ring. Furthermore, caesalpinins MM (52) and MN (53) are the first members of a rare group of cassane-type furanoditerpenes with a rearranged carbon skeleton, for which no additional representatives have been reported subsequently. The chemical constituents of C. crista from Myanmar also include 1-deacetoxy-1oxocaesalmin C (63) and 1-deacetylcaesalmin C (64), which are natural products that were previously synthesized. Interestingly, significant inhibition of the growth of the malaria parasite P. falciparum FCR-3/A2 clone in vitro (IC 50 value of 0.21 μg/cm3 ) was reported from the

Bioactive Compounds from Medicinal Plants in Myanmar Fig. 13 Structures of known cassane-type furanoditerpenes 62–76 from the seed kernels of C. crista

151 O 1

R R2 R3

R6 R5

OH R4

62 (caesalmin C) R1 = α-OAc, β-H, R2 = R3 = H, R4 = R5 = OAc, R6 = CH2 63 (1-deacetoxy-1-oxocaesalmin C) R1 = O, R2 = R3 = H, R4 = R5 = OAc, R6 = CH2 64 (1-deacetylcaesalmin C) R1 = α-OH, β-H, R2 = R3 = H, R4 = R5 = OAc,, R6 = CH2 65 (caesalpinin C) R1 = α-OAc, β-H, R2 = R4 = R5 = H, R3 = OAc, R6 = CH2 66 (caesalpinin F) R1 = O, R2 = R3 = R5 = H, R4 = OAc, R6 = α-H, β-COOMe 67 (14(17)-dehydrocaesalmin F) R1 = α-OAc, β-H, R2 = R3 = OAc, R4 = R5 = H,R6 = CH2 68 (caesalpinin J) R1 = O, R2 = R3 = H, R4 = R5 = OAc, R6 = α-H, β-COOMe 69 (ζ−caesalpin) R1 = α-OAc, β-H, R2 = R3 = H, R4 = OH, R5 = OAc, R6 = CH2 O OAc

R3 4

R OH R1

R2

70 (caesalmin E) R1 = R2 = OAc, R3 = OH, R4 = Me 71 (caesalpinin E) R1= OAc, R2 = R4 = H, R3 = COOMe 72 (caesalpinin K) R1 = R3 = H, R2 = OH, R4 = Me 73 (caesalpinin M) R1 = OH, R2 = OAc, R3= COOMe, R4 = H 74 (caesalpinin N) R1 = R4 = H, R2 = OH, R3 = CHO 75 (bonducellpin C) R1 = R4 = H, R2 = OH, R3 = COOMe 76 (7-acetoxybonducellpin C) R1 = R4 = H, R2 = OAc, R3 = COOMe

CH2 Cl2 extract of the seed kernels of C. crista grown in Myanmar [73], which is in good agreement with the traditional use of this part as an antimalarial agent locally. Furthermore, more potent in vitro activities, when compared with the well-known antimalarial drug chloroquine (IC 50 , 0.29 μM), were reported for diterpenes 67 (IC 50 , 0.20 μM), 75 (IC 50 , 0.12 μM), 83 (IC 50 , 0.26 μM), 84 (IC 50 , 0.090 μM), and 87 (IC 50 , 0.098 μM). Consequently, an acetoxy group at C-1 and a hydroxy group at C-7 have been proposed as important functionalities in norcassane-type diterpenes for enhancing growth inhibition of the malaria parasite P. falciparum FCR-3/A2 clone. The observed inhibitory activities suggest that these compounds could be the effective active components of the seed kernels of C. crista as an antimalarial drug. Further accumulation of scientific evidence is awaited concerning the traditional use of the seed kernels of C. crista as an antimalarial therapy.

152

N. N. Win and H. Morita O

O

1

R

OAc O

O

O

R1

OH 2 R 77 (caesalmin B) R1 = α-OAc, β-H, R2 = H 78 (caesalmin G) R1 = α-OH, β-H, R2 = H 79 (caesalpinin D) R1 = α-OAc, β-H, R2 = OAc 80 (caesalpinin H) R1 = α-OH, β-H, R2 = OAc 81 (caesalpinin I) R1 = O, R2 = OAc 82 (caesalpinin O) R1 = α-OAc, β-H, R2 = OH

2

OH

R

83 (norcaesalpinin B) R1 = OAc, R2 = H 84 (norcaesalpinin E) R1 = H, R2 = OH

O OAc R1

R2

OH 3 R

85 (caesaldekarin e) R1 = R3 = H, R2 = OAc 86 (2-acetoxycaesaldekarin e) R1 = R2 = OAc, R3 = H 87 (2-acetoxy-3-deacetoxycaesaldekarin e) R1 = OAc, R2 = R3 = H 88 (6-acetoxy-3-deacetoxycaesaldekarin e) R1 = R2 = H, R3 = OAc

Fig. 14 Structures of known cassane-type furanoditerpenes 77–88 from the seed kernels of C. crista

2.5 Elephantopus scaber L. Elephantopus scaber is a small herb belonging to the family Asteraceae (previously named Compositae). This plant is distributed in the neotropics, and in Europe, Asia, Africa and Australia [74–81], and can be found throughout Myanmar. In Myanmar, it is known locally as Taw-mon-lar and is prescribed as an antiviral, antimalarial, and anti-hepatitis remedy by traditional medicinal practitioners (Fig. 15) [82]. E. scaber has been used widely not only in Myanmar traditional medicine but also in those of many other countries for the treatment of various diseases. Thus, a decoction of the whole plant or the roots of E. scaber is used in countries such as India, China, Vietnam, the Philippines, Thailand, Madagascar, Nepal, and Brazil as an anti-inflammatory, antipyretic, diuretic, antitussive, antibiotic, emollient, and tonic [74–81]. The reported major constituents are sesquiterpene lactones, which are a structurally highly diverse major group of terpenoids found as characteristic constituents of the family Asteraceae [83]. Other compounds, including triterpenoids, steroids, and their glycosides, flavonoids, phenolic compounds, long-chain hydrocarbons, and components of the essential oil of E. scaber have also been reported by several research groups [84–86]. The first chemical constituents of E. scaber from Myanmar were reported in 2004 [87]. Two new sesquiterpene lactones, 17,19-dihydrodeoxyelephantopin (89)

Bioactive Compounds from Medicinal Plants in Myanmar

153

Fig. 15 Elephantopus scaber known as Taw-mon-lar in Myanmar. Typically, the whole plant or roots of E. scaber are mainly used in traditional medicine in Myanmar O

O O

O

O

O O

O

O

O O

O

89 (17,19-dihydrodeoxyelephantopin)

90 (iso-17,19-dihydrodeoxyelephantopin)

O O

O O

O O 91 (deoxyelephantopin)

Fig. 16 Structures of sesquiterpene lactones 89–91 isolated from ethanol and acetone extracts of E. scaber

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Fig. 17 Structures of fatty acid derivatives 92–95 isolated from ethanol and acetone extracts of E. scaber

O O 92 (ethyl hexadecanoate)

O O 93 (ethyl-9,12-octadecadienoate) O 94 (ethyl-(Z)-9-octadecenoate)

O O O

95 (ethyl octadecanoate)

Fig. 18 Structures of triterpene 96 and sterols 97 and 98 isolated from ethanol and acetone extracts of E. scaber HO HO 96 (lupeol)

97 (stigmasterol)

OH HO HO

O O OH 98 (stigmasterol glucoside)

and iso-17,19-dihydrodeoxyelephantopin (90) [87] (Fig. 16), a known sesquiterpene lactone, deoxyelephantopin (91) [88] (Fig. 16), four fatty acid derivatives, ethyl hexadecanoate (92) [89], ethyl-9,12-octadecadienoate (93) [90], ethyl-(Z)-9octadecenoate (94) [91], and ethyl octadecanoate (95) [92] (Fig. 17), the triterpene lupeol (96) [93], and two sterols, stigmasterol (97) [94] and its glucoside 98 [94] (Fig. 18), were isolated from ethanol and acetone extracts of the whole plants of E. scaber. Among these, 17,19-dihydrodeoxyelephantopin (89) is an epimer of iso17,19-dihydro-deoxyelephantopin (90) at C-2. In turn, deoxyelephantopin (91) is a structural analog of 17,19-dihydrodeoxyelephantopin (89) with a methacryloxy

Bioactive Compounds from Medicinal Plants in Myanmar

155

moiety at C-10 instead of an isobutoxy moiety as in 89, which is a potential antitumor agent and anti-inflammatory compound [95–102]. Three in vitro bioassays, using cytotoxicity, antibacterial, and antifungal assays, were conducted with 17,19-dihydrodeoxyelephantopin (89) and iso-17,19-dihydrodeoxyelephantopin (90) (isolated from E. scaber collected in Myanmar), together with deoxyelephantopin (91) [87]. The cytotoxicity testing against twelve cell lines derived from bladder, central nervous system, colon, gastric, head and neck, lung, mammary, ovarian, pancreatic, prostate, renal, and uterine cancers, as well as two cell lines established from melanomas and pleural mesothelioma, revealed that 17,19-dihydrodeoxyelephantopin (89) and iso-17,19-dihydro-deoxyelephantopin (90) showed activities with mean IC 70 values of 4.0 μg/cm3 and 4.3 μg/cm3 against the melanoma-derived cell line MEXF 394NL, respectively, compared with a mean IC 70 value of 1.1 μg/cm3 for 91. This biological testing work showed differences in the selectivities of 17,19-dihydrodeoxyelephantopin (89), iso-17,19-dihydrodeoxyelephantopin (90), and deoxyelephantopin (91) according to the cancer cell lines used, depending on their different functional groups present [87]. Notably, highly effective activity against the RXF 944L renal cancer cell line was found for 89, whereas marked activity for the LXFL 529L large cell lung cancer cell line was indicated for 90, the diastereomer of 89. In contrast, pronounced selective activity in the MAXF 401NL mammary cancer cell line was shown by 91. An antibacterial assay using Staphylococcus aureus and Bacillus subtilis and an antifungal assay using Candida albicans also revealed that all compounds exhibited 11-mm diameter inhibition zones at 100 μg/well against S. aureus, whereas the compounds did not show any inhibitory activities against B. subtilis or C. albicans at this concentration level [87].

2.6 Phyllanthus niruri L. Phyllanthus niruri is a small annual herb belonging to the family Euphorbiaceae (Fig. 19). It is a popular indigenous medicinal herb found in the Amazon rainforest and other tropical areas, including those in Southeast Asia, Southern India, and China [103]. P. niruri has a long history in herbal medicine systems such as Indian Ayurveda, traditional Chinese medicine, and Indonesian Jamu. The whole plant is used as a remedy for many conditions, such as dysentery, influenza, vaginitis, tumors, diabetes, jaundice, kidney stones, and dyspepsia, in addition to its use as a diuretic. The plant is also useful for treating hepatotoxicity, hepatitis B, hyperglycemia, and viral and bacterial diseases [104]. In Malaysia, P. niruri is used internally for diarrhea, kidney disorders, gonorrhea, and coughs [105]. In turn, P. niruri has been used as an antiviral and antimalarial remedy as well as for the treatment of jaundice and hepatitis in Myanmar, where it is known locally as Taung-zi-phyu [106]. A number of lignans [107–109], alkaloids [110, 111], flavonoids [112], tannins [113], benzenoids [113, 114], coumarins [115], terpenes [116, 117], phthalates [118], saponins, and phenylpropanoids [119] have been reported as the active constituents of the leaves,

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Fig. 19 Phyllanthus niruri, known as Taung-zi-phyu in Myanmar. The whole plant is used in traditional medicine in Myanmar

stems, and roots of this plant. Researchers across the world have evaluated various types of pharmacological parameters exhibited by extracts of P. niruri, beginning in the mid-1960s. Thus, these extracts exhibit a broad spectrum of biological activities, including antihypoglycemic [120], anti-HIV [121, 122], antiplasmodial [123, 124], lipid lowering [125], analgesic [126], antinociceptive [127], protection against liver damage [128], and reduction of urinary calcium [129]. The active constituents shown to date are the lignans, phyllanthin, and hypophyllanthin, with antihepatotoxic effects [130], and glycosides such as quercitrin and geraniin, which along with ellagic acid exhibit aldose-reductase activities [113, 131]. A phytochemical study on Phyllanthus niruri from Myanmar was first reported in 2006 [132]. A combination of column chromatography on both silica gel and Sephadex LH-20 of the 70% ethanol extract of P. niruri led to the isolation of a new flavone sulfonic acid, niruriflavone (99), together with nine known compounds, namely, two flavonoid glycosides, isoquercetin (100) [133] and quercetin-3-Oβ-d-glucopyranosyl(1→4)-α-rhamnopyranoside (101) [134] (Fig. 20), two ellagitannins, corilagin (102) and isocorilagin (103) [135] (Fig. 21), and the additional secondary metabolites, 6,10,14-trimethyl-2-pentadecanone (104) [136], gallic acid (105) [137], hypophyllanthin (106) [107], brevifolin carboxylic acid (107) [138], and methyl brevifolin carboxylate (108) [139] (Fig. 22). The occurrence of a flavone-6-sulfonic acid from Nature was reported for the first time. Use of an ABTS (2,2 -azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium

Bioactive Compounds from Medicinal Plants in Myanmar

157 OH OH

O O

HO O

HO

O HO3S

OH OH

O

CH2OH

O HO

OH

OH

O

99 (niruriflavone)

100 (isoquercetin) OH OH O

HO

OH O HO OH

O

O

101 (quercetin-3-O-β-D-glucopyranosyl(1

OH O OH O

OH OH

4)-α-rhamnopyranoside)

Fig. 20 Structures of flavone sulfonic acid 99 and flavonoid glycosides 100 and 101 isolated from a 70% ethanol extract of P. niruri OH

OH

OH

OH O

O O HO

O

O OH

HO O

HO

O

O

OH

O

O HO

O

HO O

HO

OH

O OH O

OH

OH HO

OH OH

102 (corilagin)

HO

OH OH

103 (isocorilagin)

Fig. 21 Structures of ellagitannins 102 and 103 isolated from the 70% ethanol extract of P. niruri

salt) cation radical reduction assay revealed that niruriflavone (99) showed radicalscavenging properties at concentration levels between 20 and 100 μM. However, niruriflavone (99) did not exhibit any antiperoxidant effects at 10 μM in an assay using the bioluminescent dinoflagellate Lingulodinium polyedrum as a test organism, although gallic acid (105) and quercetin-3-O-β-d-glucopyranosyl(1→4)α-rhamnopyranoside (101) were active in this regard [132].

158

N. N. Win and H. Morita O O

HO

OH

HO OH 105 (gallic acid)

104 (6,10,14-trimethyl-2-pentadecanone)

O

O

OH

O O

HO

OH

ROOC

O O O 106 (hypophyllanthin)

O

O

O 107 (brevifolin carboxylic acid) R = H 108 (methyl brevifolincarboxylate) R = Me

Fig. 22 Structures of secondary metabolites 104–108 isolated from the 70% ethanol extract of P. niruri

2.7 Streptocaulon tomentosum Wight & Arn. Streptocaulon tomentosum belongs to the family Asclepiadaceae. It is distributed in the Bago, Mandalay, and Yangon Regions and Mon State of Myanmar. The plant is known locally as Myin-sa-gon-ni, meaning the red variety of Myin-sa-gon (Fig. 23). In Myanmar, two Streptocaulon species, S. tomentosum and S. griffithii, occur. Of these, S. griffithii is referred to as Myin-sa-gon-phyu, meaning the white variety of Myin-sa-gon [140]. The roots of S. tomentosum have been used in Myanmar traditional medicine for the alleviation of cancer, dysentery, and stomachache, and the leaves are used externally for the treatment of snake poisoning and abscesses [141]. There has not been much scientific work performed on S. tomentosum other than the studies on Vietnamese S. juventas by Ueda et al. [142, 143] and Pham and Tran [144]. In Vietnam, S. juventas is called Ha thu o trang, and its roots are used as a tonic for various conditions, such as anemia, chronic malaria, rheumatism, menstrual disorders, neurasthenia, and dyspepsia, and is a substitute of Ha thu o do (the roots of Polygonum multiflorum Thunb., Polygonaceae). The roots of S. tomentosum have been reported to be abundant in cardenolides. Hemiterpenoids, phenylpropanoids, and phenylethanoids have also been isolated from the roots. Pharmacological studies of cardenolides have demonstrated that these compounds selectively and potently inhibit proliferation of the growth of the HT-1080 and A549 cell lines through the induction of apoptosis [142, 143]. Several investigations of the roots of S. tomentosum in Myanmar have led to the purification of 21 compounds, including triterpenoids, cardenolides, lignans, and steroidal saponins from this plant part [141, 145, 146]. Two new cardenolides, 17α-H-periplogenin-3-O-β-d-glucopyranosyl-(1→4)-2-O-acetyl-3-O-methyl-βfucopyranoside (109) [145] and 17β-H-periplogenin-3-O-β-d-digitoxoside (110),

Bioactive Compounds from Medicinal Plants in Myanmar

159

Fig. 23 Streptocaulon tomentosum (whole plant), known as Myin-sa-gon-ni in Myanmar. The roots are used in Myanmar traditional medicine

and a new pregnane glycoside, 5 -pregnene-3β,16α-diol-3-O-[2,4-O-diacetylβ-digitalopyranosyl-(1→4)-β-d-cymaropyranoside]-16-O-[β-d-glucopyranoside] (111) [141] (Fig. 24) were reported from the roots, together with seven known cardenolides, 17α-H-periplogenin (112), 17α-H-periplogenin-3-O-β-d-digitoxose (113), 17α-H-periplogenin-3-O-β-d-cymarose (114) [142, 147], 17α-H-digitoxigenin (115), 17α-H-digitoxigenin-3-O-β-d-digitoxoside (116) [148], 17β-H-periplogenin (117) [149], and 17β-H-periplogenin-3-O-β-d-cymarose (118) [142] (Fig. 24). In addition, eight known triterpenes, α-amyrin acetate (119) [150], 2α,3α,23trihydroxyurs-12-en-28-oic acid (120), β-amyrin acetate (121) [151], 2α,3βdihydroxyurs-12-en-28-oic acid (122), 2α,3β-dihydroxyolean-12-en-28-oic acid (123), 2α,3β,23-trihydroxyurs-12-en-28-oic-acid (124), 2α,3β,23-trihydroxyolean12-en-28-oic-acid (125) [152–154], and lupeol acetate (126) [155] (Fig. 25) were identified. The known sterol, cycloartenol (127) [156], and the furofuranlignan, 8-hydroxypinoresinol (128) [157], were also isolated (Fig. 26). Among these compounds, six cardenolides (109, 110, 112–115) exhibited antiproliferative activity against the MCF7 human breast cancer cell line (IC 50 values < 1–15.3 μM after two days of incubation, and 100 μM). Weak antiproliferative activity against the L929 cell line (IC 50 79.4 μM after five days incubation) was recorded also for lupeol acetate (115) [158]. Furthermore, an antiproliferative assay of the four cardenolides 110, 113, 114, and 115 on cellular viability and the cell cycle in the U937 and TUR (TPA-U937resistant, a differentiation-resistant subclone of the U937 myeloid leukemia cells) human leukemic cell lines, demonstrated that all four compounds led to induction of apoptosis at high concentrations (>10 μM) in both cell lines, whereas TUR cells were more sensitive. In this study, compound 113 exhibited the most potent activity against TUR cells versus U937 cells at a concentration of 1 μM. In addition, the assay also revealed that compounds 113 and 114 caused a blockade at the G2/M-phase at 100 μM and 10 μM in both cell lines, whereas compounds 110 and 115 caused a blockade at the G2/M-phase at 100 μM [158].

Bioactive Compounds from Medicinal Plants in Myanmar

161 R4

R2

R6

R1

R4

R1

R2 R3

R5

R3

119 (α-amyrin acetate) R1 = H, R2 = β-OAc, R3 = R4 = Me 121 (β-amyrin acetate) R1 = H, R2 = OAc, R3 = R5 = R6 = Me, R4 = H 120 (2α,3α,23-trihydroxyurs-12-en-28-oic acid) 122 (2α,3β -trihydroxyurs-12-en-28-oic acid) R1 = R2 = OH, R1 = OH, R2 = α-OH, R3 = CH2OH, R4 = COOH R3 = R4 = Me, R5 = H, R6 = COOH 123 (2α,3β -dihydroxyolean-12-en-28-oic acid) R1 = R2 = OH, R3 = R5 = Me, R4 = H, R6 = COOH 124 (2α,3β,23-trihydroxyurs-12-en-28-oic acid) R1 = R2 = OH, R3 = CH2OH, R4 = Me, R5 = H, R6 = COOH 125 (2α,3β,23-dihydroxyolean-12-en-28-oic acid) R1 = R2 = OH, R3 = CH2OH, R4 = H, R5 = Me, R6 = COOH

AcO 126 (lupeol acetate)

Fig. 25 Structures of terpenoids 119–126 isolated from the roots of S. tomentosum

Fig. 26 Structures of sterol 127 and furofuranlignan 128

O OH O OH O

HO

O

HO 127 (cycloartenol)

128 (8-hydroxypinoresinol)

Notably, Khine et al. revealed differences in the profiles of the chemical components of the air-dried roots of S. juventas and S. tomentosum [141, 145, 146]: 16 cardenolides, two hemiterpenoids, two phenylpropanoids, and a phenylethanoid have been reported from the air-dried roots of S. juventas [142] together with two triterpenoids [144], whereas nine cardenolides, eight triterpenes, a lignan and a pregnane glycoside (109–128) were isolated from the air-dried roots of S. tomentosum [141]. Three major differences were found: (I) the major cardenolides (digitoxigenin-3-O-[O-β-glucopyranosyl-(1→6)-O-β-glucopyranosyl-(1→4)-3O-acetyl-β-digitoxopyranoside], digitoxigenin-3-O-[O-β-glucopyranosyl-(1→6)O-β-glucopyranosyl-(1→4)-O-β-digitalopyranosyl-(1→4)-β-cymaropyranoside], digitoxigenin-3-O-[O-β-glucopyranosyl-(1→6)-O-β-glucopyranosyl-(1→4)-βdigitoxopyranoside], digitoxigenin sophoroside, echujin, periplogenin glucoside, corchorusoside C), and the minor cardenolides (acovenosigenin A digitoxoside, acovenosigenin A, digitoxigenin gentiobioside) of S. juventas were absent in S.

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tomentosum; (II) The major triterpenoids (120, 124, 125) and minor triterpenoids (122, 123) of S. tomentosum were not found in S. juventas, with these compounds replaced in S. juventas by (4R)-4-hydroxy-3-(1-methylethyl)pentyl rutinoside, (R)-2-ethyl-3-methylbutyl rutinoside, two phenylpropanoids, and a phenylethanoid; (III) The lignan and pregnane glycoside constituents found in S. tomentosum were not detected in S. juventas. Khine et al. thus proposed that these plants could be classified into two subspecies [141, 146], although a previous report considered S. tomentosum and S. juventas synonyms [159]. Hence, further chemical investigations, as well as botanical and genetic studies, are highly recommended for these two taxa.

2.8 Curcuma comosa Roxb. Curcuma comosa belongs to the family Zingiberaceae. It is known locally as Sanwinyaing (Fig. 27), meaning the wild variety of Curcuma longa Sanwin. It is distributed in the Bago and Mandalay Regions of Mynmar [160]. Altogether, 24 Curcuma species are distributed widely throughout Myanmar. Among these various Curcuma species, C. comosa is an important medicinal plant that has been used to treat several diseases. For example, its rhizomes have been utilized to alleviate malaria fever in combination with Artemisia annua and Aristolochia tagala by Myanmar practitioners [146]. In Thailand, C. comosa is employed in indigenous medicine as an anti-inflammatory Fig. 27 Curcuma comosa known as Sanwin-yaing. The rhizome is used in Myanmar traditional medicine

Bioactive Compounds from Medicinal Plants in Myanmar

163

O

R O

O O

O

O

O

129 (1a,5,7a-trimethyl-1a,6a,7a,8,9,9a-hexahydrobisoxireno[4,5:8,9]cyclodeca[1,2-b]furan-6(2H)-one)

O

O

O 138 (curdione)

130 (3,6,10-trimethyl-7,8,11,11a-tetra-hydrocyclodeca[b]furan-2,5(4H,6H)-dione) R = H 131 (11a-hydroxy-3,6,10-trimethyl-7,8,11,11atetrahydrocyclodeca[b]furan-2,5(4H,6H)-dione-methane) R=O

O

O

O

139 (zederone)

O

O 140 ((1S,4S,5S,10S)-germacrone1(10),4(5)-diepoxide)

Fig. 28 Structures of germacrane-type sesquiterpenoids 129–131 and 138–140 isolated from C. comosa rhizomes

agent, and for the treatment of postpartum uterine bleeding and as an aromatic stomachic [161]. Sesquiterpenoids and curcumin and other phenolic diarylheptanoids have been reported as the major constituents of the Curcuma comosa rhizomes [161, 162]. Various biological activities, including antineoplastic effects [163], nematocidal activity [164], cytotoxicity [165], topoisomerase inhibition [166], antioxidant activity [167], protection against alcohol-induced liver toxicity [168], estrogenic activity [161], and antiparasitic activity against Plasmodium falciparum Welch. and Leishmania major Friedlin [169], have been reported for curcumin and its analogs. The chemical constituents of the rhizomes of C. comosa in Myanmar have been investigated for about 15 years. A total of 29 compounds (129–157) were characterized including nine new sesquiterpenoids (129–137), 1a,5,7a-trimethyl1a,6a,7a,8,9,9a-hexahydrobisoxireno[4,5:8,9]cyclodeca[1,2-b]furan-6(2H)-one (129), 3,6,10-trimethyl-7,8,11,11a-tetrahydrocyclodeca[b]furan-2,5(4H,6H)-dione (130), 11a-hydroxy-3,6,10-trimethyl-7,8,11,11a-tetrahydrocyclodeca[b]furan2,5(4H,6H)-dione-methane (131) (Fig. 28), 1,4-dihydroxy-1,4-dimethyl-7-(1methylethylidene)octahydroazulen-6(1H)-one-methane (132), 5,8-dihydroxy3,5,8-trimethyl-4a,5,6,7,7a,8,9,9a-octahydroazuleno[6,5-b]furan-2(4H)-one (133), 5,8-dihydroxy-3,5,8-trimethyl-4a,5,6,7,7a,8,9,9a-octahydroazuleno[6,5-b]furan2(4H)-one (134), 4a,8,9,9a-tetrahydroxy-3,5,8-trimethyl-4a,5,6,7,7a,8,9,9aoctahydroazuleno[6,5-b]furan-2(4H)-one (135), 7-(1-hydroxy-1-methylethyl)1,4-dimethyl-1,2,3,3a,4,5,8,8a-octahydroazulene-1,4-diol (136) (Fig. 29), and 7-isopropenyl-1,4a-dimethyldecahydronaphthalene-1,4-diol (137) [146] (Fig. 30). In addition, 20 previously reported secondary metabolites, namely, curdione (138) [170], zederone (139) [171], (1S,4S,5S,10S)-germacrone-1(10),4(5)-diepoxide (140) [170] (Fig. 28), curcumenol (141) [172], isocurcumenol (142) [173], alismoxide (143) [174], procurcumenol (144) [175], isoprocurcumenol (145) [176], isozedoarondiol (146), zedoarondiol (147) [177], zedoalactones A (148) and B (149) [178], zedoarolide B (150) [179], gajutsulactone B (151) [179] (Fig. 31),

164

N. N. Win and H. Morita OH

OH

OH O

O

O

HO

HO

HO

OH

OH O

O

HO

HO

five possible structures of 132 (1,4-dihydroxy-1,4-dimethyl-7(1-methylethylidene)octahydroazulen-6(1H)-one-methane) 3 R1 R R4

OH OH R

HO

2

OH

O

O

HO O

O

133 (5,8-dihydroxy-3,5,8-trimethyl-4a,5,6,7,7a,8,9,9aoctahydroazuleno[6,5-b]furan-2(4H)-one) R1 = R2 = α-H, R3 = OH, R4 = Me

135 (4a,8,9,9a-tetrahydroxy-3,5,8-trimethyl4a,5,6,7,7a,8,9,9a-octahydroazuleno[6,5-b]furan-2(4H)-one)

134 (5,8-dihydroxy-3,5,8-trimethyl-4a,5,6,7,7a,8,9,9aoctahydroazuleno[6,5-b]furan-2(4H)-one) R1 = R2 = β-H, R3 = Me, R4 = OH OH

OH

OH

136 (7-(1-hydroxy-1-methylethyl)-1,4-dimethyl1,2,3,3a,4,5,8,8a-octahydroazulene-1,4-diol)

Fig. 29 Structures of new guaiane-type sesquiterpenoids 132–136 isolated from C. comosa rhizomes OH

O

OH

O H

OH

137 (7-isopropenyl-1,4a-dimethyldecahydro-naphthalene-1,4-diol)

152 (bisacumol)

153 (curcumenone)

Fig. 30 Structures of the eudesmane-type sesquiterpenoid 137, bisabolane-type sesquiterpenoid 152, and carabrane-type sesquiterpenoid 153 isolated from C. comosa rhizomes

Bioactive Compounds from Medicinal Plants in Myanmar

165 OH

OH

O

O

OH OH

141 (curcumenol)

142 (isocurcumenol)

143 (alismoxide) R1

O

O

O HO

OH

OH 144 (procurcumenol)

145 (isoprocurcumenol)

146 (isozedoarondiol) R1 = β-H, R2 = Me, R3 = OH 147 (zedoarondiol) R1 = α-H, R2 = OH, R3 = Me

OH OH

OH R

HO

R2 R 3

HO

O

O O

148 (zedoalactone A) R = α-H 150 (zedoarolide B) R = β-OH

O 149 (zedoalactone B)

O

O

151 (gajutsulactone B)

Fig. 31 Structures of known guaiane-type sesquiterpenoids 141–151 isolated from C. comosa rhizomes

bisacumol (152) [180], curcumenone (153) [181] (Fig. 30), curcumin (154), demethoxycurcumin (155), bisdemethoxycurcumin (156) [164], and (3S,5S)-3,5diacetoxy-1,7-bis(3,4-dihydroxyphenyl)heptane (157) [182] (Fig. 32), were isolated from C. comosa rhizomes [146]. Among these, five stereochemically different possible structures have been reported from the NMR data for the novel compound 132. The isolated compounds are classifiable into six germacrane-type (129–131, 138–140), sixteen guaiane-type (132–136, 141–151), one eudesmane-type (137), one bisabolane-type (152), and one carabrane-type (153) sesquiterpenoid, and four diarylheptanoids (154–157). Evaluation of sesquiterpenes 131, 133, 139, 140, 148, 149, 150 against cellular viability in a tumor cell and human U937 leukemic cell O R1 HO

O

OAc OAc R2

HO

OH

HO

154 (curcumin) R1 = R2 = OMe 155 (demethoxycurcumin) R1 = OMe, R2 = H 156 (bisdemethoxycurcumin) R1 = R2 = H

OH OH 157 ((3S,5S)-3,5-diacetoxy-1,7bis(3,4-dihydroxyphenyl)heptane)

Fig. 32 Structures of diarylheptanoids 154–157 isolated from C. comosa rhizomes

166

N. N. Win and H. Morita

line at three concentrations, 100, 10, and 1 μM, revealed that only (1S,4S,5S,10S)germacrone-1(10),4(5)-diepoxide (140) showed some growth inhibition (71.60% viability) at 100 μM. Antifungal activities of three curcuminoids (154–156) against Cladosporium cucumerinum at 20 μg/cm3 (each inhibition zone, 154 mm2 ) have also been reported [146].

2.9 Vitis repens Wight & Arn. Vitis repens (syn. Cissus repens Lam.) belongs to the family Vitaceae and is known in Myanmar as Tabin-daing-myanan (Fig. 33). This plant is distributed widely in the eastern hills of Shan State in Myanmar, and its rhizomes have been used in the local traditional medicine for the treatment of sores, carbuncles, oral and peptic ulcers, hepatitis and jaundice, tumors, hypertension, and other ailments in local traditional medicine [183]. In Chinese folk medicine, the roots and stems of C. repens are used for snakebites, rheumatic pains, and carbuncles, and the stems are applied for the treatment of nephritis, persistent coughs, and diarrhea. (E)-3-O-Methyl-resveratrol-2-C-β-glucoside, (Z)-3-Omethyl-resveratrol-2-C-β-glucoside, (E)-3-O-methyl-resveratrol-2-(2-p-coumaryl)C-β-glucoside (cissuside A), (E)-3-O-methyl-resveratrol-2-(3-p-coumaryl)-C-βglucoside (cissuside B), (E)-resveratrol, (E)-resveratrol-2-C-β-glucoside, and (Z)resveratrol-2-C-β-glucoside were isolated from Chinese Cissus repens [184]. Although various oligomers of resveratrol, as well as flavonoid, coumarin, and lignan

Fig. 33 The crude drug Vitis repens (rhizome) drug used in Myanmar traditional medicine

Bioactive Compounds from Medicinal Plants in Myanmar

167

constituents have been reported from other Vitis species [185], there are few reports concerning Vitis repens. The phytochemical and pharmacological activities of the constituents of V. repens from Myanmar were reported in 2004 [146], 2010 [186], and 2012 [187]. In 2004, eight compounds, including bergenin (158) [188], isolariciresinol (159) [189], 1-[(3methylbutyryl)phloroglucinol]-β-d-glucopyranoside (160) [190], 4-O-methylgallic acid (161) [191], protocatechuic acid (162) [192], gallic acid (105) [134], 2α,3β,23trihydroxyolean-12-en-28-oic-acid (120) [152], 4-O-galloylbergenin (163) (188), and pallidol (164) [193] were isolated from the ethyl acetate and n-butanol extracts of V. repens [146] (Fig. 34). In addition, according to the most potent antitrypanosomal activity of the methanol extract of the dried root bark of V. repens, with an IC 50 value of 8.6 ± 1.5 μg/cm3 and the greatest selectivity index of 24.4 [186], bioactivityguided fractionation of an ethanolic extract of V. repens led to the isolation of three compounds, comprising (E)-resveratrol (165) [194], 11-O-acetyl-bergenin (166) [195], and stigmast-4-en-3-one (167) [196] in 2009 [187] (Fig. 35). The biological OH OH

O

OH

O

O

OH OH

HO

OH O

HO

O

O

OH 159 (isolariciresinol)

158 (bergenin) H HO HO

O

H H

COOH

OH OH

O

H HO H

OH

R2

HO OR1

O

160 (1-[(3-methylbutyryl)phloroglucinol]β -D-glucopyranoside)

105 (gallic acid) R1 = H, R2 = OH 161 (4-O-methylgallic acid) R1 = Me, R2 = OH 162 (protocatechuic acid) R1 = R2 = H

O HO HO

OH

O HO O

HO

HO O

OH

O

O

HO HO

OH

OH

OH

OH 163 (4-O-galloyl-bergenin )

164 (pallidol)

Fig. 34 Structures of 158–164, 105, and 120 isolated from the ethyl acetate and n-butanol extracts of the V. repens rhizomes

168 Fig. 35 Structures of 165–167 isolated from ethanolic extract of V. repens rhizomes

N. N. Win and H. Morita OAc OH OH HO

OH

O

O

OH O

HO OH

O

165 ((E)-resveratrol)

166 (11-O-acetyl-bergenin)

O

167 (stigmast-4-en-3-one)

assay testing procedure used revealed for the first time that resveratrol, 11-O-acetylbergenin, and stigmast-4-en-3-one possess antitrypanosomal activities, with IC 50 values of 0.13, 0.17, and 0.15 μM, respectively [187].

2.10 Boesenbergia pandurata (Roxb.) Schltr. (Boesenbergia rotunda (L.) Mansf.) Boesenbergia pandurata (Roxb.) Schltr., a perennial herb of the Zingiberaceae family, is cultivated in some tropical countries, including Myanmar, Indonesia, Malaysia, and Thailand [160]. This plant has eight different botanical synonyms, namely, Boesenbergia cochinchinensis (Gagnep.) Loes., Boesenbergia pandurata (Roxb.) Schltr., Curcuma rotunda L., Gastrochilus panduratus (Roxb.) Ridl., Gastrochilus rotundus (L.) Alston, Kaempferia cochinchinensis Gagnep., Kaempferia ovata Roscoe, and Kaempferia pandurata Roxb. Currently, it is known as Boesenbergia rotunda (L.) Mansf. [197]. In Myanmar, this plant is referred colloquially as Seik-phoo (Fig. 36), and its rhizomes have been included in the TMF-47 (Khu-na-pah-hsay-wa-ga-lay), used for the treatment of asthma, diarrhea, indigestion, itching, and fever [198]. The rhizomes are also a popular folk medicine for diseases such as ulcers, dry mouth, stomach discomfort, leucorrhea, and dysentery in Indonesia, Malaysia, and Thailand [199]. The fresh rhizomes have a characteristic aroma and are used as flavoring agent in Thai cuisine [200]. They have also been employed as a self-medication by HIV/AIDS patients in Thailand [201]. Previous investigations on B. pandurata have reported anti-HIV [202], antibacterial [203], anti-inflammatory, analgesic, antipyretic [199, 204, 205], cytotoxic [206], antioxidant [207], and insecticidal activities [208].

Bioactive Compounds from Medicinal Plants in Myanmar

169

Fig. 36 Boesenbergia pandurata known as Seik-phoo and Seik-phoo-chin. The rhizomes are used in Myanmar traditional medicine

As a part of an anticancer drug discovery program for the treatment of pancreatic cancer, the constituents of cultivated B. pandurata rhizomes from Myanmar have been investigated since 2005 under an anti-austerity strategy, to search for candidates that eliminate preferentially tumor cell survival capabilities under low nutritional conditions [209, 210]. Accordingly, ten new secondary metabolite constituents including panduratins D–I (168–173), (1 R,2 S,6 R)-2-hydroxyisopanduratin A (174) (Fig. 37), geranyl-2,4-dihydroxy-6-phenethyl benzoate (175), 2 ,4 -dihydroxy-3 (1 -geranyl)-6 -methoxychalcone (176), and (2R)-8-geranylpinostrobin (177) [211, 212] (Fig. 38), together with 6-methoxypanduratin A (178) [213], panduratin A (179) [213], isopanduratin A2 (180) [208], hydroxypanduratin A (181) [200], panduratin C (182) [202], panduratins B1 (183) and B2 (184) [214], isopanduratin A1 (185) [208], nicolaioidesin B (186) [215] (Fig. 39), (2S)-6-geranylpinostrobin (187) [216], 6-geranylpinocembrin (188) [216, 217], pinostrobin (189) [218], pinocembrin (190) [219], alpinetin (191) [220], 7,4 -dihydroxy-5-methoxyflavanone (192) [221], (2S)-7,8-dihydro-5-hydroxy-2-methyl-2-(4 -methyl-3 -pentenyl)-8-phenyl2H,6H-benzo[1,2-b:5,4-b ]dipyran-6-one (193) [222], tectochrysin (194) [218], flavokawain C (195) [223], cardamonin (196) [220], boesenbergin A (197) [219], boesenbergin B (198) [224], and 5,6-dehydrokawain (199) [223] (Fig. 40), were isolated from an active chloroform extract of B. pandurata rhizomes collected in

170

N. N. Win and H. Morita

O

O

OH

O

O

OH

O

168 (panduratin D)

O

169 (panduratin E)

O

O

O

OH

O O

OH

O

170 (panduratin F) 171 (panduratin G)

R1 R2

OH

OH O

O

172 (panduratin H) R1 = Phenyl, R2 = COOMe 173 (panduratin I) R1 = COOMe, R2 = Phenyl

174 ((1'R,2'S,6'R)-2-hydroxyisopanduratin A)

Fig. 37 Structures of new cyclohexenyl chalcones 168–171 and 174 and cyclohexenyl derivatives 172 and 173 isolated from a chloroform extract of cultivated B. pandurata rhizomes in Myanmar

Myanmar. Among the isolated compounds, panduratins B1 (183) and B2 (184) were obtained as diastereomers. Anti-austerity assays revealed that a chloroform-soluble fraction of B. pandurata rhizomes exhibited 100% preferential cytotoxicity (PC 100 ) against the human PANC1 pancreatic cancer cell line in a nutrient-deprived medium (NDM) at 10 μg/cm3 [211]. Furthermore, in the same assay, the isolated compounds showed activity in the order: 179, 186 (2.5 μM) > 185, 188 (8 μM)> 174, 175, 176, 180, 181 (16 μM) > 178, 190, 193, 197, 198 (64 μM) > 168, 169, 171–173, 177, 183, 184, 187 (128 μM) > 170 (256 μM) > 182, 189, 191, 192, 194, 195, 196, 199 (> 256 μM). Among the flavonoids, those compounds possessing a geranyl moiety showed more potent activities (e.g, among the chalcones, 176  195, 196 and among the flavanones, 177,

Bioactive Compounds from Medicinal Plants in Myanmar

171

OH O OH

HO O

O

O

O 175 (geranyl-2,4-dihydroxy-6phenethylbenzoate)

176 (2',4'-dihydroxy-3'-(1''-geranyl)6'-methoxychalcone)

O O

O

O G=

OH 177 ((2R)-8-geranylpinostrobin)

Fig. 38 Structures of the new dihydroxyphenylbenzoate derivative 175, chalcone 176, and flavanone 177, isolated from a chloroform extract of cultivated B. pandurata rhizome in Myanmar

187, 188 > > 189, 191, 192). Based on the observed structure-activity relationships, a methoxy group at C-4 and hydroxy groups at C-2 and C-6 have been proposed as being important functionalities in cyclohexenyl chalcone derivatives for enhancing anti-austerity activity. Arctigenin, an anti-austerity-based anticancer agent [225], was used as a positive control in this study (PC 100 at 1 μM). It should be noted that paclitaxel (Taxol® ), a well-known anticancer agent, was inactive against PANC1 cells (PC 100 > 256 μM), which is consistent with a previous report (PC 100 > 3 mg/cm3 ) using this assay system [200]. Later on, in 2019, wild-type B. rotunda rhizomes occurring in the Bago Region of lower Myanmar were evaluated. These wild-type rhizomes are known locally as Seik-phoo-chin (Fig. 36), meaning a sour variety of Seik-phoo [226]. The chloroform-soluble extract of these rhizomes exhibited very weak antiproliferative activity against human lung (LK-2, A549), stomach (ECC4), breast (MCF7), cervical (HeLa), and prostate (DU145) cancer cell lines, with IC 50 values ranging from 48.7 to 73.7 μg/cm3 . Workup of the chloroform-soluble extract of these wildtype B. rotunda rhizomes revealed the occurrence of a new dinorcassane diterpenoid, seikphoochinal A (200), together with four other compounds, pinostrobin (189) [221, 227], 7,4 -dimethylkaempferol (201) [228], and galanals A (202) and B (203) [229] (Fig. 41). Interestingly, this study demonstrated that the chemical constituents of the wild-type (Seik-phoo-chin) and cultivated type (Seik-phoo) rhizomes to be quite different, despite the use of essentially the same extraction and isolation procedures as employed in previous reports on cultivated B. rotunda rhizomes [211, 212]. In addition, an antiproliferative assay conducted on the isolated compounds 189 and 201–203 (with the exception of compound 200), against six human cancer cell lines, LK-2, A549, ECC4, MCF7, HeLa, and DU145, indicated that galanals A (202) and B (203) were the most effective antiproliferative agents. For example, galanal A (202) exhibited low micromolar antiproliferative activities against the LK-2, HeLa,

172

N. N. Win and H. Morita O O

O

O O

O

O

OH 178 (6-methoxypanduratin A) R = R = H, R = R = Me 1

4

2

3

179 (panduratin A) R = R = R4 = H, R2 = Me 1

O

3

O

OH O

O

O

O 180 (isopanduratin A2) R1 = Me, R2 = R3 = R4 = H

181 (hydroxypanduratin A) R1 = R2 = R3 = R4 = H

O

O

OH

O OH

182 (panduratin C) R1 = R2 = H, R3 = Me, R4 =OH

O

HO

O

O O

183 and 184 (panduratins B1 and B2)

OH HOO

O O

O

OH

O HO

185 (isopanduratin A1) R = Me, R = H 1

2

OH

186 (nicolaioidesin B) R = H, R2 = Me 1

Fig. 39 Structures of the known cyclohexenyl chalcones 178–186 isolated from a chloroform extract of cultivated B. pandurata rhizomes in Myanmar

Bioactive Compounds from Medicinal Plants in Myanmar

173

R4 R 3O

O G=

R2 OR

1

O

187 ((2S)-6-geranylpinostrobin) R1 = R4 = H, R2 = G, R3 = Me 188 (6-geranylpinocembrin) R1 = R3 = R4 = H, R2 = G 189 (pinostrobin) R1 = R2 = R4 = H, R3 = Me 190 (pinocembrin) R1 = R2 = R3= R4 = H 191 (alpinetin) R1 = Me, R2 = R3 = R4 = H 192 (7,4'-dihydroxy-5-methoxyflavanone) R1 = Me, R2 = R3 = H, R4 = OH

R2 O

O

OH

O

O

193 ((2S)-7,8-dihydro-5-hydroxy-2methyl-2-(4''-methyl-3''pentenyl)-8-phenyl-2H,6Hbenzo[1,2-b:5,4-b']dipyran-6-one )

R1O

O

OH

O

OH

O

194 (tectochrysin)

O

195 (flavokawain) R1 = Me, R2 = OH 196 (cardamonin) R1 = R2 = H

O

O

O

OH

O

197 (boesenbergin A)

O

O

OH

O

O

O

198 (boesenbergin B)

199 (5,6-dehydrokawain )

Fig. 40 Structures of the known flavanones 187–193, the flavone 194, the chalcones 195–198, and the styryl-2-pyrone 199, isolated from a chloroform extract of cultivated B. pandurata rhizomes in Myanmar

and DU145 cell lines, with IC 50 values of 4.38, 5.10, and 3.57 μM, respectively. Galanal A (202) showed somewhat less potent activities against the A549, ECC4, and MCF7 cell lines (IC 50 values of 28.2, 32.0, and 38.6 μM, respectively). In contrast, galanal B (203) exhibited broad activity against all tested cell lines, with IC 50 values ranging from 5.26 to 8.02 μM. However, pinostrobin (189) did not inhibit the proliferation of any of the cancer cell lines utilized. The selective antiproliferative activity of 7,4 -dimethylkaempferol (201) against the LK-2 human lung cancer cell line and the ECC4 human stomach cancer cell line, with IC 50 values of 8.79 and 35.1 μM, respectively, was reported for the first time in this study [226]. From a chemotaxonomic point of view, the growth inhibitory effects for cancer cell lines of wild-type B. rotunda rhizomes may result from the presence of different chemical constituents that occur in those of the cultivated rhizomes. The chemical

174

N. N. Win and H. Morita

Fig. 41 Structures of 189 and 200–203 isolated from a chloroform extract of wild B. pandurata rhizomes in Myanmar

O

O

O

OH

O

189 (pinostrobin)

200 (seikphoochinal A) O O

O

O

R OH OH

O

201 (7,4'-dimethylkaempferol)

O

202 (galanal A) R = α-OH 203 (galanal B) R = β-OH

differences between these rhizomes are supported by the different tastes of the wild (sour taste) and cultivated (astringent taste) rhizomes, although this hypothesis may be somewhat subjective.

2.11 Kayea assamica King & Prain Kayea assamica, belonging to the family Clusiaceae, is a slow growing and tall evergreen tree known as Theraphi in Myanmar (Fig. 42). The bark of this species is light brownish-gray, often exfoliating in large square plates. The inside of the bark is fibrous and reddish with fine and close whitish veins, which soon turn brown [230]. The flower is bitter and fragrant and has been used to reduce extreme heat in the body, dizziness, dry skin, and fever [231]. The flower has been utilized as one of the constituents of TMF-15 (A-pu-naing-thee-chay-hsay), which has been especially useful for the treatment of fever [231]. In India, the fruits of the species are employed as a fish poison, and an aqueous extract of the stem bark is also a remedy for treating fevers [232]. The pollen is used for sores, fistulas, fever, and malaria [233]. Alkylated coumarins have been reported from the bark and fruit peels of this species [232]. An analysis of the phytochemicals obtained from the bark of K. assamica collected in Myanmar was reported in 2003 [233]. In this study, four new coumarin derivatives, theraphins A–D (204–207) [233], along with three known xanthones, 2hydroxyxanthone (208) [234], 1,7-dihydroxyxanthone (209) [235], and 5-hydroxy-1methoxyxanthone (210) [236], were isolated from the ethyl acetate extract of the bark (Fig. 43). Furthermore, in a cytotoxic assay of the four coumarins 204–207 against a panel of human cancer cell lines, consisting of LuI (human lung cancer), Col2 (human colon cancer), KB (epidermoid carcinoma of the nasopharynx), and LNCaP (hormone-dependent human prostate cancer) cells, compounds 204–206 exhibited

Bioactive Compounds from Medicinal Plants in Myanmar

175

Fig. 42 Kayea assamica, known as Theraphi. The flowers are used in Myanmar traditional medicine

HO OH

HO OH

O

HO

O

O

O

R

O

O

207 (theraphin D)

204 (theraphin A) R = 1-oxobutyl 205 (theraphin B) R = 2-methyl-1-oxobutyl 206 (theraphin C) R = 3-methyl-1-oxobutyl

O R4

R1 R2

O R3

208 (2-hydroxyxanthone) R1 = R3 = R4 = H, R2 = OH 209 (1,7-dihydroxyxanthone) R1 = R4 = OH, R2 = R3 = H 210 (5-hydroxy-1-methoxyxanthone) R1 = OMe, R2 = R4 = H, R3 = OH

Fig. 43 Structures of 204–210 isolated from an ethyl acetate extract of K. assamica bark from Myanmar

176

N. N. Win and H. Morita

HO OH 1

R

O

HO

O

R2

211 (kayeassamin A) R1 = G, R2 = 1-oxobutyl 212 (kayeassamin B) R1 = 1-oxobutyl, R2 = G 213 (kayeassamin C) R1 = 2-methyl-1-oxobutyl, R2 = G 214 (kayeassamin D) R1 = 3-methyl-1-oxobutyl, R2 = G 215 (kayeassamin E) R1 = 1-oxobutyl, R2 = isoprenyl 216 (kayeassamin F) R1 = 2-methyl-1-oxobutyl, R2 = isoprenyl 217 (kayeassamin G) R1 = 3-methyl-1-oxobutyl, R2 = isoprenyl

O

HO

O

O

O

O

HO

O

HO

O

O

219 (kayeassamin I)

218 (kayeassamin H)

1-oxobutyl =

O

3-methyl-1-oxobutyl =

isoprenyl =

O

2-methyl-1-oxobutyl =

O

G=

Fig. 44 Structures of the new coumarins 211–219 isolated from a chloroform extract of K. assamica flowers from Myanmar

cytotoxic effects against the Col2, KB, and LNCaP cell lines (IC 50 values in the range 3.5–13.1 μM). In contrast, compound 207 did not show any discernible activity against any of the tested cancer cell lines (IC 50 > ~50 μM). This preliminary biological assessment suggested that the presence of a 7-hydroxy group could be important for exhibition of cytotoxic activity by these coumarin derivatives. In addition, an antimalarial assay, using chloroquine-sensitive D6 and chloroquine-resistant W2 clones of Plasmodium falciparum, revealed compounds 204–207 to have moderate activities, with IC 50 values in the range of 9.7–11.1 μM against the D6 clone and IC 50 values in the range 5.1–10.4 μM against the W2 clone. The comparative values for chloroquine, which was used as a positive control, were IC 50 0.012 μM for the D6 clone and IC 50 0.13 μM for the W2 clone. Since the selectivity indices (SIs) (237)

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of the coumarin derivatives 204–206 were typically less than 1.0 and the values for 207 were 4.70 and 5.02 for the D6 and W2 clones, respectively, these coumarin derivatives have been proposed as being weak antimalarial agents. In turn, when the flowers of K. assamica in Myanmar were studied in 2008 [238, 239], the CHCl3 -soluble fraction showed anti-austerity activity, with a PC 100 value of 1 μg/cm3 against PANC-1 human pancreatic cancer cells in vitro. Subsequently, a series of chromatographic separations on this active extract resulted in the isolation of nine new coumarins, kayeassamins A–I (211–219) [238, 239] (Fig. 44), along with nine related compounds, including mammea A/AA cyclo D (220) [240, 241], mammea A/BC (221) [242], mammea B/AC (222) [243, 244], mammea A/AC

OH R O

O

O

220 (mammea A/AA cyclo D) R = 3-methyl-1-oxobutyl 224 (mammea A/AC cyclo D) R = 1-oxobutyl

O

OH

OH R1

O

HO

HO

O

O

O

R2

221 (mammea A/BC) R1 = isoprenyl, R2 = 1-oxobutyl 223 (mammea A/AC) R1 = 1-oxobutyl, R2 = isoprenyl

O

O

222 (mammea B/AC)

HO O

OH

O

O

O

HO

O

O HO

225 (mammea B/AC cyclo F)

226 (deacetylmammea E/BA cyclo D)

Fig. 45 Structures of the known coumarins 220–226 isolated from a chloroform extract of K. assamica flowers from Myanmar

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(223) [245, 246], mammea A/AC cyclo D (224) [245, 246], a mixture of theraphins B and C (205 and 206) [233], mammea B/AC cyclo F (225) [241, 243], and deacetylmammea E/BA cyclo D (226) [247] (Fig. 45). Furthermore, the anti-austerity assay used revealed that these isolated coumarins exhibited various potencies in a concentration-dependent manner. The most potent activities at levels comparable to that of the positive control, arctigenin (PC 100 of 1 μM) [225], were shown for kayeassamins A (211), B (212), D (214), E (215), and G (217), each with a PC 100 value of 1 μM, which were followed by 216, 222 (2 μM) > 213 (4 μM) > 221 (8 μM) > 223 (16 μM) > 220, 226 (32 μM) > 218, 219, 205, 206, 225 (64 μM) > 224 (>256 μM). Total preferential cell death was observed within 12 h of treatment with 211, 212, and 214 and within 24 h of treatment with 215 and 217 at 1μM. These results suggested that the presence of a geranyl moiety either at C-6 or C-8 in this type of coumarin is important for enhancing the resultant anti-austerity activity of this type of compound, which was in accordance with the observed preferential cytotoxicities in a concentration- as well as time-dependent manner for 211, 212, 214, 215 and 217. A further investigation has also revealed that kayeassamins A (211), B (212), D (214), E (215), and G (217) trigger apoptosis-like morphological changes in PANC-1 cells within 24 h at 4 μM, thereby causing the death of these cells. In addition, a wound closure assay [248], which may be utilized to evaluate the effects of compounds on the migratory ability of cancer cells, has also indicated not only the preferential cytotoxicities of 211 and 212 in NDM but also the inhibitory activities of 211 and 212 on PANC-1 cancer cell migration in a nutrient-rich medium. These additional observations suggested that geranylated coumarins are potent preferential cytotoxic agents as well as effective inhibitors against the migration of PANC-1 cells, and so they might be useful to inhibit the metastatic process evident in pancreatic cancer.

2.12 Cordia fragrantissima Kurz. Cordia fragrantissima is a timber tree that belongs to the family Boraginaceae. It occurs widely in Myanmar and is known locally as Sandawa or Taung-kalamet (Fig. 46) [249]. A lotion manufactured by grinding the wood with water has been applied to the hands and legs to relieve muscle pain. The bark is used to treat fever, diarrhea, and skin diseases and as an anthelmintic. In turn, the fruits are employed as a diuretic, expectorant, and anthelmintic and to treat lung and spleen diseases [250]. Although a number of phytochemicals such as quinones, flavonoids, and terpenoids have been reported since 1990 from other Cordia species [251–256], the chemical composition of C. fragrantissima was not reported until 2007. In a screening procedure for antileishmanial activity against Leishmania major by the heartwood constituents of C. fragrantissima, a methanol extract of the wood collected in Myanmar exhibited in vitro antileishmanial activity (MLC (minimum lethal concentration): 25 μg/cm3 ; MIC (minimum inhibitory concentration): 12.5 μg/cm3 ) [257]. Bioassay-guided fractionation of this active extract led to the isolation of three new compounds, cordiaquinols I–K (227–229) [258], and five

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179

Fig. 46 Cordia fragrantissima, known as Sandawa or Taung-kalamet. The bark or stem wood is used in traditional medicine in Myanmar

known compounds, cordiachromes A (230), B (231) [259, 260], and C (232) [261], cordiaquinol C (233) [262], and alliodorin (234) [263] (Fig. 47). Compounds 227– 234 were obtained as racemates. These racemates exhibited leishmanicidal activity OH

OH

OH HO

O OH

OH

O

227 (cordiaquinol I) O

OH

228 (cordiaquinol J) R1

OH

O 229 (cordiaquinol K) OH

O

6 5

O

11

230 (cordiachrome A) D 5,6 231 (cordiachrome B) D 5,11

R2 232 (cordiaqchrome C) R1 = R2 = =O 233 (cordiaquinol C) R1 = R2 = OH

OH 234 (alliodorin)

Fig. 47 Structures of 227–234 isolated from a methanol extract of C. fragrantissima grown in Myanmar

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against the promastigote form of L. major, L. guyanensis, and L. panamensis with IC 50 values ranging from 1.8 to 81.4 μg/cm3 . In contrast, cordiaquinol J (228) was found to be active against all three parasite species, with IC 50 values of 2.7, 3.0, and 1.8 μg/cm3 , respectively. Compounds 231, 232, 233, and 234 showed good activities against L. major with IC 50 values, in turn, of 4.1, 2.5, 4.5, and 7.0 μg/cm3 . None of these compounds was cytotoxic against COS-7 (African green monkey kidney cells, epithelial-like) and HuH-7 (human liver cancer cells, epithelial-like) cells [258].

2.13 Thanakha plants: Hesperethusa crenulata L. and Limonia acidissima L. Thanakha is a pale yellowish paste that is produced by grinding the bark, stem wood, or roots of Thanakha plants (Fig. 48) in water on a flat, smooth stone (Kyauk-pyin), which is a popular traditional cosmetic preparation used in Myanmar. Thanakha has been applied to the face, neck, hands, arms, and legs by members of the local

Fig. 48 Myanmar Thanakha plants. (a) Hesperethusa crenulata and (b) Limonia acidissima, both known locally as Thanakha plants. The bark, stem wood, or roots have use as traditional cosmetics in Myanmar

Bioactive Compounds from Medicinal Plants in Myanmar

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population for over 2000 years, as a natural cosmetic, sunscreen, skin preservation treatment, and insect repellent. Thanakha is believed to make the skin smooth, clear, and cool and increase the production of collagen and elastin to prevent wrinkles and skin aging, and ameliorate excessive facial oil, serious acne, pimples, blackheads, and whiteheads. Hesperethusa crenulata is the accepted name of a species in the genus Hesperethusa. The name is also a synonym of Limonia crenulata. In contrast, Limonia acidissima is the accepted name of a species in the genus Limonia, and its synonym O HN O

O R1 R3 R2

236 (N-{[p-(3,7-dimethyl-(6R),7-dihydroxy-(4R)-octadecanoyloxy-2octenyloxy)phenyl]ethyl} benzamide) O R1 =

R2 = R3 = OH 13

237 (N-{[p-(3,7-dimethyl-(6R),7-dihydroxy-(4R)-9’’’(E)-octadecenoyloxy-2octenyloxy)phenyl]ethyl} benzamide) O R1 =

R2 = R3 = OH 5

5

238 (N-{[p-(3,7-dimethyl-(6R),7-epoxy-(4R)-9’’’(E)-octadecenoyloxy-2octenyloxy)phenyl]ethyl}benzamide) O R2 and R3 is bonded by -O-

1

R = 5

5

AcO O

O

O

O

O

OAc

HO

HO

O O

239 (limodissimin A)

O

240 ((7'E)-(7R,8S)-4-hydroxy-3,5'-dimethoxy-4',7epoxy-8,3'-neolig-7'-en-9,9'-diyil diacetate)

Fig. 49 Structures of the three new benzamide derivatives 236–238, the dimeric coumarin 239, and the neolignan 240 isolated from an ethyl acetate extract of L. acidissima bark grown in Myanmar

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is Schinus limonia [264]. Both plants belong to the family Rutaceae and locally are known as Thanakha for the former species and Thi for the latter (Fig. 48). These plants have been used for the production of Thanakha paste. However, H. crenulata is recognized as authentic Thanakha, and L. acidissima is regarded as fake Thanakha, and their fragrances are different. These plants are both distributed in the Magway, Mandalay, and Sagaing Regions of Myanmar, as well as in India, Malaysia, and Sri Lanka [265]. Alkaloids and coumarins have been reported from the bark of H. crenulata [266, 267], while various constituents, including coumarins, steroids, triterpenoids, benzoquinones, and tyramine derivatives, have been isolated from different parts of L. acidissima [268–273].

HO HO O

HO

O

O

O

O

O

HO

O

O

O

242 ((Z)-suberenol)

241 ((E)-suberenol)

235 (marmesin)

HO

O

O

O

O

O

O

O

HO HO OH 244 ((2'R)-7-hydroxy-8-(2',3'-dihydroxy3'-methylbutyl)-2H-1-benzopyran-2-one)

243 (osthenol)

245 (columbianetin)

CHO O O

O

O

O

OR1

O OH

246 (seselin)

O O

247 (syringaldehyde) O O O

2

R O O

N

O

250 (4-methoxy-1-methyl-2(1H)-quinolinone)

248 ((+)-yangambin) R1 = R2 = Me 249 ((+)-syringaresinol) R1 = R2 = H

Fig. 50 Structures of the known coumarins 235 and 241–246, the phenol derivative 247, the furofuran lignans 248 and 249, and the known quinolinone 250, isolated from an ethyl acetate extract of L. acidissima bark grown in Myanmar

Bioactive Compounds from Medicinal Plants in Myanmar

183

A study in 2004 of H. crenulata of Myanmar origin [274] led to a report of the crystalline compound marmesin (235) as being present in a chloroform-methanol (1:1) extract of the bark, and its structure was confirmed as 2,3-dihydro-2(1-hydroxy1-methylethyl)-furanocoumarin [275, 276]. Marmesin (235) has three UV-absorbing chromophores, namely, an aromatic ring, a double bond between C-3—C-4, and a carbonyl at C-2, and hence gives a λmax at 335 nm. Since the powdered Thanakha bark used may have cosmetic use without causing any toxicity, Joo et al. proposed that marmesin (235) could be useful commercially as a natural UVA (320–380 nm)blocking product and as a starting material to synthesize even more effective UVAfiltering compounds through structural modification [274]. The chemical constituents of the bark of L. acidissima from Myanmar were reported in 2009 and 2010 [277–279]. Phytochemical investigation of the EtOAcsoluble fraction of the ethanol extract led to the isolation of three new benzamide derivatives, N-{[p-(3,7-dimethyl-(6R),7-dihydroxy-(4R)-octadecanoyloxy2-octenyloxy)phenyl]ethyl} benzamide (236), N-{[p-(3,7-dimethyl-(6R),7dihydroxy-(4R)-9 (E)-octadecenoyloxy-2-octenyloxy)phenyl]ethyl} benzamide O O O

OH

O O

O

HO

O

R2 R1 252 (limonin) R1 = H, R2 = =O, 253 (rutaevin) R1 = =O, R2 = β-OH

251 (13α,14β,17α-lanosta7,9,24-triene-3β,16α-diol)

HO

R3 R

R1 HO

OH

4

HO R2

254 (hederatriol) R1 = R3 = H, R2 = R4 = CH2OH 255 (bassic acid methyl ester) R1 = OH, R2 = CH2OH, R3 = H, R4 = COOMe 256 (3β-hydroxyolean-12-en-11-one) R1 = H, R2 = R4 = Me, R3 = =O

257 (13α,14β,17α-lanosta-7,24-diene3β,11β,16α-triol)

Fig. 51 Structures of the known terpenoids 251–257 isolated from an ethyl acetate extract of L. acidissima bark grown in Myanmar

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(237), and N-{[p-(3,7-dimethyl-(6R),7-epoxy-(4R)-9 (E)-octadecenoyloxy-2octenyloxy)phenyl]ethyl} benzamide (238) [277]. Also obtained were a new dimeric coumarin, limodissimin A (239) [278] as well as a new neolignan stereoisomer, (7 E)-(7R,8S)-4-hydroxy-3,5 -dimethoxy-4 ,7-epoxy-8,3 -neolig-7 -en-9,9 -diyil diacetate (240) [279] (Fig. 49), together with 23 other compounds (241–262). The other 23 compounds were identified as the previously reported coumarins, marmesin (235) [274–276], (E)-suberenol (241) [280], (Z)-suberenol (242) [281], osthenol (243) [282], (2 R)-7-hydroxy-8-(2 ,3 -dihydroxy-3 -methylbutyl)-2H1-benzopyran-2-one (244) [283], columbianetin (245) [284], and seselin (246) [285]; a phenol derivative, syringaldehyde (247) [286]; the furofuran lignans (+)-yangambin (248) [287] and (+)-syringaresinol (249) [137], a quinolinone, 4-methoxy-1-methyl-2(1H)-quinolinone (250) [288] (Fig. 50); the terpenoids, 13α,14β,17α-lanosta-7,9,24-triene-3β,16α-diol (251) [289], limonin (252) [290], rutaevin (253) [291], hederatriol (254) [292], bassic acid methyl ester (255) [293], 3β-hydroxyolean-12-en-11-one (256) [294], and the fatty acids (9E)-octadecenoic acid (258) [295], cascarillic acid (259) [296], (+)-α-dimorphecolic acid (260) [297], (8R)-hydroxylinoleic acid (261) (299), and (6Z,9Z,12Z)-pentadecatrienoic acid (262) [299] (Figs. 51 and 52). O HO 258 (E)-9-octadecenoic acid

OH O 259 (cascarillic acid) OH OH 260 ((+)-α-dimorphecolic acid)

O

O OH OH 261 ((8R)-hydroxylinoleic acid) O OH 262 ((6Z,9Z,12Z)-pentadecatrienoic acid)

Fig. 52 Structures of the known fatty acids 258–262 from an ethyl acetate extract of L. acidissima bark grown in Myanmar

Bioactive Compounds from Medicinal Plants in Myanmar

185

Among these isolates, compounds 240, 254, and 260–262 exhibited the most potent NO production inhibitory activities (IC 50 (μM): 8.08, 7.28, 8.82, 6.73, 7.01) against LPS-activated BV-2 cells, followed by 250, 251, 257 (IC 50 /μM: 9.03, 10.7, 12.8) and 243–246 [IC50 /μM: 22.3, 21.6, 33.5, 23.1], respectively [277]. The inhibitory effects of compounds 240, 250, 254, and 260–262 were greater than that of the positive control, l-NMMA (IC 50 9.50 μM). However, cytotoxicity was found from compounds 236–238 at 10 μM. Since activated microglial cells produce excessive inflammatory substances, such as NO, cytokines, and prostaglandins, NO derived from inducible nitric oxide synthase in LPS-activated microglia is an important mediator of inflammation and neuronal cell death [300]. Thus, Limonia acidissima and its active constituents may be good lead compounds for the development of novel neuroprotective agents.

2.14 Soymida febrifuga (Roxb.) A. Juss. Soymida febrifuga belongs to the family Meliaceae. It is distributed widely in tropical areas of Asia, inclusive of India, Sri Lanka, and Myanmar. In Myanmar, the plant is referred to colloquially as Dan-da-kuu-ni (Fig. 53), and the bark powder is a major ingredient of TMF-16 (A-pu-that-hsay) and TMF-20 (Mahar-kalyarni-hsay), and a common ingredient of TMF-18 (Ganar-doneba-hsay), TMF-25 (Thone-sae-kunitpar-ziwila-winga-hsay), and TMF-43 (Akin-sae-par-hsay). TMF-16 and TMF-20 have their use in eliminating extreme heat in the body and are effective in the treatment of fever, influenza, diarrhea, indigestion, and headache [301]. This species is also

Fig. 53 Soymida febrifuga, known as Dan-da-kuu-ni. The bark is used in Myanmar traditional medicine

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well known as Indian redwood and is a popular traditional medicine in India [302]. In the Ayurveda system of traditional medicine, the bark is used for blood purification and is also recommended for ulcers, leprosy, dysentery, and as an anti-inflammatory agent [303]. It is considered to be as effective as cinchona bark for the treatment of malaria [304]). Limonoids and their insect antifeedant activities have been reported from the leaves of Soymida febrifuga [305, 306]. Prior investigations of the various parts of S. febrifuga have led to the isolation of lupeol, sitosterol, methyl angolensate, deoxyandirobin, and two tetranortriterpenoids with a modified furan ring from the bark [307–310]. The bark of S. febrifuga from Myanmar has been investigated, with the results published in 2009 [311]. A 70% ethanol extract of S. febrifuga bark was found to inhibit 50% of human pancreatic PANC-1 cancer cells preferentially, under nutrition-deprived conditions at a concentration of 10 μg/cm3 . Twenty-seven compounds including four new ones, (3R)-6,4 -dihydroxy-8-methoxyhomoisoflavan (263), 7-hydroxy-6-methoxy-3-(4 -hydroxybenzyl)coumarin (264), and 6hydroxy-7-methoxy-3-(4 -hydroxybenzyl)coumarin (265) (Fig. 54), and (2R)7,4 -dihydroxy-5-methoxy-8-methylflavan (266) (Fig. 55), together with 23 known compounds, inclusive of 6,4 -dihydroxy-7-methoxyhomoisoflavan (267) [312], 7,4 dihydroxyhomoisoflavan (268) [313], 5,7-dihydroxy-4 -methoxyhomoisoflavanone (269) [314], 5,7,4 -trihydroxy-6-methoxyhomoisoflavanone (270) [315] 5,7,4 trihydroxyhomoisoflavanone (271) [314], 7,4 -dihydroxyhomoisoflavanone (272) [313] (Fig. 54), 7,4 -dihydroxy-8-methylflavan (273) [315], 7,4 -dihydroxyflavan (274) [316], 4 -hydroxy-7-methoxyflavan (275) [317], 5,7-dihydroxyflavanone (276) [318] (Fig. 55), 2 ,4 -dihydroxychalcone (277) [318], 2,2 ,4 -trihydroxychalcone (278) [319], 2,4 -dihydroxy-2 -methoxychalcone (279) [320], 2,4 -dihydroxy-4methoxychalcone (280) [317], 4 -hydroxy-2,4-dimethoxydihydrochalcone (281) [321], 4-hydroxy-2,6,4 -trimethoxydihydrochalcone (282) [322], 2,4 -dihydroxy4-methoxydihydrochalcone (283) [317], 2,4,4 -trihydroxydihydrochalcone (284) [323], 4,4 -dihydroxy-2,6-dimethoxydihydrochalcone (285) [324] (Fig. 56), 4 hydroxy-3,5-dimethoxystilbene (286) [325], 3,4 -dihydroxy-5-methoxystilbene (287) [326], p-hydroxybenzoic acid (288) [327], and guaiacylglycerol (289) [328] (Fig. 57), were isolated from a CH2 Cl2 -soluble fraction of the active EtOH extract. The isolated compounds were mainly flavonoid derivatives that were classified as three homoisoflavans (263, 267, 268), two 3-benzylcoumarins (264, 265), four homoisoflavanones (269–272), four flavans (266, 273–275), one flavanone (276), four chalcones (277–280), five dihydrochalcones (281–285), and two stilbenes (286, 287). Also, one phenol derivative (288), and one glycerol derivative (289) were obtained. Among these compounds, 2 ,4 -dihydroxychalcone (277) displayed the most potent preferential cytotoxicity (PC 50 19.0 μM) against PANC-1 cancer cells under nutrition-deprived conditions. In contrast, no cytotoxicity was observed with 2,2 ,4 trihydroxychalcone (278) and 2,4 -dihydroxy-2 -methoxychalcone (279). These observations indicated that specific chalcone substituents are required for activity. Similar results have been reported also for 4 -hydroxy-3,5-dimethoxystilbene (286),

Bioactive Compounds from Medicinal Plants in Myanmar

187

R1 R2

O

OH

R3 263 ((3R)-6,4'-dihydroxy-8-methoxyhomoisoflavan) R1 = OMe, R2 = H, R3 = OH 267 (6,4'-dihydroxy-7-methoxyhomoisoflavan) R1 = H, R2 = OMe, R3 = OH 268 (7,4'-dihydroxyhomoisoflavan) R1 = R3 = H, R2 = OH HO

OR3

O

R1 R2

O

269 (5,7-dihydroxy-4'-methoxyhomoisoflavanone) R1 = H, R2 = OH, R3 = OMe 270 (5,7,4'-trihydroxy-6-methoxyhomoisoflavanone) R1 = OMe, R2 = OH, R3 = H 271 (5,7,4'-trihydroxyhomoisoflavanone) R1 = R3 = H, R2 = OH 272 (7,4'-dihydroxyhomoisoflavanone) R1 = R2 = R3 = H

R2 O

O

O

OH

R1 O 264 (7-hydroxy-6-methoxy-3-(4'-hydroxybenzyl)coumarin) R1 = Me, R2 = H 265 (6-hydroxy-7-methoxy-3-(4'-hydroxybenzyl)coumarin) R1 = H, R2 = Me

Fig. 54 Structures of the homoisoflavans 263, 267, and 268, the homoisoflavanones 269–272, and the 3-benzylcoumarins 264 and 265, isolated from a dichloromethane extract of S. febrifuga bark grown in Myanmar OH

R1

OH

O

HO

O

O

R2

266 ((2R)-7,4'-dihydroxy-5-methoxy-8-methylflavan) R1 = Me, R2 = OMe 273 (7,4'-dihydroxy-8-methylflavan) R1 = Me, R2 = H 274 (7,4'-dihydroxyflavan) R1 = R2 = H

275 (4'-hydroxy-7-methoxyflavan)

O

HO

OH

O

276 (5,7-dihydroxyflavanone)

Fig. 55 Structures of the flavans 266 and 273–275 and the flavanone 276, isolated from a dichloromethane extract of S. febrifuga bark grown in Myanmar

188

N. N. Win and H. Morita R3 HO R2 R1

O

277 (2',4'-dihydroxychalcone) R1 = OH, R2 = R3 = H 278 (2,2',4'-trihydroxychalcone) R1 = R2 = OH, R3 = H 279 (2,4'-dihydroxy-2'-methoxychalcone) R1 = OMe, R2 = OH, R3 = H 280 (2,4'-dihydroxy-4-methoxychalcone) R1 = H, R2 = OH, R3 = OMe

R4

OR3

R1O OR2 O

281 (4'-hydroxy-2,4-dimethoxydihydrochalcone) R1 = R4 = H, R2 = R3 = Me 282 (4-hydroxy-2,6,4'-trimethoxydihydrochalcone) R1 = R2 = Me, R3 = H, R4 = OMe 283 (2,4'-dihydroxy-4-methoxydihydrochalcone) R1 = R2 = R4 = H, R3 = Me 284 (2,4,4'-trihydroxydihydrochalcone) R1 = R2 = R3 = R4 = H 285 (4,4'-dihydroxy-2,6-dimethoxydihydrochalcone) R1 = R3 = H, R2 = Me, R4 = OMe

Fig. 56 Structures of the chalcones 277–280 and the dihydrochalcones 281–285, isolated from a dichloromethane extract of S. febrifuga bark grown in Myanmar

O

O

O OH OR

OH

HO OH

HO

286 (4'-hydroxy-3,5-dimethoxystilbene) R = Me 287 (3,4'-dihydroxy-5-methoxystilbene) R = H

OH

HO

288 (p-hydroxybenzoic acid)

289 (guaiacylglycerol)

Fig. 57 Structures of the stilbenes 286 and 287, the phenol derivative 288, and the glycerol derivative 289, isolated from a dichloromethane extract of S. febrifuga bark grown in Myanmar

which showed a more potent activity (PC 50 44.4 μM) than that determined for 3,4 dihydroxy-5-methoxystilbene (287) (PC 50 71.4 μM). In addition, an in vitro cytotoxicity assay was performed for compounds 263–289 (excluding 265) against a panel of five cancer cell lines, viz., two murine cancer cell lines (colon 26-L5 carcinoma (colon 26-L5), B16-BL6 melanoma (B16-BL6)) and three human cancer cell lines (lung A549 adenocarcinoma (A549), cervix HeLa adenocarcinoma (HeLa), and HT1080 fibrosarcoma (HT-1080)). This revealed that 4 -hydroxy-3,5-dimethoxystilbene

Bioactive Compounds from Medicinal Plants in Myanmar

189

(286) showed the most potent activity against the HeLa (IC 50 9.43 μM), colon 26-L5 (IC 50 2.96 μM) and B16-BL6 (IC 50 15.9 μM) cell lines. In particular, the observed IC 50 value of 286 against colon 26-L5 cells was comparable to that of the positive control doxorubicin (IC 50 3.12 μM).

2.15 Vitex negundo L. Vitex negundo, belonging to the family Verbenaceae, is a popular herbal plant that occurs commonly throughout Myanmar, where it is known as Kyaung-ban-gyi (Fig. 58). The fruits, leaves, and roots have been recognized as useful parts in human healthcare. The dried fruit powder is taken orally with honey, at a dose of 5–10 g for diarrheal diseases. For menstrual disorders, urinary disorders, and indigestion, the dried powder is ingested orally with roasted common salt and warm water at the same dose as above. The leaves have been used as a curry substitute. The root powder is made into a slurry with alcohol and is externally applied for muscle cramps [329]. V. negundo is also a well-known medicinal herb in the Indian medicinal system, Fig. 58 Vitex negundo, known as Kyaung-ban-gyi. The fruits, leaves, and roots have their use in traditional medicine in Myanmar

190

N. N. Win and H. Morita OR OH O

O

O

O OH

O

290 (chrysoplenetin) R = Me 291 (chrysosplenol D) R = H

Fig. 59 Structures of the flavones 290 and 291 isolated from a methanol extract of V. negundo fruits grown in Myanmar

where it is referred to as Five-Leaved Chaste Tree or Monk’s pepper. V. negundo extracts have been used in the Unani system of medicine for anti-inflammatory, expectorant, tranquilizer, antispasmodic, anticonvulsant, rejuvenative, antiarthritic, anthelminthic, antifungal, and antipyretic purposes. In the Unani system, the seeds are recommended for controlling premature ejaculation and enhancing male libido. The Ayurvedic and Unani Pharmacopoeia of India has documented the use of the leaves, seeds, and roots to treat excessive vaginal discharge, edema, skin diseases, pruritus, helminthiasis, rheumatism, and puerperal fever [330]. Potent preferential cytotoxic activity against PANC-1 human pancreatic cancer cells using the anti-austerity strategy was observed for a crude MeOH extract of the V. negundo fruits obtained in Myanmar [331]. Subsequent bioactivity-guided phytochemical investigation led to the isolation of two known flavones, chrysoplenetin (290) [332] and chrysosplenol D (291) [333] (Fig. 59). This was the first report of the occurrence of chrysoplenetin (290) in V. negundo collected in Myanmar as well as in the genus Vitex. Compounds 290 and 291 have been identified as the active constituents with PC 50 (50% preferential cytotoxicity) values of 3.4 μg/cm3 and 4.6 μg/cm3 , respectively, against PANC-1 cells, and apoptosis-like morphological changes by both compounds were observed in PANC-1 cells. Furthermore, of a panel of 39 human cancer cell lines (JFCR-39) available at the Japanese Foundation for Cancer Research, 25 of these, inclusive of lung, breast, CNS, colon, melanoma, ovarian, prostate, and stomach cancer cell lines, revealed high susceptibilities to chrysoplenetin (290) in the submicromolar range. In particular, chrysoplenetin (290) exhibited potent sensitivities against NCIH522 lung, OVCAR-3 ovarian, and PC-3 prostate cancer cells, with 50% growth inhibition (GI 50 ) values of 0.12, 0.18, and 0.17 μM, respectively. Comparative analysis using the JFCR-39 panel also suggested that the molecular mode of action of chrysoplenetin (290) is different when compared with those of several clinically used anticancer drugs [331].

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Fig. 60 Diospyros burmanica, known as Hpun-mang, Mai-mak-ho-ling, Mai-mate-si or Te in Myanmar. There is no information on the use of this species in traditional medicine O

O

O

O

O

R O

O

O OH

O

O

OH

OH

292 (burmanin A)

O

293 (burmanin B) R = H 294 (burmanin C) R = OMe

OH R1

R4

O

R2

O

R3 R1

R2 R4

R3

295 (4,8-dimethoxy-3-methyl-1-naphthol) R1 = R4 = H, R2 = Me, R3 = OMe 296 (2,8-dimethoxy-3-methyl-1-naphthol) R1 = OMe, R2 = Me, R3 = R4 = H 297 (8-methoxy-3-methyl-1-naphthol) R1 = R3 = R4 = H, R2 = Me 298 (5,8-dimethoxy-3-methyl-1-naphthol) R1 = R3 = H, R2 = Me, R4 = OMe 299 (4-hydroxy-5-methoxy-2-naphthalenecarboxaldehyde) R1 = R3 = R4 = H, R2 = CHO

300 (3-hydroxy-5-methoxy-2-methyl-1,4-naphthalenedione) R1 = OH, R2 = OMe, R3 = R4 = H 301 (5,8-dimethoxy-2-methyl-1,4-naphthalenedione) R1 = R3 = H, R2 = R4 = OMe 302 (7-hydroxy-8-methoxy-2-methyl-1,4-naphthalenedione) R1 = R2 = H, R3 = OH, R4 = OMe 303 (5-methoxy-2-methyl-1,4-naphthalenedione) R1 = R3 = R4 = H, R2 = OMe

Fig. 61 Structures of the bisnaphthoquinone analogs 292–294, the naphthols 295–299, and the naphthoquinones 300–303, isolated from a chloroform-soluble fraction of D. burmanica wood grown in Myanmar

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2.16 Diospyros burmanica Kurz. Diospyros burmanica is a tree that belongs to the family Ebenaceae. It is widely distributed in the Bago and Mandalay Regions of Myanmar and is locally known as Hpun-mang, Mai-mak-ho-ling, Mai-mate-si or Te (Fig. 60) [334]. It is well known as the plant of wild persimmon fruits, and there is no information on its traditional medicinal use. The chemical composition of D. burmanica were first reported in 2012 [335]. The methanol extract of the wood of D. burmanica in Myanmar exhibited potent activity against Leishmania major (MLC 6.25 μg/cm3 ; MIC 1.25 μg/cm3 ) in a previous screening [257]. Furthermore, three unprecedented bisnaphthoquinone analogs, burmanins A–C (292–294), five naphthols, 4,8-dimethoxy-3methyl-1-naphthol (295) [337], 2,8-dimethoxy-3-methyl-1-naphthol (296) [338], 8-methoxy-3-methyl-1-naphthol (297) [338], 5,8-dimethoxy-3-methyl-1-naphthol (298) [339], and 4-hydroxy-5-methoxy-2-naphthalenecarboxaldehyde (299) [340], and four naphthoquinones, 3-hydroxy-5-methoxy-2-methyl-1,4-naphthalenedione (300) [336, 341], 5,8-dimethoxy-2-methyl-1,4-naphthalenedione (301) [342], 7hydroxy-8-methoxy-2-methyl-1,4-naphthalenedione (302) [343], and 5-methoxy-2methyl-1,4-naphthalenedione (303) [336, 344] were isolated from the CHCl3 -soluble fraction of the active methanol extract by a combination of various chromatographic methods [335] (Fig. 61). A leishmanicidal assay of 292–303 against the promastigote form of L. major suggested that burmanin A (292) was the most active compound (IC 50 0.053 ± 2.7 × 10−3 μM), followed by the dimeric analogs 293 (IC 50 0.18 ± 5.4 × 10−3 μM) and 297 (IC 50 0.15 ± 11 × 10−3 μM) and the monomeric naphthoquinone 303 (IC 50 3.3 ± 0.19 μM). In contrast, much weaker activity (> 38 μM) was observed from the other monomeric naphthoquinone (302) and the naphthols (295–299). A cytotoxic assay of 292–303 against the murine macrophage-like cell line RAW264.7 also indicated that burmanins A–C (292–294) showed weak toxicity (IC 50 values 24 ± 1.7, 31 ± 0.45, and 22 ± 0.64 μM, respectively), which is required for potential antileishmanial drug candidates in addition to high leishmanicidal activity, since the Leishmania protozoan parasite dwells and multiplies within mammalian macrophages [336]. The selectivity indexes of burmanins A–C (292–294) of 453, 172, and 147, respectively, were relatively higher than those of reported natural monomeric naphthoquinones determined from amastigotes of GFP-transfected L. major versus BMM (SI = 1.2–7.0) [345]. These results have suggested that burmanins A–C (292–294) could be potential lead compounds for drugs in the treatment of parasitic diseases caused by L. major.

2.17 Cinnamomum inunctum Kurz. Cinnamomum inunctum belongs to the family Lauraceae. It is referred to as Kara-way (Fig. 62) and distributed in the states of Kachin, Mon, and Shan of Myanmar [346],

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Fig. 62 Cinnamomum inunctum known as Kara-way. The leaves, fruits, and bark have use in Myanmar traditional medicine

where its dried fruits and leaves are utilized quite widely in traditional medicine. The dried fruits are a common ingredient of TMF-8 (Thway-dular-hsay), TMF29 (Thabba-zay-ya-hsay-gyi), and TMF-39 (Mahar-than-wutta-bayar-hsay). These formulations are prescribed for the treatment of fever, influenza, food poisoning, indigestion, diarrhea, dysentery, malaria, rheumatism, menstrual disorders, and blood purification after childbirth [347]. Although the components of the essential oil are known as the main constituents of many Cinnamomum species [348, 349], there has been no report regarding the phytochemical profile of C. inunctum until quite recently. The constituents of C. inunctum fruits in Myanmar were reported for the first time in 2015 [350]. In this work, four previously unknown monoterpene lactones, 5-(2,3dihydroxy-3-methylbutyl)-4-hydroxy-4-methyldihydrofuran-2(3H)-one (304), 5(2,3-dihydroxy-3-methylbutyl)-4-methylfuran-2(5H)-one (305), 8-hydroxy-4,7,7trimethyl-1,6-dioxaspiro[4.4]non-3-en-2-one (306), and 8-hydroxy-4,7,7-trimethyl1,6-dioxaspiro[4.4]non-3-en-2-one (307) [350], together with 3-hydroxy-4,4dimethyl-4-butyrolactone (308) [351], were isolated from a methanol-soluble extract of C. inunctum (Fig. 63). Among them, compounds 306 and 307, with unique spirolactone moieties, were determined as being racemates. However, the biological activities of both the crude plant extracts and the above-mentioned compounds have yet to be reported.

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HO O

O

HO HO

HO

O

O 305 (5-(2,3-dihydroxy-3-methylbutyl)-4methylfuran-2(5H)-one)

304 (5-(2,3-dihydroxy-3-methylbutyl)-4hydroxy-4-methyldihydrofuran-2(3H)-one) HO

HO O

O

O

O

O

O 307 (8-hydroxy-4,7,7-trimethyl-1,6dioxaspiro[4.4]non-3-en-2-one)

306 (8-hydroxy-4,7,7-trimethyl-1,6dioxaspiro[4.4]non-3-en-2-one)

O HO

O

308 (3-hydroxy-4,4-dimethyl-4-butyrolactone)

Fig. 63 Structures of the monoterpene lactones 304–308 isolated from a methanol extract of C. inunctum fruits grown in Myanmar

2.18 Kaempferia pulchra Ridl. Kaempferia pulchra is a perennial herb of the Zingiberaceae family. It is cultivated in some tropical countries, including Myanmar, Indonesia, Malaysia, and Thailand. In Myanmar, it is known colloquially as Shan-pan-oot (Fig. 64), and has been used extensively for the treatment of coughs, diabetes mellitus, urinary infections, in blood stimulation, as a carminative, a heat quencher, deodorant, and diuretic. It Fig. 64 Kaempferia pulchra known as Shan-pan-oot. The rhizomes have use in traditional medicine in Myanmar

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is also one of the major ingredients in TMF-21 (Hsee-hsay-phyu) and a common ingredient of TMF-20 (Mahar-kalyarni-hsay), TMF-23 (Hsay-pale-kalat), TMF-32 (Nan-dwin-shar-put-hsay-gyi), TMF-39 (Mahar-than-wutta-bayar-hsay), and TMF43 (Akin-sae-bah-hsay). These formulations have been employed for the treatment of various fevers, influenza, and indigestion, as a diuretic, and for diabetes, diarrhea, and purification of the blood [352]. The rhizomes reportedly possess anti-inflammatory and antitumor activities [353, 354] and have been used in a restricted manner for self-medication by cancer, diabetes, and AIDS patients. Sandaracopimaradiene diterpenoids and ethyl 4-methoxy-(E)-cinnamate have been detected in extracts from this plant grown in Thailand [353–355]. A phytochemical and pharmacological investigation of K. pulchra rhizomes grown in Myanmar was reported in 2015 [356]. In this study, a CHCl3 soluble fraction of the rhizomes and/or the isolated compounds were tested for their various biological activities, such as antiproliferative activity [356, 357], viral protein R (Vpr) inhibitory activity [360], and NO production and nuclear factor-kappa B (NF-κB) expression inhibitory activities [360, 361]. Thirty-one compounds, including 23 new isopimarane diterpenoids, kaempulchraols A-W (309–331) [356–359] (Fig. 65), together with 9α-hydroxyisopimara8(14),15-dien-7-one (332) [362], 7β,9α-dihydroxypimara-8(14),15-diene (333) [363], (1S,5S,9S,10S,11R,13R)-1,11-dihydroxypimara-8(14),15-diene (334) [364], sandaracopimaradien-1α,2α-diol (335) [353], (1R,2S,5S,9S,10S,11R,13R)-1,2,11trihydroxypimara-8(14),15-diene (336) [364], 7α-hydroxyisopimara-8(14),15-diene (337) [365], (2R)-ent-2-hydroxyisopimara-8(14),15-diene (338) [366], and ethyl 4methoxy-(E)-cinnamate (339) [353] (Fig. 66), were isolated from the active CHCl3 soluble fraction of K. pulchra rhizomes. The isolated compounds were mainly classifiable into isopimara-8(9),15-dienes (309–312, 315, 322, 323, 327, 329) and isopimara-8(14),15-dienes (313, 314, 316, 317–321, 324–326, 328, 330–338). In an evaluation of the antiproliferative activity using the CCK-8 assay [356, 357, 367], the CHCl3 extract of K. pulchra showed weak inhibition of the proliferation of all tested human cancer cell lines, including A549 (human lung cancer), HeLa (human cervical cancer), PANC-1 and PSN-1 (human pancreatic cancer), and MDA-MB-231 (human breast cancer), with IC 50 values of 30.2, 26.7, 37.9, 20.4, 39.0 μg/cm3 (unpublished data), respectively. Furthermore, the assay also revealed various antiproliferative activity potency levels, with IC 50 values ranging from 12.3 to 99.3 μM for compounds 309–339. In this assay, kaempulchraol B (310) was the most active compound against A549 and PANC-1 cells, with IC 50 values of 32.9 and 25.4 μM, respectively, whereas the proliferation of PSN-1, HeLa, and MDA-MB231 cells was selectively inhibited by kaempulchraols F (314), M (321), and T (328), with IC 50 values of 12.3, 28.4, and 48.2 μM, respectively. However, antiproliferative activities were lacking against all the tested cell lines by compounds 322, 323, 329–333, at the concentration levels used [356, 357, 367]. As in the case of the antiproliferative activity testing, anti-Vpr activity evaluation using HeLa cells harboring, the Vpr expression plasmid (TREx-HeLa-Vpr cells) also was conducted on not only the CHCl3 -soluble fraction of K. pulchra rhizomes but also the isolated compounds 309–339 [359, 367]. Vpr is a small basic protein

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N. N. Win and H. Morita R1 R4 R3 R2

309 (kaempulchraol A) R1 = R3 = H, R2 = β-OH, R4 = α-OMe 310 (kaempulchraol B) R1 = R3 = H, R2 = β-OH, R4 = β-OMe 311 (kaempulchraol C) R1 = R3 = H, R2 = β-OH, R4 = α-OH 312 (kaempulchraol D) R1 = R3 = H, R2 = β-OH, R4 = β-OH 315 (kaempulchraol G) R1 = R3 = H, R2 = β-OH, R4 = =O 322 (kaempulchraol N) R1 = R4 = α-OH, R2 = β-OH, R3 = H 323 (kaempulchraol O) R1 = α-OH, R2 = β-OH, R3 = H, R4 = β-OMe 327 (kaempulchraol S) R1 = R2 = H, R3 = =O, R4 = α-OH 329 (kaempulchraol U) R1 = R2 = R3 =H, R4 = α-OH

R1 R2 R6 R3

R5 R4

313 (kaempulchraol E) R1 = α-OH, R2 = R3 = R5 = R6 = H, R4 = β-OH 314 (kaempulchraol F) R1 = R3 = α-OH, R2 = R4 = R5 = R6 = H 316 (kaempulchraol H) R1 = R3 = α-OH, R2 = R5 = R6 = H, R4 = β-OH 317 (kaempulchraol I) R1 = α-OH, R2 = R3 = R4 = R5 = R6 =H 318 (kaempulchraol J) R1 = α-OH, R2 = R3 = R4 = R6 =H, R5 = =O 319 (kaempulchraol K) R1 = R2 = R3 = R5 = H, R4 = β-OAc, R6 = OH 320 (kaempulchraol L) R1 = R2 = R3 = R5 = H, R4 = β-OH, R6 = OMe 321 (kaempulchraol M) R1 = R2 = R6 = α-OH, R3 = R4 = R5 = H 324 (kaempulchraol P) R1 = R2 = R3 = R5 = R6 = H, R4 = β-OH 325 (kaempulchraol Q) R1 = α-OAc, R2 = R3 = R5 = R6 = H, R4 = β-OH 326 (kaempulchraol R) R1 = R2 = R3 = R4 = H, R5 = α-OAc, R6 = OH 328 (kaempulchraol T) R1 = R2 = R3 = R6 = H, R4 = β-OH, R5= α-OAc 330 (kaempulchraol V) R1 = R2 = R3 = H, R4 = β-OH, R5= β-OAc, R6 = OH 331 (kaempulchraol W) R1 = R2 = R3 = H, R4 = R5 = β-OH, R6 = OH

Fig. 65 Structures of the new isopimara-8(9),15-dienes 309–312, 315, 322, 323, 327, 329 and the isopimara-8(14),15-dienes 313, 314, 316, 317–321, 324–326, 328, 330, 331, isolated from a chloroform-soluble fraction of K. pulchra rhizomes grown in Myanmar

(14 kDa) that is conserved in HIV-1, HIV-2, and simian immunodeficiency virus (SIV) [368, 369], and has been suggested as a promising drug target for comprehensive HIV/AIDS therapy [370, 371]. Fumagillin, damnacanthal, vipirinin, and quercetin had been reported as Vpr inhibitors earlier [372–375]. The CHCl3 -soluble fraction of K. pulchra inhibited Vpr in TREx-HeLa-Vpr cells at an effective dose of 25 μg/cm3 . Furthermore, the assay results also revealed that kaempuchraols B (310), D (312), G (315), Q (325), T (328), U (329), and W (331) exhibited anti-Vpr activities at concentrations ranging from 1.56 to 6.25 μM. The inhibitory potencies of compounds 310, 312, 315, 325, 328, 329, and 331 were comparable to that of

Bioactive Compounds from Medicinal Plants in Myanmar R1

197

R5

R2

HO R4 R3

332 (9α-hydroxyisopimara-8(14),15-dien-7-one)

338 ((2R)-ent-2-hydroxyisopimara-8(14),15-diene)

R1 = R2 = R5 = H, R3 = =O, R4 = OH 333 (7β,9α -dihydroxypimara-8(14),15-diene) R1 = R2 = R5 = H, R3 = β-OH, R4 = OH

O

334 ((1S,5S,9S,10S,11R,13R)-1,11-dihydroxypimara-8(14),15-diene)

OEt

R1 = R5 = α-OH, R2 = R3 = R4 = H 335 (sandaracopimaradien-1α,2α -diol) R1 = R2 = α-OH, R3 = R4 = R5 = H

O

339 (ethyl 4-methoxy-(E)-cinnamate)

336 ((1R,2S,5S,9S,10S,11R,13R)-1,2,11-trihydroxypimara-8(14),15-diene) R1 = R2 = R5 = α-OH, R3 = R4 = H 337 (7α-hydroxyisopimara-8(14),15-diene) R1 = R2 = R4 = R5 = H, R3 = α-OH

Fig. 66 Structures of the known isopimara-8(14), 15-dienes 332–338 and the cinnamate derivative 339, isolated from a chloroform-soluble fraction of K. pulchra rhizomes grown in Myanmar

the positive control, damnacanthal, at 5 μM. Structure-activity relationship studies using this assay have suggested that the presence of a β-OH group at C-6 and C-14 and the absence of either a methoxy or carbonyl group in an isopimara-8(9),15-diene skeleton are important structural requirements for enhancing the activities of this type of compound, whereas an α-OAc or β-OAc group at either C-1 or C-7 enhanced the overall activity of the 6β-hydroxy-isopimara-8(14),15-dienes. Anti-inflammatory activity has been investigated for nine isopimara-8,(9),15diene diterpenoids, 309–312, 315, 322, 323, 327, and 329, and 21 isopimara-8,14(15)diene diterpenoids, 313, 314, 316–321, 324–326, 328, and 330–338 [360, 361]. The CHCl3 -soluble extract of K. pulchra exhibited NO inhibitory activity, with an IC 50 value of 31.43 μg/cm3 , without showing any cytotoxicity in LPS-induced RAW264.7 cells. Furthermore, three isopimara-8(9),15-diene diterpenoids, kaempulchraols B–D (310–312), and two isopimara-8(14),15-diene diterpenoids, kaempulchraols P (324) and Q (325), were effective as NO inhibitory agents. In these assays, kaempulchraols B–D (310–312) and kaempulchraols P (324) and Q (325) exhibited NO inhibitory activities with IC 50 values of 47.69, 44.97, 38.17, 39.88, and 36.05 μM, respectively, again without showing any cytotoxicity to LPS-induced RAW264.7 cells. In addition, investigations of the mechanisms of action of 310–312 and kaempulchraols P (324) and Q (325) demonstrated their individual ability to inhibit the NF-κB-mediated transactivation of a luciferase reporter gene, interleukin-6 (IL-6) production, and cyclooxygenase-2 (COX-2) expression, with an effective dose of 25 μM [360, 361]. These findings provide new insights into the potential anti-inflammatory activities of isopimara-8(9), 15-diene diterpenoids and isopimara-8(14),(15)-diene diterpenoids.

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2.19 Picrasma javanica Blume Picrasma javanica is a medium-sized tree belonging to the family Simaroubaceae. Plants of this family contain structurally diverse and biologically active quassinoids and indole alkaloids, with significant cytotoxic and antimalarial activities. P. javanica is distributed widely in the tropical regions of Asia, including Myanmar, Indonesia, and India. In Myanmar, it is known as Nann-paw-kyawt or Taung-kamaka [376] (Fig. 67), and has been used quite extensively for self-medication by malaria, cancer, and HIV/AIDS patients. Decoctions of its bark are used in folk medicine as a febrifuge and a substitute for quinine. Numerous quassinoids and alkaloids have been reported as chemical constituents of P. javanica [377–394]. Two reports of the constituents of P. javanica bark appeared in 2015 and 2016 [395, 396]. In a preliminary screening procedure, a CHCl3 -soluble fraction of P. javanica bark exhibited generally potent antiproliferative activities against a small panel of five different cancer cell lines, including A549 (human lung), HeLa (human cervical), PANC-1 and PSN-1 (human pancreatic), and MDA-MB-231 (human breast), with Fig. 67 Picrasma javanica, known as Nann-paw-kyawt or Taung-kamaka. The bark is used in Myanmar folk medicine

Bioactive Compounds from Medicinal Plants in Myanmar

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O O

O R3

O

R

O

1

O

R2

O

O R4

R1

O

R3 O

R1

O

O

R2 340 (picrajavanicin A) R1 = R2 = R4 = H, R3 = CHO 344 (picrajavanicin E) R1 = OMe, R2 = R4 = H, R3 = Me 345 (picrajavanicin F) R1 = OMe, R2 = H, R3 = Me, R4 = OH 347 (picrajavanicin H) R1 = R2 = R4 = H, R3 = CH2OH 349 (picrajavanicin J) R1 = OMe, R2 = R4 = H, R3 = CH2OH 350 (picrajavanicin K) R1 = R4 = H, R2 = =O, R3 = Me 351 (picrajavanicin L) R1 = R4 = H, R2 = β-OH, R3 = Me 352 (picrajavanicin M) R1 = R4 = H, R2 = α-OH, R3 = Me

341 (picrajavanicin B) R1 = R3 = H, R2 = Me 342 (picrajavanicin C) R1 = R2 = Me, R3 = H 343 (picrajavanicin D) R1 = H, R2 = Me, R3 = OH 348 (picrajavanicin I) R1 = R3 = H, R2 = CH2OH

O

O O

O

OH

O O 346 (picrajavanicin G)

O

O

O

O O

R

358 ((16R)-methoxyjavanicin B) R = α-OMe 359 ((16S)-methoxyjavanicin B) R = β-OMe

Fig. 68 Structures of the new quassinoids 340–352, 358 and 359 isolated from P. javanica grown in Myanmar

IC 50 values of 8.3, 12.9, 1.6, 22.1, and 14.2 μg/cm3 , respectively. Thirteen new quassinoids, picrajavanicins A–M (340–352) [395, 396] (Fig. 68), together with five related quassinoids, javanicins B (353) [386], F (354) [388], and I (355) [385], picrasin A (356), and 2 -isopicrasin A (357) [397] (Fig. 69), were isolated from this active CHCl3 extract. In contrast to the CHCl3 extract, most of the isolated quassinoids lacked antiproliferative activity against A549, HeLa, PSN-1, and MDAMB-231 cells. However, compounds 347–352, 356, and 357 were selectively active against PANC-1 cells with IC 50 values ranging from 3.25 to 17.4 μM. In turn, compounds 347, 356, and 357 also inhibited selectively the proliferation of HeLa cells, with IC 50 values of 9.50, 10.2, and 3.98 μM, respectively. Hence, the strong antiproliferative activity of the extract may be due to the effects of β-carboline alkaloids reported as major constituents of P. javanica. Interestingly, anti-Vpr activity (treated dose, 5 μg/cm3 ) was also observed by the chloroform-soluble extract of P. javanica, which is in good agreement for the traditional uses of this plant for self-medication by HIV/AIDS patients. Furthermore, the evaluation of the Vpr inhibitory effects of picrajavanicins A–K (340–350) and M (352) and javanicins B (353), F (354), and I (355) using TREx-HeLa-Vpr cells suggested that the compounds had the ability to inhibit Vpr activity in the potency order: 355 > 353 > 342, 343 > 340, 341, 350, 354 > 347 > 344, 345, 346, 348, 352 > 349, at a treated dose of 2.5 μM [398]. The potency of the most active inhibitor

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O O

O

O

O

O

O O

353 (javanicin F)

353 (javanicin B) O O

O

O

OH

O

O

HO O

O

O

O O

O

O

O

353 (javanicin B)

O

353 (picrasin A)

O HO O

O

O

O O

O 353 (2’-isopicrasin A)

O 360 (lanosta-7,24-dien-3-one)

O HO

N O

O

N O

361 (scopoletin)

362 (canthin-6-one)

Fig. 69 Structures of the known quassinoids 353–357, the sterol 360, the coumarin 361, and the indole alkaloid 362 isolated from P. javanica grown in Myanmar

among the picrasane quassinoids tested, javanicin I (355), was comparable to that of the positive control, damnacanthal. Structure-activity relationship conclusions on the 2,12,14-triene-1,11,16-trione-2,12-dimethoxy-18-norpicrasane quassinoids, picrajavanicin G (346) and javanicin I (355), suggested that the presence of a methyl group at C-13 is crucial for exhibiting significant anti-Vpr activity in this type of compound. It should be noted that javanicin I (355) was obtained as the least polar compound among the isolates from the CHCl3 extract. In addition, the presence of a methyl group at C-13, hydroxy or carbonyl groups at C-4, a hydroxy group at C-14 and the absence of a methoxy group at C-3 in other picrasane-type quassinoids, all have been proposed as being important structural requirements for potent Vpr inhibitors [398]. These observations suggest that the polarity of the C-13 side chain is related to the overall Vpr inhibitory effects.

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The constituents of P. javanica wood were described in 2019 [399]. Two unprecedented quassinoids, (16R)-methoxyjavanicin B (358) and (16S)-methoxyjavanicin B (359) [399] (Fig. 68), and the known compounds, lanosta-7,24-dien-3-one (360) [400], scopoletin (361) [401], and canthin-6-one (362) [402] (Fig. 69), were isolated from a CHCl3 -soluble portion of the MeOH extract of this plant part. P. javanica wood contains also picrajavanicins B (341) and C (342) [396] and javanicins B (353) [386] and F (354) [388], the same chemical constituents as found in P. javanica bark (Fig. 69). Lanosta-7,24-dien-3-one (360) showed some activity against the human lung (A549), breast (MCF7), and cervical (HeLa) cancer cell lines and the normal fibroblast cell line tested, with IC 50 values ranging from 48.6 to 65.9 μM. In contrast, (16R)-methoxyjavanicin B (358) and (16S)-methoxyjavanicin B (359) exhibited higher activity (each with an MIC value of 1.6 μM), whereas 353 and 341 had inhibitory activity (MIC values of 6.3 and 3.1 μM) against the Gram-positive bacterium, Bacillus subtilis.

2.20 Curcuma amada Roxb. Curcuma amada is an important member of the genus Curcuma genus (family Zingiberaceae) and is known colloquially as mango ginger, due to the raw mango-like aroma of the rhizomes. This herbaceous perennial plant is referred to in Myanmar as Thayetkin [160] (Fig. 70), and has a long history of traditional uses ranging from folk medicine to culinary preparations. The fresh rhizomes of C. amada are consumed as a dipping vegetable. The rhizomes are also utilized in ethnomedicine for the treatment of skin diseases, stomach ailments, coughs, inflammation, and rheumatism. Phytochemical studies of C. amada have revealed the presence of labdane diterpenoids, β-sitosterol, and curcumin [403]. The constituents of C. amada occur also in other members of the genera Curcuma [166, 404–407], in addition to Alpinia [408–417], Hedychium [418–422], Zingiber [423], and Aframomum [424, 425]. Various biological activities, such as antiobesity, antiamnesic, neuroprotective Fig. 70 Curcuma amada known as Thayetkin. The rhizomes are used in traditional medicine in Myanmar

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363 (12β-hydroxy-15-norlabda8(17),13(14)-dien-16-oic acid)

O HO

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364 ((E)-15-ethoxy-15-methoxylabda8(17),12-dien-16-al)

O O O

O O OH

365 ((E)-15α-ethoxy-14α-hydroxylabda8(17),12-dien-16-olide)

366 (15-ethoxy-12β-hydroxylabda8(17),13(14)-dien-16,15-olide)

Fig. 71 Structures of the new labdane diterpenoids 363–366 isolated from a methanol extract of C. amada rhizomes grown in Myanmar

[426]), antioxidant, cytotoxic, antimicrobial, platelet aggregation inhibitory [427], anti-inflammatory, antiallergic, biopesticide, and hyperglyceridemia effects, have been described previously [428]. Phytochemical and pharmacological studies of C. amada native to Myanmar were reported in 2017 [429]. Disernible antiproliferative activities were displayed by a MeOH extract of C. amada rhizomes, with IC50 values ranging from 31.7 to 49.2 μg/cm3 , against the A549 (human lung cancer), HeLa (human cervix cancer), MCF7 (human breast cancer), and PANC-1 and PSN-1 (human pancreatic cancer) cell lines. Furthermore, phytochemical workup of this bioactive methanol extract resulted in the isolation of 17 compounds, including four new labdane diterpenoids (12β-hydroxy-15-norlabda-8(17),13(14)-dien-16-oic acid (363), (E)-15-ethoxy15-methoxylabda-8(17),12-dien-16-al (364), (E)-15α-ethoxy-14α-hydroxylabda8(17),12-dien-16-olide (365), and 15-ethoxy-12β-hydroxylabda-8(17),13(14)-dien16,15-olide (366) [429]) (Fig. 71), together with the 13 known labdane diterpenoids ((E)-15,16-bisnorlabda-8(17),11-dien-13-one (367) [409], (E)-14,15,16trinorlabda-8(17),11-dien-13-al (368) [418], (E)-14,15,16-trinorlabda-8(17),11dien-13-oic acid (369) [423], 16-oxolabda-8(17),12-dien-15,11-olide (370) [419], (E)-15,15-diethoxylabda-8(17),12-dien-16-al (371) [425], (E)-labda-8(17),12-dien15,16-dial (372) [430], (E)-14-hydroxy-15-norlabda-8(17),12-dien-16-al (373) [423], zarumin A (374) [412], methyl (12E)-16-oxolabda-8(17),12-dien-15-oate (375) [417], (E)-labda-8(17),12-dien-15-ol-16-al (376) [421] coronarin D ethyl ether

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(377) [431], coronarin D methyl ether (378) [418], and (E)-labda-8(17),12,14-trien15,16-olide (379) [423]) (Fig. 72). Among these isolated labdane diterpenoids, compounds 364–366 and 376–379 were somewhat active as cytotoxic agents against the cell lines in which they were evaluated, displaying IC50 values ranging from 19.7 to 96.1 μM. Interestingly, (E)-14-hydroxy-15-norlabda-8(17),12-dien-16-al (373) inhibited selectively the growth of HeLa, PANC-1, and PSN-1 cells, with IC 50 values of 5.88, 1.00, and 3.98 μM, respectively, at potency levels comparable to those of the positive control, 5-fluorouracil (IC 50 values of 8.34, 11.7, and 7.11 μM, respectively). Compound 373 has been isolated from the rhizomes of Zingiber ottensii [423] and the roots of Aframomum melegueta [432]. In contrast, compounds 363 and 368 were not active at all at the highest treatment dose used (100 μM) against all of the cancer cell lines employed in the investigation. Structure-activity relationship conclusions among the labdane diterpenoids 371–376 against PANC-1 and PSN-1 O O

O

R

H O

367 ((E)-15,16-bisnorlabda-8(17),11dien-13-one) R = Me 368 ((E)-14,15,16-trinorlabda-8(17),11dien-13-al) R = H 369 ((E)-14,15,16-trinorlabda-8(17),11dien-13-oic acid) R = OH

370 (16-oxolabda-8(17),12-dien15,11-olide) RO O O

O R

H

377 (coronarin D ethyl ether) R = Et 378 (coronarin D methyl ether) R = Me O

371 ((E)-15,15-diethoxylabda-8(17),12dien-16-al ) R = CH(OEt)2 372 ((E)-labda-8(17),12-dien-15,16-dial) R = CHO 373 ((E)-14-hydroxy-15norlabda-8(17),12-dien-16-al ) R = OH 374 (zarumin A) R = COOH 375 (methyl (12E)-16-oxolabda-8(17),12dien-15-oate ) R = COOMe 376 ((E)-labda-8(17),12-dien-15-ol-16-al) R = CH2OH

O

379 ((E)-labda-8(17),12,14trien-15,16-olide)

Fig. 72 Structures of the known labdane diterpenoids 367–379 isolated from a methanol extract of C. amada rhizomes grown in Myanmar

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cells suggested that a hydroxymethyl functionality at C-13 is important to the inhibition of the proliferation of human pancreatic cancer cell lines, within this compound class [429].

2.21 Vitex trifolia L. Vitex trifolia belongs to the family Verbenaceae. It is an aromatic coastal deciduous shrub. In Myanmar, it is a popular herb where it is known locally as Kyaung-banlay (Fig. 73). The leaves, fruits, and roots have been used for the alleviation of indigestion, dyspepsia, diarrhea, dysentery, menstrual disorders, urinary disorders, maintaining men’s health, and muscle cramps [435]. The leaves and fruits have been reported to have positive effects in treating amnesia [434], cancer [435–438], inflammation [439, 440], and parasitic infections [441], in addition to having wound healing [442] and antibacterial effects [443, 445]. Several types of compounds, such as flavonoids [438], labdane-type diterpenoids [446], lignans [447], terpenoids [448], and essential oil components [449], have been reported from the different parts Fig. 73 Vitex trifolia known as Kyaung-ban-lay. The leaves, fruits, and roots have use in traditional medicine in Myanmar

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of V. trifolia. This species is used as an anti-inflammatory agent in Chinese folk medicine to treat headaches, colds, migraine, eye pain, rheumatic pains, and cancer. The fruits have utilization in the treatment of amenorrhea, anxiety, the common cold, headaches, watery eyes, mastitis, and as an emmenagogue (stimulate blood flow in the pelvic area and uterus). V. trifolia is known as sambhalu in Unani medicine and is employed for treating decreasing libido. The inner bark has use against dysentery, as an expectorant, to lower blood pressure and to treat the common cold, and for prosopalgia, chronic tracheitis, sinusitis, periodontitis, rheumatism, and pulmonary tuberculosis. The roots are used as an anthelmintic, insecticidal, and diuretic agent [450]. In 2017, a report was published on the leaves of V. trifolia from Myanmar [451]. In this study, an ethyl acetate extract was shown to be an inducer of adipogenesis similar to rosiglitazone (ROS), an antidiabetic thiazolidinedione drug, in 3T3-L1 preadipocytes. Purification of this active ethyl acetate extract by various chromatographic methods also afforded three compounds, vitexilactone (380) [452], vitexicarpin (381) [453], and oleanolic acid (382) [454] (Fig. 74). Among the isolated compounds, the ROS-like action of 380 was of some interest. The effects of vitexilactone on 3T3-L1 cells during adipogenesis were compared with those of ROS. Furthermore, 380 has also been shown to increase lipid accumulation and the expression of adiponectin and glucose transporter type 4 (GLUT4) in the cell membrane, and to decrease both the size of adipocytes and the phosphorylation of IRS-1, extracellular signal-regulated kinase 1/2 (ERK1/2) and JNK in 3T3-L1 cells, as observed for ROS. However, in contrast to ROS, the induction of proteins involved in the lipogenesis of vitexilactone (380) was partial. The ROS-like effects of 380 in 3T3-L1 O OH

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383 ((–)-hinokinin) R = =O 384 ((–)-O-methylcubebin) R = OMe 385 ((–)-cubebin) R = OH

Fig. 74 Structures of compounds 380–385 isolated from an ethyl acetate extract of V. trifolia leaves grown in Myanmar

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cells were suppressed by the addition of bisphenol A diglycidyl ether, a peroxisome proliferator-activated receptor (PPAR) antagonist, suggesting that the action of 380 on adipocytes is mediated by PPAR. Since PPAR is a ligand-activated transcription factor that plays a crucial role in the regulation of glucose homeostasis and lipid metabolism and is considered to be one of the important targets for the treatment of metabolic disorders such as type 2 diabetes, and vitexilactone has been proposed as a novel insulin-sensitizer candidate [451]. Further work on V. trifolia by the same group in order to elucidate its antidiabetic components has been published [455]. Three lignans, (−)-hinokinin (383) [456], (−)-O-methylcubebin (384) [457], and (−)-cubebin (385) [460], were isolated from the ethyl acetate extract of the leaves of V. trifolia, using various chromatographic methods (Fig. 74). The structure of 384 was also determined by X-ray crystal structure analysis. The occurrence of these three lignans from V. trifolia and single-crystal X-ray analysis of (–)-O-methylcubebin were reported for the first time [455]. As in the case of 380, an upregulation of intracellular lipid accumulation assay revealed that 383–385 possess the effect of increasing intracellular lipid accumulation in a concentration-dependent manner, in the potency order of 384 > 383 > 385. However, interestingly, while compound 384 had the most potent effect, further study has revealed that 384 is able to not only reduce the diameter of differentiated adipocytes significantly but also increase the expression of adiponectin, which is known to be an important molecule to reduce insulin resistance. Notably, the activity of (−)-O-methylcubebin (384) was inhibited by antagonists of PPARγ and improved by inhibitors of the classical mitogen-activated protein kinase (MAPK) pathway and p38 MAPK pathway, while this lignan did not show any migration of GLUT4 from the cytoplasm to the cell membrane observed for ROS, even though it showed similar effects to those of ROS. Thus, compound 384 has been proposed as a PPARγ agonist with the effect of promoting adipogenesis via the inhibition of ERK1/2 and p38 MAPK phosphorylation. Hence, (−)-O-methylcubebin (384) could be a lead compound that does not show weight gain, which is a side effect of ROS [455].

2.22 Mansonia gagei Drumm. Mansonia gagei is a tree belonging to the family Sterculiaceae. It is found in both Thailand and Myanmar. Locally in Myanmar it is known as Kala-met (Fig. 75), and is one of the components of TMF-15 (Thee-chay-hsay). This formulation has been prescribed for insomnia, menopause, swelling, reducing body heat, and improving the appetite and condition of the skin [459]. Furthermore, a liquid paste obtained by grinding the heartwood of M. gagei with water is utilized as a popular lotion that is effective in cooling the skin and relieving muscle pain. Several mansonone onaphthoquinones and coumarins have been reported from M. gagei, and the individual constituents obtained from this species display antiestrogenic, larvicidal, antioxidant, antifungal, and cytotoxic activities [460–466].

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Fig. 75 Mansonia gagei known as Kala-met. The bark and stem wood have use in traditional medicine in Myanmar

A phytochemical study of M. gagei bark from Myanmar was reported in 2018 [467]. The hexane and methanol/water (95:5, v/v) extracts exhibited relatively strong melanogenesis inhibition effects (IC 50 26 μg/cm3 ) against α-melanocyte-stimulating hormone (α-MSH)-induced B16 cells. Four compounds, mansonone E (386) [461], mansorin I (387) [464], populene F (388) [468], and mansonone G (389) [465], were isolated from the hexane extract of the bark (Fig. 76). In this study, mansorin B (390) [461] was also obtained from the bark methanol/water extract (Fig. 76). The occurrence of populene F (388) in M. gagei was reported for the first time. All compounds were shown to be more active compounds than arbutin (IC 50 > 100 μM) in the bioassay system used, with the order of potency being mansonone G (389) (IC 50 7 μM) > mansorin B (390) (IC 50 28 μM) > mansorin I (387) (IC 50 48 μM) > mansonone E (386) (IC 50 83 μM) > populene F (388) (IC 50 83 μM). Also, the order of cytotoxic potency of these five compounds against B16 melanoma cells

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388 (populene F)

389 (mansonone G)

386 (mansonone E)

O

O

HO

O

O

HO O

387 (mansorin I)

390 (mansorin B)

Fig. 76 Structures of compounds 386-390 isolated from M. gagei bark

decreased in an almost parallel manner to that observed for their melanogenesis inhibitory activities (mansonone G (389) (IC 50 26 μM) > mansorin I (387) (IC 50 56 μM) ≥ mansorin B (390) (IC 50 58 μM) > populene F (388) (IC 50 68 μM) > mansonone E (386) (IC 50 > 100 μM)). However, in this case, mansonone E (386) lacked cytotoxic activity. A hydroxy group at C-6 has been suggested as being a key feature structurally, to determine both the cytotoxic and melanogenic effects of this series of isolated compounds. Among the isolated compounds, the mechanism of the melanogenesis inhibitory effects was investigated for the least active and non-cytotoxic compound mansonone E (386), which dose-dependently inhibited the expression levels of tyrosinase (TYR), tyrosinase-related protein 1 (TRP-1), TRP-2, cAMP response element binding protein (CREB), and microphthalmia-associated transcription factor (MITF). These levels were increased by α-MSH stimulation. Furthermore, this compound rescued the phosphorylation of Akt and p38 MAPK, which were up- or downregulated by α-MSH stimulation, in a dose-dependent manner. In contrast, co-treatment of 386 with the phosphoinositide 3-kinase (PI3K)/Akt inhibitor wortmannin enhanced melanogenesis inhibition by mansonone E. Thus, mansonone E has been suggested as a suppressor of α-MSH-induced melanogenesis in B16 cells with the effect of inhibiting both phosphorylation in the PI3K/Akt pathway and the expression of melanogenesis-related proteins [467].

2.23 Premna integrifolia L. (syn.: P. serratifolia L.) Premna integrifolia (syn: Premna serratifolia) is a tree belonging to the family Verbenaceae [471]. It is distributed in the southern part of Myanmar, India, Malaysia, and Sri Lanka. In India, most of its plant parts have use in traditional medicine

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Fig. 77 The stem wood of Premna integrifolia known as Taung-tangyi. The stem wood is used to produce Thanakha cream used as a traditional cosmetic in Myanmar

to treat various infectious diseases. P. integrifolia reportedly possesses anticoagulant [470], antiarthritic [471], antihyperglycemic [472], and antimicrobial properties [473]. The various plants parts of P. integrifolia elaborate a variety of alkaloid, isoprenoid, flavonoid, and lignan constituents [474, 475]. In Myanmar, this plant is referred to as Taung-tangyi (Fig. 77), and is used to produce Thanakha cream. Women, especially those in the Tanintharyi Region, use the wood of P. integrifolia to produce a natural cosmetic, sunscreen, skin preservation treatment, and insect repellent. However, phytochemical and pharmacological studies relating to this type of cosmetic use have not been investigated yet in a scientific, evidence-based manner. In 2018, the results of an investigation of a sample of P. integrifolia from Myanmar were reported. A CHCl3 -soluble extract of the P. integrifolia stem wood was found to exhibited quite potent antimelanogenesis activity, with an IC 50 value of 28.3 μg/cm3 [476]. Ten compounds, including two new tetrahydrofuran lignans, taungtangyiols A (391) and B (392) (Fig. 78), and eight known furofuran lignans, sesamin (393) [477], paulownin (394) [478], 4α-hydroxysesamin (395), 4α,8α-dihydroxysesamin (396) [479], asarinin (397) [477], xanthoxyol (398) [480], 1α-hydroxy-6-epipinoresinol (399), and 1α-hydroxypinoresinol (400) [481] (Fig. 79), were isolated by utilizing a combination of various chromatographic techniques [476]. The structure of 391 was confirmed by acetylation of 391, which provided 391a and 391b (Fig. 79). In addition, eleven compounds, including four new lignans, premnans A (401) and B (402), taungtangyiol C (403) [482], 7,9-dihydroxydolichanthin B (404) [483], in adition to

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R1 R4

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391 (taungtangyiol A) R1 = H, R2 = R3 = R4 = OH 392 (taungtangyiol B) R1 = R3 = R4 = OH, R2 = H 391a (6,7-diacetoxytaungtangyiol A) R1 = H, R2 = OH, R3 = R4 = OAc 391b (5,6,7-triacetoxytaungtangyiol A) R1 = H, R2 = R3 = R4 = OAc O O

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405 (premnan C)

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406 ((3R,4S)-4-(1,3-benzodioxol-5-ylcarbonyl) -3-[(R)-1-(1,3-benzodioxol-5-yl) -1-hydroxy methyl]tetrahydro-2-furanone)

Fig. 78 Structures of the new lignans 391, 392, and 401–406, isolated from a chloroform-soluble extract of P. integrifolia stem wood grown in Myanmar, and 391a and 391b obtained by acetylation of 391

premnan C (405) [482], which was assumed to be an artifact and one natural product lignan, (3R,4S)-4-(1,3-benzodioxol-5-ylcarbonyl)-3-[(R)-1-(1,3-benzodioxol-5-yl)1-hydroxymethyl]tetrahydro-2-furanone (406) [484] (Fig. 78), together with five previously known plant constituents ((−)-aptosimon (407), (−)-diasesamin-di-γ lactone (408) [485], and (+)-epi-sesaminone (409) [486] (Fig. 79), oleanonic acid

Bioactive Compounds from Medicinal Plants in Myanmar

211 O

R1

O

O

R2 O

R3

O

O

393 (sesamin) R1 = R2 = R3 = H 394 (paulownin) R1 = R3 = H, R2 = OH 395 (4α-hydroxysesamin) R1 = OH, R2 = R3 = H 396 (4α,8α-dihydroxysesamin) R1 = R3 = OH, R2 = H O O

O O

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397 (asarinin)

398 (xanthoxyol) OH

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O O O

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407 ((–)-aptosimon)

399 (1α-hydroxy-6-epipinoresinol) 2-Hα 400 (1α-hydroxypinoresinol) 2-H

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O O

O

O

O

O O

O

O

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408 ((–)-diasesamin-di-γ-lactone)

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HO

409 ((+)-epi-sesaminone)

Fig. 79 Structures of the known lignans 393–400 and 407–409, isolated from a chloroform extract of P. integrifolia stem wood grown in Myanmar

(410) [487] and (2α,3α)-dihydroxyolean-12-en-28-oic acid (411) [488]) (Fig. 80), were reported [482, 483]. Melanogenesis inhibitory effects against melanocyte-stimulating hormone (αMSH) and a potent cyclic nucleotide phosphodiesterase inhibitor, 3-isobutyl-1methylxanthine (IBMX)-induced B16F10 mouse melanoma cells, have been shown for the furofuran lignans 393–398, which exhibited potencies with IC 50 values less than the minimum treated dose (10 μM). Most of the compounds were more potent than the positive control arbutin (IC 50 364.4 μM). In contrast, tetrahydrofuran lignans 391 and 392 inhibited the production of melanin, with IC 50 values of 50.7 and 40.9

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O

O OH

HO

OH

HO

O 410 (oleanonic acid)

411 ((2α,3α )-olean-12-en-28-oic acid)

Fig. 80 Structures of the known triterpenes 410 and 411 isolated from a chloroform extract of P. integrifolia stem wood grown in Myanmar

μM, respectively, without showing any cytotoxicity. However, the acetate derivatives of 391 (391a and 391b) did not inhibit the deposition of melanin, even at the maximum dose used (100 μM). Accordingly, these findings support the traditional use of P. integrifolia wood extract as a cosmetic agent and melanogenesis inhibitor [476]. The 3,7-dioxobicyclo[3.3.0] and methylenedioxy moieties have been proposed as being important functionalities for the potent inhibitory activity of furofuran lignans, although the lack of a methylenedioxy group on the aryl ring of this compound type, as in 399 and 400, decreases the activity observed. In addition, this study also suggested that ring cleavage of furofurans to tetrahydrofuran diols (391, 392), as well as esterification to the diol (391a) or triol (391b), dramatically weakened the inhibitory effects. In addition, antimelanin deposition activities without any concomitant cytotoxicity were found for 7,9-dihydroxydolichanthin B (404) and (2α,3α)-olean-12-en-28oic acid (411), which exhibited IC 50 values of 18.4 and 11.2 μM, respectively [483]. Dose-dependent antimelanin deposition activities have also been shown for oleanonic acid (410), with an IC 50 value of 17.7 μM, although this compound possessed very slight cytotoxicity against the B16-F10 cell line at concentrations above 50 μM in a manner similar to that in a previous study [489]. Moreover, investigation of the effects of 404 and 411 on mushroom tyrosinase as well as on Tyr, Mitf , Trp-1, and Trp-2 mRNA expression suggested that the antimelanin deposition activity of 396 was caused by the downregulation of Tyr mRNA expression, while that of 411 was caused by the downregulation of Mitf mRNA expression located upstream of Tyr, Trp-1, and Trp-2 in the cAMP pathway, although both compounds lacked the ability to inhibit tyrosinase activity [490]. In contrast, compounds 401–403, 405, and 407 have been found to be the active compounds that enhance melanin production [482]. Among them, compounds 403 and 405 increased melanin production by 31 and 50% at a concentration of 50 μM, respectively, while 393 and 397 showed slight cytotoxicity at a concentration of 100 μM, which would account for the reduction of melanin production. Furthermore, compounds 402, 403, and 407 exhibited dose-dependent enhancement of melanin production by 67%, 30%, and 45%, respectively, at a concentration of 100 μM, compared with cells treated only with IBMX and α-MSH [482]. The results from these studies suggest that the chemical types of the constituents reported from P.

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integrifolia are important for melanin deposition or melanin-enhancing effects, but the mechanism(s) of action of the most potent compounds in this regard should be clarified.

2.24 Picrorhiza kurroa Royle ex Benth. Picrorhiza kurroa is a small perennial medicinal herb belonging to the family Scrophulariaceae. However, this is an unresolved name on www.theplantlist.org where it is currently classified in the Plantaginaceae [491]. The plant is distributed in the eastcentral parts of Myanmar, and in India, the Himalayan region (Garhwal to Bhutan), southeast Tibet, west China, and Sri Lanka. P. kurroa, known in Myanmar as Saungmay-ga [492] (Fig. 81), is an important medicinal plant and a common component of TMF-8 (Thway-dular-hsay), TMF-17 (Thway-hsay-ni-gyi), TMF-25 (Ma-te‘-myintmo-hsay), TMF-37 (Kat-pu-yar-di-thu-dathana-hsay), and TMF-38 (Kat-pu-yar-dimyin-thay-myat-ke‘-hsay), which have been used for blood purification and the treatment of fevers, liver cirrhosis, diarrhea, and dysentery [493]. The crude extracts and

Fig. 81 The crude drug Picrorhiza kurroa (stems) known as Saung-may-ga. The stems and roots have use in Myanmar traditional medicine

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O OH

OH 412 (saungmaygaoside A) R = COOH 413 (saunmaygaoside B) R = CHO 414 (saungmaygaoside C) R = CH(OMe)2 415 (saungmaygaoside D) R = CH2OH

Fig. 82 Structures of the new bis-iridoid glycosides 412–415 isolated from a n-butanol extract of P. kurroa stems grown in Myanmar

constituents of this plant have been reported to have cytotoxic [494], hepatoprotective [495], anti-inflammatory [496], anti-asthma [497], immunostimulatory [498], free-radical scavenging [499], and anti-hepatitis [500] activities. The rhizomes of P. kurroa contain iridoids [501], acetophenones [502], and cucurbitacins [503–505] as major constituents, with picrosides I and II being the major bioactive compounds [506–508]. An investigation of the stems of P. kurroa collected in Myanmar was reported in 2017 [509], and a n-butanol extract inhibited the expression of Vpr in TREx-HeLaVpr cells with effective doses of 5 and 10 μg/cm3 [509]. Ten compounds, comprising four new bis-iridoid glycosides, saungmaygaosides A–D (412–415) [509] (Fig. 82), and six known compounds (abelioside A methyl acetal (416) [510], abelioside A (417) [510], sweroside (418) [511], 8-epi-loganin (419) [512], 8-epi-loganic acid (420) [513], and sylvestroside IV dimethyl acetal (421) [514]) (Fig. 83), were isolated from this active n-butanol extract, using a combination of various chromatographic techniques. In an earlier contribution, [509], the structures and names of 416 and 417 were incorrectly reported as abeliosides A and B, so herein they have been corrected as indicated above. The dimethyl acetals 414, 416, and 421 were suggested to be extraction artifacts, since methanol was often used throughout the extraction and isolation processes. From the work on the sample of P. kurroa from Myanmar, some of the isolated iridoids were found to inhibit the expression of Vpr in TREx-HeLa-Vpr cells [509]. At a 5 μM treatment dose, sylvestroside IV dimethyl acetal (421) and sweroside (418) exhibited the most potent activities, followed by saungmaygaoside D (415), 8-epi-loganic acid (420), saungmaygaoside C (414), and abelioside A (417). The potencies of 418 and 421 were comparable to that of the positive control damnacanthal. However, the other isolates, saungmaygaosides A (412) and B (413), abelioside

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O

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O

O

O

O

R O

O O

O HO HO HO

HO HO HO

O OH

O OH 418 (sweroside)

416 (abelioside A methyl acetal) R = CH(OMe)2 417 (abelioside A) R = CHO

O O

O

OR O

O HO

O

O

O

O

O

O

O O

HO HO HO

O OH

419 (8-epi-loganin) R = Me 420 (8-epi-loganic acid) R = H

HO HO HO

O OH 421 (sylvestroside IV dimethyl acetal)

Fig. 83 Structures of the known iridoid and bis-iridoid glycosides 416–421 isolated from a nbutanol extract of P. kurroa stems grown in Myanmar

A methyl acetal (416), and 8-epi-loganin (419), did not inhibit the expression of Vpr. Similar effects were also observed when the cells were treated with a 10 μM dose. Notably, the presence of the secoiridoid moiety in the bis-iridoid glycosides 412, 413, and 415 was unexpected, because these kinds of compounds have not been reported previously as constituents of P. kurroa. Two main routes have been proposed for the biosynthesis of iridoids: route I involves iridodial, iridotrial, and deoxyloganic acids derived from precursors of many carboxylic iridoids with 8β-stereochemistry, including loganin, loganic acid, secologanin, secologanic acid, secoiridoids, and monoterpenoid indole alkaloids. In turn, the precursors in route II are 8-epi-iridodial, 8-epi-iridotrial, and 8-epi-deoxyloganic acid [515, 516]. Although the precursor loganin was not obtained from P. kurroa stems grown in Myanmar, sweroside (418) is regarded as an intermediate in the biosynthesis of the seco-iridoid portion of 412, 413, 415, and 417, and was isolated also from this same stem sample. Thus, Win et al. proposed that bis-iridoid glycosides 412, 413, and 415 are biosynthesized via biosynthesis routes I and II (Scheme 1), whereas 419 and 420 are derived from the biosynthesis pathway only using route II [509]. This study was the first report not only of the isolation of bis-iridoid glycosides generated by dimerization of secoiridoid and 8-epi-loganin units from the stems of P. kurroa from Myanmar, but also of bis-iridoid and iridoid glucosides with anti-Vpr inhibitory activities.

216

N. N. Win and H. Morita O

route I

COOH

O

O

O

OH

OGlc

OH

iridodial

iridotrial

deoxyloganic acid

HO

HO

O

O

O

OGlc

OGlc

OGlc

loganin

loganic acid

secologanin

O

route II

O

COOH

OGlc

OH 8-epi-iridotrial COOH

HO

O

O

OH 8-epi-iridodial

O COOMe

COOMe

COOH

O OGlc

8-epi-loganic acid

8-epi-deoxyloganic acid COOMe

HO

O OGlc 8-epi-loganin

Scheme 1 Biosynthesis pathway of iridoid precursors secologanin (route I) and 8-epi-loganin (route II)

2.25 Jatropha multifida L. Jatropha multifida, a member of the family Euphorbiaceae, is a tree 2–3 m in height that is distributed widely in subtropical and tropical areas throughout Asia and Africa [517]. It is known popularly known as Say-ma-khan in Myanmar (Fig. 84) and has been used in folk medicine as a purgative and to treat fevers, indigestion, colic, wounds, and skin infections. This plant is one of the common ingredients in TMF-16 (A-pu-thet-hsay) and TMF-27 (Pyi-lone-chan-thar-hsay-gyi). These formulations are prescribed to treat various fevers, influenza, diarrhea, hemorrhoids, paralysis, gout, and also as a diuretic [518]. The seed oil, latex, and leaves are effective purgatives and abortifacients and have been used as wound dressings and for the treatment of neurodermatitis, eczema, and itch [517]. The roots and

Bioactive Compounds from Medicinal Plants in Myanmar

217

Fig. 84 Jatropha multifida is known as Say-ma-khan. The stems are used in Myanmar traditional medicine

stems have been reported to have antimicrobial, antimalarial, antitumor, antileishmanial, antiulcer [517, 519], anti-inflammatory, and antioxidant activities [520]. In addition, J. multifida biosynthesizes several characteristic types of secondary metabolites, such as acylphloroglucinols (multifidol and multifidol glucoside) [521], diterpenoids (multifolone, (4E)-jatrogrossidentadione acetate, jatrophone, citlalitrione, 3β-acetoxy-12-methoxy-13-methyl-podocarpa-8,11,13-trien-7-one, (4E)-jatrogrossidentadione, 15-epi-(4E)-jatrogrossidentadione [522], jatromulone A, 3β,12-dihydroxy-13-methylpodocarpane-8,10,13-triene, (2S,3R,5S,10R)-2,3dihydroxy-15,16-di-nor-ent-pimar-8,11,13-triene, (2S,3R,5S,10R)-2-acetoxy3-hydroxy-15,16-di-nor-ent-pimar-8,11,13-triene, gossweilone, jatrointelone A, jatrophodione A, 1,2-dihydroheudelotinol, heudelotinone) [523], and a coumarinolignan (cleomiscosin A) [524]). In 2017, the anti-influenza effects of different extracts of a sample of J. multifida native to Myanmar were reported [525]. The survival of influenza A H1N1 virus (A/PR/8/34)-infected MDCK cells was evaluated with extracts of different polarities

218

N. N. Win and H. Morita O O

R

O

RO

O

R2

O

361 (scopoletin) R = H 422 (scoparone) R = Me

O

423 (salicifoliol) R1 = H, R2 = OH 424 (glaberide I) R1 = OMe, R2 = H O

O OH O

O

HO

1

R

R2

O

1

OH O

427 (matairesinol)

O

O HO

HO O

249 (syringaresinol) R1 = R2 = OMe 425 (pinoresinol) R1 = R2 = H 426 (medioresinol) R1 = H, R2 = OMe

OH OH

HO O

O

428 (secoisolariciresinol)

Fig. 85 Structures of the coumarins 361 and 422 and the lignans 423–428 isolated from a chloroform extract of P. multifida stems

(n-hexane, CHCl3 , EtOAc, and H2 O) prepared from J. multifida stems. These were subjected to naphthol blue black staining and determinations of MDCK cell viability (with a MTT assay), influenza A viral infection and growth (using an immunofluorescence staining method), and viral growth in virus-infected MDCK cells (by measuring viral titers in an influenza A viral growth assay) [525]. In the viral infection and cell viability assays, an increase in the survival of influenza A virus-infected MDCK cells and inhibition of influenza A viral binding to host cells was shown for the H2 O extract when the influenza virus and 3.1–25 μg/cm3 of each extract was co-incubated in advance. In contrast, in the viral growth assay the CHCl3 extract at a concentration of ≤12.5 μg/cm3 inhibited influenza A viral replication in host cells. Thus, Shoji et al. proposed that the H2 O extract could include compounds that inhibit influenza A viral binding to the host cell surface, endocytosis, membrane fusion, or uncoating by inhibiting viral HA, while the CHCl3 extract includes compounds that inhibit influenza viral replication in the host cells by inhibiting viral RNA polymerase or NA activities. These results indicated that the different polarity H2 O and CHCl3 crude extracts of J. multifida inhibited viral infection or growth by different mechanisms. In a later study, a moderate anti-melanin deposition activity (IC 50 value of 22.7 μg/cm3 ) was observed for the CHCl3 extract of the J. multifida stems [526]. Furthermore, subsequent isolation of the active CHCl3 extract furnished nine compounds, including two coumarins, scoparone (422) [527] and scopoletin (361) [528], and seven lignans salicifoliol (423) [529], glaberide I (424) [530], pinoresinol (425), medioresinol (426), syringaresinol (249) [531], matairesinol (427) [528], and

Bioactive Compounds from Medicinal Plants in Myanmar

219

secoisolariciresinol (428) [532] (Fig. 85). Among the isolated compounds, glaberide I (424) and secoisolariciresinol (428) displayed anti-melanin deposition activities, with IC 50 values of 49.9 and 37.5 μM, respectively, without showing any cytotoxicity in the anti-melanin deposition assay against the α–MSH- and IBMX-induced mouse melanoma cell line (B16-F10) [526]. Further detailed analyses of the effects of 424 and 428 on tyrosinase activity, as well as MITF, TYR, TRP-1, and TRP2 mRNA expression, have suggested that the anti-melanin deposition activity of 428 could result from the downregulation of Tyr mRNA expression, while the antimelanin deposition activity of 424 may result from the inhibition of other melanogenesis pathways. J. multifida does not appear to be used as a traditional cosmetic. However, the results of this study may be helpful for the application of J. multifida and its phytoconstituents as one of the ingredients of potential skin-whitening agent formulations.

2.26 Swertia chirata Buch.-Ham. ex Wall. Swertia chirata is an annual/biennial medicinal herb (Gentianaceae) and is distributed widely in the Himalayan mountains, Pakistan, India, Nepal, Bhutan, Tibet, and Myanmar. In Myanmar, this plant is referred to as Pan-khar or Thinbaw-sega-gyi (Fig. 86) and it occurs at Taungoo Township in the Bago Region. It has been utilized in traditional medicine for the treatment of liver disorders, malaria, chronic fever, anemia, bronchial asthma, hepatitis, diabetes, cancer, and HIV/AIDS [533, 534]. Previous phytochemical studies have reported the presence of xanthones, flavonoids, terpenoids, alkaloids, iridoids, secoiridoids, steroids, and phenolic compounds from whole plants of S. chirata. These secondary metabolites showed antitumor, antiviral, antidiabetic, anti-HIV, and anti-hepatitis bioactivities [533–536]. A report appeared in 2019 on the phytochemistry of the whole plants of S. chirata collected in Myanmar [537]. The CHCl3 -soluble extract of S. chirata inhibited the expression of Vpr at an effective concentration level of 10 μg/cm3 . Four xanthones, decussatine (429) [538], methylbellidifolin (430) [539], 3,5-dimethoxy1-hydroxyxanthone (431) [540], and bellidifolin (432) [541], and two triterpenoids, oleanolic acid (382) [454] and 12-hydroxyoleanolic lactone (433) [542], were isolated from this active CHCl3 extract (Fig. 87). In the reported paper [537], the name of 430 was incorrectly given as methylswertianin, and, herein, it has been corrected as methylbellidifolin. In this study, the isolated compounds 382, 429, 430, 432, and 433 were examined for their inhibitory effects on the expression of Vpr in TREx-HeLa-Vpr cells, conducted according to a previously reported protocol [359]. Among the compounds tested, only oleanolic acid (382) and bellidifolin (432) exhibited anti-Vpr activities. Bellidifolin (432) and oleanolic acid (382) inhibited Vpr expression at a 5 μM concentation level. Similar inhibitory effects were observed when the cells were treated with a 10 μM dose. In particular, 10 μM oleanolic acid (382) exhibited more

220

N. N. Win and H. Morita

Fig. 86 The crude drug Swertia chirata (whole plant). The original plant is known as Pan-khar or Thinbaw-sega-gyi, and the whole plant is used in traditional medicine in Myanmar

potent activity than that of the positive control, damnacanthal. In another investigation, Kashiwada et al. reported [543] that 382 inhibited HIV-1 replication in acutely infected H9 cells (EC 50 value of 1.7 μg/cm3 ) and inhibited H9 cell growth (IC 50 value of 21.8 μg/cm3 ). The finding of oleanolic acid as a Vpr inhibitor is supportive of further studies being carried out on compound 382 and/or its derivatives as possible anti-HIV agents. Naturally occurring xanthones have emerged as an important class of organic compounds in terms of their documented pharmacological and biological activities. The structure-activity relationship conclusions among the isolated xanthones (decussatine (429), methylbellidifolin (430), and bellidifolin (432)), suggested that the presence of hydroxy groups at C-5 and C-8 in 432 favored the inhibition of the expression of Vpr, rather than the presence of the methoxy substituents at C-5, C-7, and C-8 in 429 and 430, which led to decreases in their activities. Bellidifolin (432) is a simple oxygenated xanthone that reportedly possesses strong hypoglycemic activity [539] and monoamine oxidase inhibitory activity [544]. This compound also reportedly inhibits the production of the proinflammatory cytokines IL-6 and tumor necrosis factor-α (TNF-α) and the production of prostaglandin E2 (PGE2) by suppressing the protein expression of COX-2 in LPS-stimulated RAW264.7 macrophages [545]. Treatment with 432 also reportedly suppressed the phosphorylation of the inhibitor κB kinase-β (IKK-β), Akt, and the p65 subunit of NF-κB [546]. Thus, these Vpr inhibition findings of 432 have provided a new insight into its bioactivity. The detection

Bioactive Compounds from Medicinal Plants in Myanmar R1

O

221

OH

R2 O

O

R3

429 (decussatine) R1 = R2 = OMe, R3 = H 430 (methylbellidifolin) R1 = OH, R2 = H, R3 = OMe 431 (3,5-dimethoxy-1-hydroxyxanthone) R1 = R2 = H, R3 = OMe 432 (bellidifolin) R1 = R3 = OH, R2 = H

OH O

CO

OH O HO

HO

382 (oleanolic acid)

433 (12-hydroxyoleanolic lactone)

Fig. 87 Structures of the xanthones 429-432 and the terpenoids 382 and 433, isolated from a chloroform extract of Swertia chirata whole plants

of anti-Vpr activity in this plant is in good agreement with ethnopharmacological reports of its anti-HIV, antiviral, and anti-hepatitis activities [534, 546].

3 Synthesis Aspects As described in Sect. 2, 433 compounds identified from Myanmar medicinal plants have been described, and regardless of whether these were new or known previously, the chemical syntheses of several of these compounds have been carried out, as indicated by the following examples. Panduratin A (179), a cyclohexenyl chalcone with antiausterity activity from Boesenbergia pandurata, was synthesized via six steps including a high pressure Diels-Alder reaction. The related natural products, panduratins H (172) and I (173), 2-hydroxyisopanduratin A (174), 4-hydroxypanduratin A (181), and nicolaioidesin B (186) were also produced using the same general procedures (Schemes 2 and 3) [547]. Of the active coumarins from the flowers of K. assamica, the more recently described geranylated and isoprenylated coumarins remain to be synthesized. However, the known active coumarins 220–225 have been synthesized in order to confirm the chemical structures proposed at the time of their initial structure elucidation. Pechmann condensation was applied to synthesize members of those classes of alkylated or phenylated coumarins. Compound 220 has been partially synthesized from mammeisin (Scheme 4) [548], while 221 was synthesized from 5,7-dihydroxy-4-phenyl-2H-[1]benzopyran-2-one as the key starting

222

N. N. Win and H. Morita

O O

O

19 kbar CH2Cl2 RT, 3 d 93%

O

O

O +

methyl cinnamate

1:2.9 172 (panduratin H)

173 (panduratin I)

LiAlH4, THF RT, 2 h

O

79%

OH DMP, CH2Cl2 RT, 1 h 78%

HO +

MOMO i) nBuLi, MOM-bromobenzenes THF, -78 °C; R ii) DMP, CH2Cl2 RT, 2 h

MOMO R MOMO

Br

R = OMe or OMOM MOM:methoxymethyl ether

MOMO

OH O

HCl, MeOH 50°C, 2 h 179: 70% 181: 71%

O R OH

179 (panduratin A) R = OMe 181 (4-hydroxypanduratin A) R = OH

Scheme 2 Synthesis of panduratins H (172), I (173), A (179), and 4-hydroxypanduratin A (181)

material (Scheme 5) [242]. Compound 222 was semi-synthesized from 6-butyryl5,7-dihydroxy-4-propylcoumarin (Scheme 6) [244], and further converted to 225 (Scheme 6) [244]. Compounds 223 and 224 were generated from 6-butyryl5,7-dihydroxy-4-phenylcoumarin (Scheme 7) [245]. The synthesis of additional bioactive compounds from Myanmar medicinal plants is expected in the future.

Bioactive Compounds from Medicinal Plants in Myanmar

O

223 O

19 kbar CH2Cl2 RT, 3 d 62%

O +

cinnamaldehyde

1:7.2

i) n-BuLi, MOM-bromobenzenes THF, –78°C; R ii) DMP, CH2Cl2 RT, 2 h

OH

MOMO Br MOMO

O

R = OMe or OMOM MOM:methoxymethyl ether

R HCl, MeOH 50°C, 2 h

OH

174: 71% 186: 69%

MOMO O R MOMO

174 (4-hydroxypanduratin A) R = OH 186 (nicolaioidesin B) R = OMe

Scheme 3 Synthesis of 2-hydroxyisopanduratin A (174) and nicolaioidesin B (186) O

OH

HO

O

Ph

O

DDQ O benzene, RT, 2 h

mammeisin

Scheme 4 Synthesis of mammea A/AA cyclo D (220)

O

OH

Ph

O

O

220 (mammea A/AA cyclo D)

224

N. N. Win and H. Morita OH

Ph

O

HO

OH

O

CH3CH2CH2COCl/AlCl3/ CS2,CH3NO2/reflux

O

HO

5,7-dihydroxy-4phyenyl-2H[1]benzopyran-2one

Ph

O

O

(CH3)2C=CHCH2Br/10% KOH/0°C

OH

Ph

O

HO

O

O

221 (mammea A/BC)

Scheme 5 Synthesis of mammea A/BC (221)

O

O

OH

OH

prenyl bromide HO

O

6-butyryl-5,7dihydroxy-4propylcoumarin

O

2,2,2-trifluoroethanol 10% aq KOH 0 °C, 90 min

HO

O

O

222 (mammea B/AC) m-chloroperbenzoic acid CH2Cl2, RT, overnight

O

OH

O

O

HO 225 (mammea B/AC cyclo F)

Scheme 6 Synthesis of mammea B/AC (222) and mammea B/AC cyclo F (225)

Bioactive Compounds from Medicinal Plants in Myanmar HO

dioxane, BF3-etherate, 50°C, 90 min O

OH

225 O

OH

Ph

O

HO

O

Ph 223 (mammea A/AC)

HO

O

O

6-butyryl-5,7dihydroxy-4phenylcoumarin O

OH

Ph

O pyridine, 110°C, 10 h

O

O

O

224 (mammea A/AC cyclo D)

Scheme 7 Synthesis of mammea A/AC (223) and mammea A/AC cyclo D (224)

4 Conclusions Human health and well-being are to a great extent directly dependent on biodiversity. The southeast Asian country of Myanmar has a high level of biodiversity distributed within a wide variety of habitats, including lowland tropical forests, mangroves, and marine ecosystems. Various species of medicinal plants occur in Myanmar, and local inhabitants have been using these plants for a long time based on traditional knowledge forwarded through generations. In drug discovery, it is becoming a challenging scientific task to find robust and viable lead molecule candidates, and one approach to this process flows from the screening of natural product extracts to new isolates, which requires expertise and experience. Natural products have afforded key compounds in drug discovery and development in the past. Anticancer drugs such as Taxol (paclitaxel; Taxus brevifolia) and vinblastine (Catharanthus roseus) and antimalarial drugs such as quinine (Cinchona spp.) and artemisinin (Artemisia annua) were all discovered from plant-derived natural products and they or their synthetic derivatives are effective in treating the respective diseases mentioned above. During the last two decades (2000–2020), scientists from Myanmar, Japan, Germany, and Korea have investigated medicinal plant samples found in Myanmar that are constituents of local traditional medicinal preparations. Of these, after screening, 29 samples were found to be active in the respective bioassay system or systems used. Altogether, 433 compounds were obtained from these active samples, of which 147 compounds were characterized as new compounds at the time of their initial isolation, and their structures were published in international peer-reviewed journals. The types of compounds to have been discovered are classifiable into monoterpene

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lactones, diterpenoids, sesquiterpene lactones, triterpenoid saponins, fatty acids, and esters, various types of flavonoids (cyclohexenyl chalcones, chalcones, flavanones), coumarins, xanthones, quinones, quinols, curcuminoids, benzoic acid derivatives, and lignans. These structurally diverse phytochemicals were found to possess various pharmacological effects, such as cytotoxicity, NO inhibitory activity, antimalarial activity, antitrypanosomal activity, anti-austerity activity, leishmanicidal activity, HIV-1 viral protein R inhibitory activity, antimelanogenesis activity, and antibacterial activity. On the basis of internationally recognized documentation, only about 0.25% of the 11,800 plant species listed as indigenous, or are naturally occurring or cultivated in Myanmar, have been investigated in a scientific manner so far. Thus, further efforts and investment should be made to investigate the remaining Myanmar medicinal plants in order to potentially discover new drug candidate molecules. The bioactive compounds obtained from the medicinal plants of Myanmar may serve also as lead compounds for new analogs with either improved therapeutic activity or reduced toxcity. Acknowledgments The authors are grateful to Professors Douglas Kinghorn and Heinz Falk for their enormous assistance and encouragement with the preparation of the manuscript and whose contributions to this volume went far beyond their editorial duties. The authors would like to express their gratitude to Professor Yoshinori Asakawa (Tokushima Bunri University), Professor Ikuro Abe (University of Tokyo), and Professor Dr Daw Hla Ngwe (Yangon University) for valuable suggestions. The authors also acknowledge the researchers from Myanmar, Germany, Japan, and Korea for their great contributions to the scientific research publications regarding the medicinal plants from Myanmar. Special thanks are owed to Professor Dr Ni Ni Than (Yangon University), Dr Myint Myint Than (Department of Traditional Medicine), Dr Yi Yi Win (Dawei University), Dr Khine Zar Wynn Lae (Yangon University), and Dr Ei Ei Thwin (Taunggoke Degree College), who helped to take the photographs of the Myanmar medicinal plants.

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Nwet Nwet Win studied chemistry in Myanmar and obtained a B.Sc. (Hons.) degree with first-class honors and a M.Sc. degree with credit, both from Mawlamyine University. She then received further graduate degrees from Dagon University, Myanmar (M.Res.), Yangon University (Ph.D.), and the University of Toyama, Japan (Ph.D. in pharmaceutical sciences). She has been actively involved in the discovery of bioactive compounds from medicinal plants of Myanmar since she was a doctoral student with Professor Daw Hla Ngwe at the Department of Chemistry, University of Yangon, Myanmar (2002– 2004) and then with Professor Shigetoshi Kadota (2005–2008), and also as a postdoctoral fellow with Professor Hiroyuki Morita (2014–2018), both of them at the Institute of Natural Medicine, University of Toyama. In her continuing investigations of biologically active medicinal plant constituents from Myanmar, she has focused on potential anti-austerity, anti-Vpr, antineoplastic, and anti-melanogenesis activities. She has authored and coauthored 34 international papers and approximately 30 domestic publications. Throughout her career, she has been awarded scholarships from the Japanese Government (Monbukagakusho, MEXT), the Tokyo Biochemical Research Foundation, the Takeda Science Foundation, and the Japan Society for the Promotion of Science (JSPS). She is a member of the Japanese Society of Pharmacognosy and the Pharmaceutical Society of Japan. She worked 20 years (1998–2018) for the National Universities of Myanmar, such as at Dagon University, Taungoo University, and Yangon University, starting as a Demonstrator, and moving to Assistant Lecturer, Lecturer, and Associate Professor positions. Currently, she has been a researcher at the Institute of Natural Medicine, University of Toyama since 2018. Hiroyuki Morita obtained his Ph.D. in 2001 from the University of Shizuoka under the direction of Professor Hiroshi Noguchi. After one year of postdoctoral research with Professor John C. Vederas at the University of Alberta in Canada (2001–2002), he returned to Japan to investigate the structural enzymology of protein kinases and plant polyketide synthases as a Postdoctoral Fellow at Mitsubishi Chemical Corporation (2002–2004) and Mitsubishi Kagaku Institute of Life Sciences (2004–2008). In 2008, he returned to the University of Shizuoka as an Assistant Professor (2008–2009) and then moved to the Laboratory of Natural Products Chemistry at the Graduate School of Pharmaceutical Sciences, The University of Tokyo, as an Assistant Professor (2009–2012). Currently, he is Professor at the Institute of Natural Medicine, University of Toyama (2012–). His current research is focused on the study of the biosynthesis of natural products derived from plants, including X-ray crystal structure analysis and engineering of secondary metabolite enzymes involved in the biosynthesis of natural products to produce new compounds, as well as the isolation of bioactive compounds from natural sources such as plants and marine organisms.

New Techniques of Structure Elucidation for Sesquiterpenes Julio C. Pardo-Novoa and Carlos M. Cerda-García-Rojas

Contents 1 2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Techniques Based on NMR Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Calculation of Chemical Shifts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Calculation of Coupling Constants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Residual Dipolar Couplings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Advanced Mosher’s Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Use of Cryoprobes for NMR Measurements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Techniques Based on Chiroptical Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Vibrational Circular Dichroism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Electronic Circular Dichroism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Techniques Based on X-Ray Diffraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Calculation of the Flack Parameter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Calculation of the Flack and Hooft Parameters . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

254 254 255 262 268 270 274 276 276 280 291 292 298 303 304

J. C. Pardo-Novoa · C. M. Cerda-García-Rojas (B) Department of Chemistry, Centro de Investigación y de Estudios Avanzados del Instituto Politécnico Nacional, Avenida Instituto Politécnico Nacional 2508, Mexico City 07360, Mexico e-mail: [email protected] J. C. Pardo-Novoa e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 114, https://doi.org/10.1007/978-3-030-59444-2_3

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1 Introduction Throughout the history of Chemistry, a series of techniques has been developed to decipher the molecular structure of new compounds of natural origin, starting from chemical degradation up to highly sophisticated NMR methodologies in combination with quantum mechanical calculations of their spectroscopic properties. This contribution reviews the most relevant techniques that have been used recently in determining the structure of a group of representative natural products such as the sesquiterpenes. Essentially, the papers dating back to this new century will be referenced from January 2001 until May 2020. A frequent characteristic of these new methodologies consists of the combined use of precise experimental determinations and accurate theoretical data obtained by molecular modeling calculations for a property of a given compound. This has been particularly valuable for nuclear magnetic resonance (NMR) spectroscopy that has expanded its potential for solving complex structural problems. At this time, it is possible to say that if the structure of a studied new compound is correct, its calculated data will easily match with the experimental ones, thus validating the structural hypothesis for a given molecule. Some other lines of research on sesquiterpenes, particularly those dedicated to the determination of their absolute configuration, involve the comparison of calculated and experimental chiroptical properties, namely, electronic and vibrational circular dichroism as well as the use of X-ray diffraction analysis with emphasis on calculation of anomalous dispersion effects, as applicable to all organic molecules that crystallize in non-centrosymmetric space groups, not only to those containing “heavy” atoms. Although many investigations described in this chapter utilize more than one modern technique for the structure elucidation of isolated molecules, they are discussed in the section that is considered as the most pertinent and illustrative for each case.

2 Techniques Based on NMR Spectroscopy The power of NMR spectroscopy has been enhanced greatly by its combination with theoretical parameters calculated using quantum mechanics mainly with Density Functional Theory (DFT) and Hartree–Fock (HF) protocols. In most cases, the calculated 1 H and 13 C chemical shifts and coupling constants (J) are compared with the experimental values by means of basic statistical analysis allowing the determination of the correct configuration among several structural hypotheses. Other strategies include the measurement of residual dipolar couplings that generate information about the orientations between internuclear vectors, in correlation with the molecular three-dimensional arrangement of a structure. This section also comprises the review of modified Mosher methods in NMR tubes that allow the use of small amounts of materials and the combination of this methodology with molecular models for the evaluation and visualization of the diamagnetic shielding effects present in the ester derivatives. Several articles about the use of cryoprobes that have enhanced the power

New Techniques of Structure Elucidation for Sesquiterpenes

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of NMR spectrometers to unprecedented levels of quantitation and accuracy are also included.

2.1 Calculation of Chemical Shifts A remarkable example of the application of these NMR-DFT methodologies was described for the structural determination of a triquinane derivative isolated from Pulicaria vulgaris [1]. The structure of this sesquiterpenoid was deduced as presilphiperfolane-7α,8α-diol (1) by combining solvent-induced removal of chemical shift degeneracy and computational DFT-gauge including atomic orbital (GIAO) prediction of NMR spectra with the analysis of 1 H NMR splitting patterns obtained by the full spin simulation using the MestReNova program [2]. The chemical shifts and coupling constants of presilphiperfolane-7α,8α-diol (1) and seven other geometryoptimized diastereomers were calculated at the DFT B3LYP 6-311++G(d,p) level of theory and the solvent effects were modeled by the polarizable continuum model selfconsistent reaction field (PCM-SCRF) protocol. The values of correlation coefficients and root mean-square errors between calculated and experimental data supported 1 as the correct structure [1]. All DFT calculations were accomplished using the Gaussian 09 program package [3]. Altogether nine guaianolides, some of them with anti-inflammatory properties, were isolated from the aerial parts of Ormenis mixta collected from Constantine, Algeria [4]. Of this group of compounds, the stereochemical assignments of 2–4 were based mainly on a combined quantum mechanical/NMR approach by comparison of the experimental 13 C/1 H NMR chemical shifts and J H-H coupling constants with the predicted values. The calculations for each compound involved the conformational search of all possible diastereoisomers of 2–4 followed by optimization of their geometries and calculation of the 13 C and 1 H NMR chemical shifts for every selected conformer using the MPW1PW91 functional and 6-31G(d,p) basis set in methanol also using the Gaussian 09 program [3]. Calculations of the weighted averaged chemical shifts were carried out taking into account the Boltzmann distributions according to the conformational energies and considering the calculated chemical shifts of tetramethylsilane as the reference compound [4].

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HO

O

OH

OH

HO

HO O O

1 (presilphiperfolane-7,8-diol)

O

2

OH

HO

O

OH

HO

HO

HO

O

O

O

O 4

3

The structures of the sesquiterpenoids corianlactone (5) and boletunone B (6) were studied by comparing their 1 H and 13 C NMR chemical shifts and coupling constants with those calculated with several DFT and second-order Møller-Plesset (MP2) perturbation theory methods [5]. It was of particular interest to validate the case of boletunone B (6), which was the subject of a structural revision. Furthermore, it was relevant to find that including the solvent by means of the Integral Equation-Formalism Polarizable Continuum Model (IEF-PCM) led to a substantial improvement in the comparisons of theoretical and experimental data [5]. O

O

O

CO2H

O O O

O O

5 (corianlactone)

OH O 6 (boletunone B)

Five sesquiterpene lactones of the guaianolide-type (7–11) were studied in solution and in the solid state using NMR spectroscopy in combination with theoretical calculations of their chemical shifts together with X-ray diffraction analysis [6]. The experimental 1 H and 13 C chemical shifts in solution were efficiently calculated using the GIAO method at the DFT B3LYP/6-31++G** level of theory, while for the solid state, the gauge-including projector-augmented wave (GIPAW) method [7] was used for calculations of the 13 C chemical shifts with remarkable results. The study in the solid state revealed that mexicanin I (7) was a standard case, with only one symmetrically independent molecule in the unit cell, while helenalin (8) and 6α-hydroxydihydroaromaticin (9) showed the presence of two polymorphs. Geigerinin (10) exhibited multiple asymmetric units in the crystal lattice, in which the symmetry-independent molecules were connected by a series of hydrogen bonds, while symmetry-independent molecules of badkhysin (11) differed in the conformation of the sidechain [6].

New Techniques of Structure Elucidation for Sesquiterpenes

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O

O O

O

HO

O

7 (mexicanin I)

8 (helenalin)

HO

O O

O

HO

O

HO

O O

HO

9 (6-hydroxydihydroaromaticin)

10 (geigerinin)

O

O O O O 11 (badkhysin)

Two new germacranes named ketopelenolides C (12) and D (13) were isolated from Artemisia arborescens, collected in Cagliari, Italy. Crucial points in their structure elucidation were solved by NMR spectroscopy in combination with molecular modeling and quantum–mechanical calculations including comparisons between experimental and DFT-calculated 13 C NMR data [8]. For the configurational assignment of these structures, which comprise highly flexible medium-sized rings, a detailed conformational analysis was undertaken. The calculation protocol involved geometry optimization of all possible conformers followed by calculation of chemical shifts with the GIAO approach using the DFT MPW1PW91/6-31G(d,p) method [8]. HO

O

O

O O O 12 (ketopelenolide C)

O O 13 (ketopelenolide D)

The guaiane-type sesquiterpenoids 14–18 were isolated from Daphne genkwa collected in Xingyi City, Guizhou Province, People’s Republic of China [9]. Calculation of their 13 C NMR chemical shifts was achieved using the GIAO method at the mPW1PW91/6-311+G(d,p) level of theory, while application of the statistical method DP4+ [10] was an effective strategy for the assignment of the correct diastereoisomers. Additionally, the absolute configurations were determined by comparing the experimental and calculated electronic circular dichroism spectra [9].

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J. C. Pardo-Novoa and C. M. Cerda-García-Rojas HO

OH

HO

O

O OH

14

O

15

O

OH OH

OH

16

OH

17

O

18

OH

OH

The structures of three new bergamotane (19–21) derivatives were ascertained [11] by the use of 13 C NMR chemical shift calculations in combination with the DP4+ probability method [10]. These tetracyclic sesquiterpenoids, designated as purpurolides D–F (19–21), were isolated from a strain of the endophytic fungus Penicillium purpurogenum. The three metabolites exhibited significant inhibitory activities against pancreatic lipase, although the most potent compound was purpurolide F (21), for which its enhanced activity was attributed to the presence of a 3-hydroxydecanoic acid moiety [11]. HO O R

O O

O O

O

O

O O

O

OH 19 R = OH (purpurolide D) 20 R = H (purpurolide E)

21 (purpurolide F)

An important aspect of the experimental vs. calculated NMR data protocols for structure elucidation is the use of scaling factors. These are essential elements in the conversion of calculated NMR isotropic shielding tensors to chemical shifts. A series of scaling factors was computed for ten DFT methods and ten different solvents were commonly used in NMR. The study included 23 reference compounds and the sesquiterpene lactone mexicanin I (7) to show the utility of such methodology [12]. Another study for the parameterization of scaling factors used ten diverse sesquiterpenes (22–31) that included the antimalarial drug artemisinin (25). This protocol involved GIAO-DFT calculations of chemical shifts with the mPW1PW91/6-31G (d) functional/basis set. Application of these parameterized factors ensured the accurate structural determination for this class of natural products [13].

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259

Artemisinin (25) has been the subject of an investigation that included the 1 H and 13 C NMR calculation of absolute shielding constants with the continuous set of gauge transformations (CSGT) method at the DFT/B3LYP level of theory and the 6-311+G(2d,p) basis set. The calculated magnetic shielding tensors were converted into chemical shifts by taking into account the 1 H and 13 C absolute shielding tensors for tetramethylsilane calculated at the same level of theory [14]. O O

O O

O

O

OH

OH O O

O

23

22

O O

HO

O

O O

O

OH

25 (artemisinin)

24

O O O O

O

HO

O OH

OH 26

O HO

27

O HO

O O

O

OH 29

28 H

O

HN O OH 30

31

The structures of fifteen sesquiterpenoids were revised as depicted in 32–46 as part of an extensive investigation that examined more than one-hundred halogenated compounds [15]. The lack of efficient methods to calculate 13 C NMR data for carbons bearing heavy atoms was resolved by the parametric corrections of DFT-computed

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chemical shifts, which in combination with relativistic force field-computed spin– spin coupling constants, permitted validation or revision for all these compounds. The method was termed as DU8+ [15] and also recommended as a convenient tool for structure validation or revision of terpenoids. O

O

O O Cl

Br Cl

32

33 Br

Cl

OH

HO

OH OH

Br HO

35

Br

34

CHBr2 Br

O

Cl Br

O

O Br

37

36

Br O

OOH

Cl

HO

Br

Br

O

O

HO 39

38

O

Br

40

HO

Cl

Cl O

Br

Br

OH 42

41

Br

43

Br O

O Cl

Cl

Br 44

HO

Cl Br

HO 45

46

The total synthesis of compounds 47–49 allowed the structural revision of dichrocephones A and B [16]. Since spectroscopic data for the synthetic material was different from the natural isolate, a detailed scrutiny was undertaken. NMR calculated spectra using DFT methods, mPW1PW91/6-311+G(2d,p) for geometry optimization and B3LYP/631+G(d,p) for chemical shift calculations, were helpful in revising the relative configuration of these compounds, which was also supported by biosynthesis considerations. Comparison of NMR spectroscopic data and optical rotations of the synthetic compounds with those of the natural products revealed that the structure of dichrocephone A corresponded to ent-49, while that of dichrocephone B is represented by ent-48 [16].

New Techniques of Structure Elucidation for Sesquiterpenes

O

OH OH

O

261

OH

O

O

HO 48

47

49

The relative configurations of the tricyclic sesquiterpenes isohirsut-1-ene (50) and isohirsut-4-ene (51), obtained from an engineered Streptomyces strain, were assigned by comparison of experimental and density functional theory 1 H and 13 C chemical shifts [17]. To achieve this complex task, due to the hydrocarbon nature of the structures, DFT calculations were carried out for eight possible stereoisomers of each compound at the B3LYP/6-31+G(d,p) level of theory followed by scaling the calculated isotropic values for each atom. Mean absolute deviations between the calculated and experimental chemical shifts as well as DP4 [18] statistic values were considered in the discernment of the correct structures [17].

51 (isohirsut-4-ene)

50 (isohirsut-1-ene)

The absolute configuration of a nor-sesquiterpenoid named artarborol (52), isolated from the Mediterranean shrub Artemisia arborescens, was established using a combination of chemical derivatization, NMR data, molecular modeling, and quantum–mechanical calculations [19]. Comparison of the experimental 13 C NMR chemical shifts with those calculated by the DFT MPW1PW91/6-31G(d,p) method validated the stereochemical assignment. Given that the structure contains a ninemembered ring that confers flexibility, the study required the evaluation of all possible conformers and the averaging of their calculated chemical shifts. This protocol was recommended for sesquiterpenes containing flexible medium-sized rings such as germacranes, humulanes, and caryophyllanes that exist in conformational equilibria [19]. O O

HO 52 (artarborol)

53 ((1R,2S,4S,5R,9R,11R)-3-ishwarone)

The structure of (1R,2S,4S,5R,9R,11R)-3-ishwarone (53) isolated from Peperomia scandens was elucidated by comparison between experimental and theoretical NMR data that included chemical shift 13 C NMR calculations for four stereoisomers. The molecular modeling protocol also involved the vibrational frequencies data calculated with the B3LYP functional and the cc-pVTZ basis set [20].

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A conformational analysis of longifolene (54) and a synthetic precursor (55) was carried out by ab initio calculations with the RHF/6-311G(d) method. The study also included the 1 H and 13 C NMR chemical shift ab initio calculations performed at the same level of theory, allowing the stereospecific 1 H NMR assignments of the two molecules [21].

O 54 (longifolene)

55

Molecular modeling of peribysins C and D, using a quantum mechanics approach with an ab initio protocol at the 6-31G(d,p) basis set, allowed revision of their structures as depicted in 56 and 57, respectively. The CAST/CNMR system, which predicts 13 C NMR chemical shifts based on the data for related structures included in a database, was an essential tool to detect the structural misassignment of these bioactive compounds obtained from a strain of Periconica byssoides [22]. O OH

O

OH

OH

O

OH

R1

R2 O O

56 (peribysin C)

57 (peribysin D)

58 R1 = CH3, R2 = OH (vulgarin) 59 R1 = OH, R2 = CH3 (epivulgarin)

An outstanding review regarding the determination of relative configuration in organic compounds, involving NMR and computational approaches, summarized a detailed examination of numerous relevant methods for solving a wide range of stereochemical problems [23]. The methodologies include quantum mechanical calculation of NMR parameters, coupling constants-based analyses, and the Universal NMR Database. The review embraced diverse groups of molecules such as the sesquiterpene lactones vulgarin (58) and epivulgarin (59), for which the configurational assignments were achieved by a molecular modeling conformational search combined with a quantum mechanics protocol that included 13 C NMR chemical shift calculations using the GIAO theory [24].

2.2 Calculation of Coupling Constants Vicinal spin–spin coupling constants (3 J H-H ) have played a fundamental role in establishing the relative configurations and conformational analyses of natural products

New Techniques of Structure Elucidation for Sesquiterpenes

263

due to the dihedral angle dependence [25, 26], which has been revisited theoretically to evaluate its dependence on the Fermi-contact [27]. Calculation of spin–spin coupling constants significantly enhances the stereochemical information provided by the quantum mechanics molecular models, thus allowing the correct assignment of a structure. A systematic analysis to incorporate 3 J H-H couplings into the DP4 formalism (J-DP4) revealed that the combined use of two approaches, one involving a DP4like equation including an additional term given by 3 J H-H and the second that comprises the original DP4 method, but with a restricted conformational search, afforded optimum results considering performance and computational cost [18]. The combined method was tested on a set of 69 examples that included sesquiterpenoids 60–63, and gave good results in the selection of the correct stereoisomer in each case [18]. O OH O O OH O O 60

61(cordycepol A) OH

O

62

OH

O

63

The structure of cordycepol A (61), isolated from the fungus Cordyceps ophioglossoides, was subjected to revision in light of the results provided by the DFT calculation of its NMR spectroscopic parameters. The chemical shifts of 61 were calculated at the mPW1PW91/6-311+G(d,p) level of theory, while the calculated coupling constants were obtained with relativistic force field DU8c parametric methodology, which provided high accuracy with deviations usually below 1 Hz [28]. The structure elucidation of lippifolianones 64–66, isolated from the aromatic shrub Lippia integrifolia, was accomplished by DFT molecular modeling [29]. Their conformational distribution was modeled using the Monte Carlo random search [30] as implemented in the Spartan program (Wavefunction, Inc.). The minimum energy conformers were geometry and energy optimized by using the B3LYP/6-31G(d) level of theory. The dihedral angles for each conformation, measured in the molecular models, were used to calculate the vicinal 1 H–1 H coupling constants by means of a generalized Karplus-type equation [26, 31], followed by Boltzmann-averaging according to the DFT energy, to obtain the calculated coupling constants for each compound. The calculated values were compared with the experimental ones to validate the conformations of 64–66 as listed in Table 1 [29]. Additionally, the absolute configuration of these lippifolianones was assigned by analysis of the circular

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Table 1 Calculated and experimental coupling constants of lippifolianones (64–66), valeranone (69), and morelianone (70)a 3J

H,H /Hz

64 Calc

65 Obs

66

Calc

Obs

Calc

Obs

4.0

4.8

5.8

5.7

2.6

3.3

11.5

11.4

2α,3α

3.5

–

b

2α,3β

3.0

–

b

2β,3α

13.2

11.6

12.3

11.4

2.3

1.7

2β,3β

3.8

5.5

4.2

4.8

5.3

6.3

3α,4

12.3

13.0

11.7

12.8

–

–

3β,4

3.4

5.5

3.2

4.8

–

–

8α,9

2.2

2.0

5.3

5.8

5.6

5.8

8β,9

6.2

5.0

8.4

8.2

8.9

8.3

3J

H,H

3J

69 Calc

Obs

1α,2α

3.6

3.9

1α,2β

2.8

1β,2α

13.1

1β,2β

H,H

70 Calc

Obs

1α,2α

6.4

5.9

2.8

1α,2β

11.7

13.2

13.4

1α,11

3.0

2.4

4.1

5.3

1β,2α

1.1

1.0

2α,3α

4.8

4.8

1β,2β

6.4

5.4

2α,3β

13.0

13.5

1β,11

3.7

4.0

2β,3α

2.0

2.1

2α,3

5.3

–

3β,3β

4.4

7.5

2β,3

11.4

12.7

6α,7

3.0

3.5

3,4

2.2

2.4

6β,7

12.3

12.7

4,5

1.0

1.0

7,8α

3.4

3.5

4,9

0.6

1.0

7,8β

12.3

12.5

5,11

1.2

1.3

b

7,11

5.8

5.8

8α,9

3.3

3.4

8α,9α

4.1

4.5

8β,9

2.8

2.9

8α,9β

2.8

2.6

9,10

3.4

3.1

8β,9α

13.1

13.2

10,11

6.9

5.9

8β,9β

3.9

4.1

a Measured b Not

at 300 MHz in CDCl3 observed due to signal overlapping

dichroism data and the anomalous dispersion effect detected in the X-ray diffraction analysis of brominated derivative 67. The structural information obtained for the lippifolianes was also useful in establishing a biosynthetic relationship with africanane derivatives, such as 68, also isolated from the same species [29].

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265

O 64 14

HO 2

3 4 15

11 10

1 6

7

9 8

5

O

13

HO

12

65

O 66

O

Br Br Br

O

O 67

68

A similar molecular modeling approach was applied to valeranone (69), a rearranged sesquiterpenoid with antispasmodic activity isolated from Valeriana officinalis [32] and to morelianone (70), a sesquiterpenoid with an interesting and intense woody odor [33]. This tricyclic structure was obtained by a Wagner-Meerwin rearrangement of the longipinane derivatives isolated from Stevia salicifolia. Table 1 shows the calculated vicinal coupling constants of 69 and 70 in comparison with the experimental values, and Fig. 1 displays the minimum energy molecular models of lippifolianone (65), valeranone (69), and morelianone (70).

Fig. 1 Minimum energy molecular models of lippifolianone (65), valeranone (69), and morelianone (70)

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J. C. Pardo-Novoa and C. M. Cerda-García-Rojas 14

14 1 2 3

1

9 8

10 4

5

7

2 11

O

15

12

69 (valeranone)

10 9 8

3

13 15

6

11

4

5 12

6

7

O 13

70 (morelianone)

The structure and conformation of the longipinene diester 71 isolated from Stevia eupatoria (Fig. 2) [34] was studied by a Monte Carlo search protocol [30] followed by DFT molecular modeling calculations at the B3LYP/6-31G* level of theory that revealed the ester residue orientations and the preferred conformation of the seven-membered ring. The results with respect to the sesquiterpene framework were validated by the comparison between calculated and experimental vicinal 1 H–1 H coupling constant analytical data [34]. The conformational analysis of 11αH-dihydrozaluzanin (72) was carried out by molecular dynamics to locate all possible conformations, followed by DFT calculations at the B3LYP/6-31G(d,p) level, to optimize the molecular geometry of the minimum energy structures [35]. This molecular modeling approach also allowed the calculation of the theoretical coupling constants, as well as the generation of the molecular electrostatic potential map to rationalize the susceptibility of electrophilic attack on the molecule [35]. The coupling constants, which were essential elements

Fig. 2 Inflorescence of Stevia eupatoria Willd. Photograph courtesy of Dr. J. Martín TorresValencia

New Techniques of Structure Elucidation for Sesquiterpenes

267

in the conformational analysis of 72, were also calculated by means of the KarplusCalculator method and by the MestRe-J program graphical tool for prediction of vicinal proton–proton 3 J H, H coupling constants [2]. O O

O

O OH O O

O O O 72 (11α H-dihydrozaluzanin)

71

A conformational study of cedranolides 73–82 was carried out by molecular modeling with the Complete Basis Set 4 Minimal Population Localization (CBS4M) method together with the comparison between experimental and calculated coupling constants [36]. It was found that in cedranolides 73, 75, 76, 78, 79, 81, and 82, the conformational equilibrium of the five-membered ring bearing the secondary methyl group is shifted toward a twisted-envelope conformation with the methyl group in a pseudo-equatorial orientation. In contrast, in cedranolides 74, 77, and 80, this five-membered ring is flipped and the methyl group mainly adopts a pseudoaxial orientation. The complete assignments of the 1 H NMR chemical shifts and experimental coupling constants were achieved by an iterative full spin analysis [36] employing the PERCH NMR software [37] (PERCH Solutions Ltd., Kuopio, Finland). R1

R2

O

R1

73 R1 = O, R2 = OH 74 R1 = H2, R2 = H

75 α-CH3 76 β-CH3

OAc OH

77 78 79 80

β-ΟAc, α-CH3 α-OAc, β-CH3 α-ΟAc, α-CH3 β-ΟAc, β-CH3

81 α-CH3, β-OH 82 β-CH3, α-OH

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2.3 Residual Dipolar Couplings The residual dipolar couplings (RDCs) technique is based on the property of gels such as poly(methyl methacrylate) to orient organic molecules when swollen in NMR tubes in a solvent as CDCl3 . The one-bond 13 C–1 H RDCs for a compound, as well as its degree of alignment, depend on the cross-link density and provide information about the orientations between internuclear vectors regardless of the distance between them. The utility of this method was exemplified with the sesquiterpene lactone ludartin (83) isolated from Stevia yaconensis var. subeglandulosa, which in combination with a 3 J coupling constant analysis, allowed determination of its relative configuration [38]. Application of the RDCs protocol was conclusive for the stereochemical analysis and the assignment of the diastereotopic hydrogen atoms of this bioactive sesquiterpene lactone [38].

O O O 83 (ludartin)

The structures of nine new eremophilanolides (84–92), isolated from the aerial parts of Senecio volckmannii var. volckmannii, were studied mainly by NMR spectroscopic techniques [39]. An interesting protocol named Computer-Assisted 3D Structure Elucidation (CASE-3D) was employed for the configurational and conformational analysis of several of these eremophilanolides, taking advantage of the information provided by the RDC results and DFT-calculated 1 H and 13 C chemical shifts. Application of the RDC method to 84 and its stereoisomer 93 illustrated some aspects about differential ordering in small molecules in the alignment medium [39].

New Techniques of Structure Elucidation for Sesquiterpenes

269

OH O

O

O

O

O

O

84

85

86

OH O

O

O O

O

OH

O

87

89 6α,7α-epoxy 90 6β,7β-epoxy

88 O

OH O

O O O

91 8β-hydroxy 92 8α-hydroxy

93

The soft coral Lemnalia flava collected near Xisha Island, China, afforded six new sesquiterpenoids (94−99), for which their structures were established by NMR spectroscopy, X-ray diffraction analysis, chemical transformations, and comparison with theoretical NMR data and DFT electronic circular dichroism calculations [40]. The RDCs analysis was employed to evaluate the relative configuration of 96, which was aligned in an oligopeptide phase (AAKLVFF) that was compatible with MeOH [40]. RDCs were measured using the [1 H–13 C]-CLIP-HSQC experiment [41] that allowed extraction of nine values with high accuracy. On the other hand, a molecular modeling search protocol followed by DFT optimization at the B3LYP/6-31G(d) level of theory of 96 and three of its stereoisomers revealed group structures to be analyzed according to NOE and RDC values [40]. The alignment tensor of each structure was calculated using the singular value decomposition method as implemented in the RDC module of the MSpin program [42]. The predicted RDCs were compared with the experimental ones providing the Q-factor as a quantitative evaluation for each stereoisomer, supporting the (4S*5S*6S*7S*11S*) configuration for 96 [40]. OH

O O

OH

O

O O

O O O

O

94

95

96 O OH O

NH

OH

O 97

O 98

99

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2.4 Advanced Mosher’s Methods The widely used Mosher’s method for the absolute configuration determination of chiral compounds [43, 44] has benefited with the introduction of two improvements that have been important in the structural elucidation of natural products and derivatives. These are (1) the implementation of this procedure for using directly in NMR tubes [45], which has been crucial when the amount of material is limited, and (2) the interpretation of the results in conjunction with molecular models, which have been fundamental tools to evaluate the three-dimensional arrangement of the ester derivatives, for a better understanding of the diamagnetic shielding effects. Thus, the aerial parts of Mikania thapsoides afforded six sesquiterpene lactones (100–105) with an unusual cis,cis-germacranolide skeleton and two melampolides (106 and 107), all of them bearing a trans C-8 lactone ring closure. Their structural elucidation was carried out using 1D and 2D NMR measurements and the absolute configuration of 100 was determined through the Mosher ester methodology in combination with a molecular model calculated with molecular mechanics [46]. The (R)- and (S)-α-methoxy-α-trifluoromethylphenylacetate (MTPA) esters of 100 were prepared to determine the absolute configuration of the carbon atom bearing the hydroxy group. The configuration at C-5 was deduced as (R) due to the high shielding experienced by H-8 (δ 1.34 ppm) of the (R)-MTPA ester as compared to the (S)-MTPA ester, which indicated that H-8 is strongly affected by the phenyl group of the MTPA moiety. The H-7 shielding (δ 0.21 ppm) was smaller than that of H-8, which was in agreement with the trans lactone ring closure [46]. OR1

14

8

2 7

3 4

O

O

10 9

1

5

O

12

11

O O

O

6 13

O OSen

OH

15

2

OR

100 R1 = Ac, R2 = Sen 101 R1 = Ac, R2 = i-Val 102 R1 = H, R2 = Sen

103

OAc

O O

O

O

O

OH

OSen

OR 104 R = Sen 105 R = i-Val

106 R = Sen 107 R = i-Val

O Sen =

(

O i-Val =

(

New Techniques of Structure Elucidation for Sesquiterpenes

271

Two furanoeremophilanes 108 and 109 together with twenty-one known constituents were isolated from Senecio chionophilus [47]. The structural assignment was carried out based on spectroscopic data and chemical transformation methods. The absolute configuration of 108 was determined by the Mosher ester methodology which was performed directly in NMR tubes using deuterated pyridine as the solvent. Compound 108 was treated with the (–)-(R)- and (+)-(S) α-methoxyα-(trifluoromethyl)phenylacetyl chlorides to obtain the (S)- and (R)-esters, respectively. The analysis of the differential chemical shift (δ H ) data showed positive changes for H-10 and H-15 and negative effects for H-2 and H-3. These effects indicated that the absolute configuration at C-1 of 108 was (S). According to a ROESY experiment in combination with the Mosher ester data, the absolute configurations at C-4, C-5, and C-10 were deduced as (S), (R), and (R), respectively, Thus, the structure of 108 was established as (1S,4S,5R,10R)-1-hydroxy-6-isobutyryloxy10H-9-oxofuranoeremophilane. The structure of 109 was elucidated by NMR data that also included a ROESY experiment. This compound showed mild activity against Mycobacterium tuberculosis [47]. O

HO 1

3

4

10 5 15

8

O

O 12

6

7

O

14

O

HO

9 2

11

O

13

O

108

O

109

The structure elucidation of the isomeric sesquiterpenes godotol A (110) and godotol B (111), isolated from Pluchea arabica, was carried out by analysis of the NMR data [48]. For 110, the connectivity was established using a HMBC plot, which showed correlations of H-5 with C-3 and C-13, and H-7 with C-5 and C-12. The connectivity of 111 was determined by HMBC correlations of H-5 with C-3, C10, C-14, and C-15, and H-11 with C-2, C-9, and C-12. The absolute configurations were established by the Mosher ester approach based on the δ H = (δ S – δ R ) values of the hydrogen atoms on both sides of the carbinol carbon atoms and with the help of molecular models. For godotol A (110), they were determined as C-2 (R) and C-9 (S) while for godotol B (111) they were assigned as C-3 (R) and C-8 (R). Both compounds showed weak antibacterial activity, inhibiting the growth of Staphylococcus aureus [48]. 9

10 12

11 7 6

OH 12 1

8

13 1

5

2 4

15

2

OH

3

HO

10

11

3

4

6 15

14

9 8

5

7

13

14

110 (godotol A)

111 (godotol B)

OH

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Three new sesquiterpenoids were obtained from the plant species Saussurea laniceps, two guaianolides (112) and (113), and one eudesmane derivative (114). Their structural elucidation was carried out by spectroscopic methods and the absolute configurations of 112 and 113 were determined by application of the Mosher’s method [49]. In the case of compound 112, the negative δ H values for H-15α (– 0.02 ppm) and H-15β (–0.03 ppm) and the positive δ values for H-2α (+0.02 ppm) and H-2β (+0.03 ppm) indicated that C-3 has the (R)-configuration. Similarly, for compound 113, the negative δ values for H-15α (–0.04 ppm) and H-15β (−0.04 ppm) and the positive δ values for H-2α (+0.03 ppm) and H-2β (+0.02 ppm) showed that C-3 has the (R)-configuration. Compound 114 displayed a substantial inhibition of the proliferation of murine T and B cells in vitro [49]. 14

HO

3 4

O

9

2

10

1 5

8

6

O

1'

O 2'

3'

OH

O

HO

7 4'

15

O

13

12 11

O

O

O

112

113 HO OH OH

114

The plant species Artemisia absinthium yielded a new sesquiterpene lactone named artabolide (115) that exhibited a significant inhibition of polar auxin transport in radish hypocotyls [50]. Its structure determination was carried out by NMR spectroscopy and X-ray diffraction analysis. This analysis in combination with NOESY data revealed that the 10-membered ring of 115 exists in a chair–chair conformation with the C-3 hydroxy group oriented in an axial position. The conformational information, supported by the X-ray analysis, was important for the interpretation of the Mosher method, which was used to establish the absolute configuration of artabolide 115 as (3R,4S,6R,7S) [50]. 14 1 3

HO

9 10

2

14 4

15

5

O

8 7

6

OH 7

8

OH 9

11

6

O

5

13

4

3

2

12

O 115 (artabolide)

12

11

1

10

15

13

116 (alyterinone)

The stems of Alyxia schlechteri collected in the Phu Wiang District of Thailand, afforded a germacrane sesquiterpene, which was named alyterinone (116) [51]. Its structure elucidation was achieved by 1D and 2D NMR spectroscopy including a NOESY experiment, which supported the orientation of the hydroxy groups at C-8

New Techniques of Structure Elucidation for Sesquiterpenes

273

and C-9 that exist in a trans configuration. Also, H-6α showed NOESY correlations with H-4α and H-12 of the α-isopropenyl group, while H-8 and Me-14 showed a correlation that indicated their axial β-orientation. Application of the Mosher methodology gave the corresponding monoesters at C-8, revealing that this chiral center was (S) and allowing the full assignment of the absolute configuration of alyterinone (116) [51]. A new hirsutane sesquiterpenoid with an uncommon variation in the hydrocarbon skeleton was isolated from the cultures of the basidiomycete fungus Lentinus cf. fasciatus [52]. This tricyclic compound, named lentinulactam (117), exhibited a conjugated bridged lactam as the distinctive feature. Its structure and absolute configuration were established by NMR spectroscopy in combination with Mosher ester analysis and calculation of a molecular mechanics model. The analysis of the absolute configuration indicated that lentinulactam (117) has the opposite configuration to that of related hirsutane derivatives [52]. HO

O NH OH

117 (lentinulactam)

The chemical study of the tropical rainforest basidiomycete Marasmiellus troyanus, isolated from spore fall of a mature basidiocarp found in Rio Palenque Forest Reserve, Ecuador, gave three new sesquiterpene lactones (118–120) with an unusual hydrocarbon framework related to cis-fused caryophyllane. Their structures were determined by detailed NMR spectroscopy and the absolute configuration of 118 was established by a single-crystal X-ray structural analysis in combination with a modified Mosher’s ester approach. Treatment of 118 with (S)- and (R)-methoxyphenylacetic acid (MPA) only yielded the monoesters at C-2, probably due to steric hindrance effects, revealing the (S) configuration at this chiral center [53]. 12 13

HO

HO

OH

2

O

HO

3 11 10

1

15 9

5 8

O

7

118

4

HO 6

O

OH

14

O 119

O

O 120

The marine sponge Dysidea septosa collected from Lingshui Bay, Hainan Province, China, gave five new sesquiterpenes, which were named lingshuiolides A (121) and B (122), lingshuiperoxide (123), isodysetherin (124), and spirolingshuiolide (125). Their structures were elucidated by 1D and 2D NMR spectroscopy by analogy with related known compounds. Molecular models of 123 and 125 were useful in visualizing the ROESY correlations that accounted for their relative configurations, while the absolute configuration of lingshuiolide B (122) was determined

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J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

by application of the modified Mosher’s methodology. It is interesting to note that spirolingshuiolide (125) was classified as the first case of a sesquiterpene with a rearranged drimane framework [54]. R1 R2

O

O

O O

O

O

3

R

OH

121 R1 = R3 = OH, R2 = H (lingshuiolide A) 122 R1 = H, R2 = R3 = OH (lingshuiolide B)

123 (lingshuiperoxide)

O

O

O

O

O O 124 (isodysetherin)

125 (spirolingshuiolide)

A chemical study on the gorgonian coral Menella woodin, collected from Vietnamese coastal areas, allowed two new sesquiterpene lactones of the guaiane-type to be obtained. Their structures, represented by 126 and 127, were elucidated by extensive spectroscopic analysis that included the comparison between experimental and DFT-calculated NMR spectra. Their absolute configurations were determined by comparison between experimental and calculated optical rotations and electronic circular dichroism spectra, using quantum-chemical methods with the timedependent DFT calculations at the B3LYP/6-311G(d,p) level of theory and using the polarizable continuum model for solvation effects. The stereochemical information was supplemented with application of the Mosher’s method on compound 126 that provided results consistent with those obtained by the quantum calculations [55]. HO

HO HO O

O O

O O

126

O 127

2.5 Use of Cryoprobes for NMR Measurements Microscale NMR techniques using cryoprobes have expanded the versatility of new methods for structure elucidation of complex natural compounds that are present in very small amounts in rare organisms. Microprobe NMR spectroscopy, microscale degradation and synthesis, are synergistic tools for the discovery of bioactive natural

New Techniques of Structure Elucidation for Sesquiterpenes

275

products from uncultured microbes, rare invertebrates, and environmental samples [56]. The chemical examination of the saprotrophic wood decay fungus Granulobasidium vellereum afforded the illudane sesquiterpenoids 128–135, for which their structure elucidation was achieved mainly by NMR spectroscopy taking advantage of a highly sensitive cryoprobe [57]. Molecular models of the new compounds were obtained with a molecular mechanics energy minimization procedure to rationalize the ROESY correlations used for the structure determination, while their absolute configurations were analyzed by application of the Mosher methodology. Compounds 128 and 131 were shown to be the enantiomers of the known compounds illudin M and dihydroilludin M, while 129 and 130 corresponded to diastereomers of illudin M and illudin S, and compounds 132 and 133 were two previously undescribed illudanes. The cytotoxicity of 128–131 and 133 was evaluated against two tumor cell lines, the Huh7 hepatocarcinoma cell line and the MT4 T-cell line, and showed that 128–131 have potent cytotoxic activity [57]. HO

HO

HO

HO

O

OH

O

128

OH

O 130

129

HO

HO

OH

OH

OH HO

OH

OH

131

132

133

OH

OH OH 134

135

The chemical study of the Cameroonian medicinal plant Alafia multiflora gave the aromadendrene sesquiterpenoids 136 and 137. Their structures were determined mainly by 1D and 2D NMR spectroscopy that included COSY, NOESY, HSQC, and HMBC experiments using an NMR instrument equipped with a cryoprobe [58]. Aromadendrene 136 displayed weak growth inhibitory activity against the B16F10 murine skin melanoma cell line [58].

276

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas O

HO

R 136 R = COOH 137 R = CHO

The phytochemical study of Centaurea ragusina collected from two wild habitats of Split, Croatia, yielded the sesquiterpene lactones 138–140. Compounds 139 and 140 were new and received the trivial names of ragusinin and hemistepsin, respectively. Their structure elucidation was carried out mainly by NMR using a spectrometer furnished with a cryoprobe [59]. Ragusinin (139) showed high specific cytotoxicity against the murine SCCVII and human HeLa and Caco-2 tumor cell lines, in being also not active against normal V79 fibroblasts. The biological activity of this natural compound was examined in detail [59]. O OH

HO

O

HO

HO

O O

O

O

O O

O

O HO

138

139

140

3 Techniques Based on Chiroptical Properties 3.1 Vibrational Circular Dichroism Remarkable advances in both experimental determination and theoretical calculation of vibrational circular dichroism (VCD) have positioned this technique as one of the most promising tools for determining the absolute configuration of organic compounds. These studies have been extremely useful in the field of structure elucidation of organic natural products and chiral drugs. The use of VCD spectroscopy has several advantages over other methodologies, such as the significant number of measurable transitions that provide multiple data for contrasting experimental and calculated values, which allow a better evaluation of the spectra. Each vibrational band in the infrared spectrum will have an associated sign, either positive or negative, which will depend on the chirality of the analyzed compound. The enantiomers of a given compound will show experimental VCD spectra that will be mirror images of each other and the absolute configuration of each enantiomer can be assigned by comparison with the calculated spectra.

New Techniques of Structure Elucidation for Sesquiterpenes

277

Several reviews have covered articles on vibrational circular dichroism of organic compounds up to 2015 [60–65]. Therefore, this section includes the relevant work on sesquiterpenes generated from 2015 to May 2020. The absolute configuration of inuloxin A (141), a phytotoxin produced by the plant species Inula viscosa, and seiricardine A (142), a phytotoxin isolated from the fungus Seiridium cardinale, was determined by chiroptical experimental measurements in comparison with theoretical calculations [66]. Vibrational CD spectroscopy was essential in this stereochemical study that was conducted in combination with optical rotatory dispersion and electronic circular dichroism. The conformational search of 141 was accomplished at the molecular mechanics level to give 15 conformers that were optimized at the DFT/B3LYP/6-31G and then at the DFT/B3LYP/TZVP levels of theory providing the three most populated conformers. Vibrational CD calculations were performed also using DFT methods at the M06-2X/TZVP level of theory with implicit solvation models IEF-PCM in chloroform and acetonitrile. The study of seiricardine A (142) was carried out on the 4-bromobenzoate derivative 143, which contains a convenient chromophore for the ECD spectrum, higher ORD and VCD values, and less intermolecular interactions. The absolute configuration of (+)inuloxin A (141) was established as 7R,8R,10S, while that of (–)-seiricardine A (142) and its derivative 143 was assigned as (1S,2R,3aS,4S,5R,7aS) [66]. 12

11 15 1

O 12

8

2

10

11

O

3

5

4

OH

3

9

RO

O

7

6

2

3a 7a

1

8

141 (inuloxin A)

5

13 6

7

10 13

14

4

9

142 R = H (seiricardine A) 143 R = 4-BrPhCO

The absolute configuration of the sesquiterpene lactone (–)-centratherin (144) was determined by a theoretical and experimental study that included VCD, optical rotatory dispersion, and electronic circular dichroism [67]. The VCD study of this furanoheliangolide was conducted according to a protocol that comprised a conformational search, carried out with the MMFF94s force field, followed by DFT geometry optimization and spectral calculation achieved at the B3LYP/aug-cc-pVDZ level of theory. The vibrational and electronic dissymmetry factors were used to evaluate the results and validate the absolute configuration of (–)-centratherin as (6R,7R,8S,10R,2 Z)-144 [67].

O

14 10

2

O 5

O

13

7 6

4 15

O

8

3

HO

O

9

1

11

O

O O

12

OH

O

144 ((—)-(6 R,7R,8S,10R,2Z)-centratherin)

145 (farinosin)

278

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

The absolute configuration of the biologically active sesquiterpene lactone farinosin (145) isolated from the plant species Encelia farinosa, collected near Hermosillo, Mexico, was studied with VCD spectroscopy [68]. The experimental conditions for the measurement involved the use of 100% atom-D CDCl3 solutions of 145 in BaF2 cells with an acquisition time of 20 h. The sample stability was monitored by 1 H NMR analysis immediately before and after the VCD measurements. The VCD calculations involved conformational searches by means of molecular mechanics force field calculations (MMFF94), followed by structural optimization of the more stable conformers and calculation of the VCD spectra. These tasks were carried out with DFT calculations using the functional B3PW91 and the DGDZVP basis set. The stereochemical study of this eudesmanolide (145) was reinforced by calculation of the Hooft X-ray parameters [68]. The volatile components present in the steam distillation oil of the roots of Vetiveria zizanioides, commonly known as vetiver oil, yielded three new sesquiterpenoids named vetiverianines A (146), B (147), and C (148) [69]. Their structures were determined by NMR analysis, X-ray crystallography, and VCD spectroscopy. The conformational search of the novel (4S,5S,6S,7S,10S)-146 was performed using the Monte Carlo protocol with the MMFF94S molecular mechanics force field to yield four conformers that were DFT geometry optimized at the B3PW91/DGDZVP2 level. The averaged calculated spectrum was compared with the experimental one, showing a strong agreement that validated the proposed absolute configuration. A similar protocol was applied to eremophilanes 147 and 148, which also gave good results, allowing the assignment of their absolute configuration as (4R,5S,7R)-7,11epoxy-α-vetivone and (4R,5S,7R,11R)-2-deoxo-7,11-epoxy-13-hydroxy-α-vetivone, respectively [69]. 16

1

14 9

O 15

2 3

4

11

O

8

10 5

7 6

12

2 3

OH 13

146 (vetiverianine A)

1

4

10

9

6 15

14

OH

8 7

5

11

13

O

O 12

147 (vetiverianine B)

148 (vetiverianine C)

A theoretical procedure to discern between correct and incorrect chemical structures of chiral compounds was developed on the basis of VCD spectroscopy and Raman optical activity [70]. The methodology involved the analysis of the spectral similarity overlap between experimental spectra and those calculated with quantumchemical theories that included the DFT/B3LYP/TZVP level of theory with the polarizable continuum model for solvent effects after Boltzmann weighted average spectra of all conformers. Raman optical activity tensors were calculated with the DFT at the B3LYP/aug-cc-pVDZ level with force constants calculated at the B3LYP/TZVP level also considering the solvent effects. The structures and conformations of the sesquiterpenoids aquatolide (149), caespitenone (150), and sporol (151), among other natural products, were studied by application of this useful methodology [70].

New Techniques of Structure Elucidation for Sesquiterpenes

279

O

O O

O O

O O

OH

O 149 (aquatolide)

150 (caespitenone)

151 (sporol)

Aquatolide 149 was isolated for the first time from Asteriscus aquaticus [71], and years later, its structure was revised by quantum-chemical NMR calculations, NMR analysis, and X-ray crystallography [72]. This molecule was taken as a relevant example to emphasize the importance of archiving the 1 H NMR FID files for structural analysis and dereplication [73]. Five new sesquiterpenoids of the botryane group (152–156) were obtained from the ethyl acetate extracts of the endophytic fungus Nemania bipapillata that was isolated from the marine red alga Asparagopsis taxiformis found in the Falkenbergia stage [74]. It is worth mentioning that compounds 155 and 156 are in fact nor-sesquiterpenoids. The five structures were determined by NMR and mass spectrometry, while the absolute configuration of 152 and 156 were established by VCD spectroscopy. The high level of concordance between the experimental and DFTcalculated IR and VCD spectra at the B3LYP/6-31G(d) level of theory with the polarizable continuum model in methanol permitted the assignment of the absolute configuration of 152 as (2R,4S,5R,8S) and 156 as (2R,4S,8S). The absolute configurations of 153–155 were accomplished by comparison of their experimental and calculated electronic circular dichroism spectra. All fungal metabolites were tested as cholinesterase inhibitors, revealing 153–156 to be more active toward human acetylcholinesterase than human butyrylcholinesterase. The most active compound was 155 with 27.7% inhibition at 100 μM against acetylcholinesterase [74]. 10 11

2 3

1

4

O 14 9 5

O

O

15

OH

OH

OH

8 7 6 13

OH OH 12

OH OH

OH

153

152 13

O 10

1

2 3

O

14

OH

OH

9

8

5 6

4

154

7 12

OH

11

155

OH 156

The new eudesmane derivative 157, bearing a cyclic carbonate moiety, was isolated from marine filamentous cyanobacterial mats found in the waters of the marine mammal zoological park in the Florida Keys, USA. Grazing on the mats by captive bottlenose dolphins caused apparent ingestion-related intoxication, which prompted the researchers to undertake a chemical study of these ecosystems.

280

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Sequencing of 16S rDNA disclosed that the mats contained closely related Oscillatoriacean species, including an unknown cyanobacterium belonging to the genus Neolyngbya. The structure of 157 was determined by NMR spectroscopy, singlecrystal X-ray diffraction, and mass spectrometry. Its absolute configuration was established by comparison of experimental and DFT calculated VCD spectra at the B3LYP/6-31G(d) level of theory, from which their agreement allowed the assignment of 157 as the (4S,5R,6R,7S,10S)-eudesman-(4,6)-cyclocarbonate. Compound 157 was evaluated in a zebrafish embryo/larval model showing neurotoxic effects qualitatively similar to those observed for brevetoxin-2 and consistent with neurobehavioral impairment observed in the dolphins [75]. O 12 11 13

O 7

6

O 5

4

10

8

14 3 2

1

9 15

157 ((4S,5R,6R,7S,10S)-eudesman(4,6)-cyclocarbonate)

3.2 Electronic Circular Dichroism Electronic circular dichroism (ECD) has been one of the most exploited methodologies for the absolute configuration determination of organic natural products because only small amounts of materials are required for analysis. Besides its wide application taking into consideration the empirical rules based on numerous examples, the incorporation of molecular modeling protocols, inclusive of the calculation of time-dependent DFT ECD spectra has enhanced the potential of this technique in the structure elucidation of new molecules including conformationally flexible and restrained sesquiterpenoids [76]. A selection of the most illustrative publications on this field is summarized in this section.

3.2.1

Time-Dependent DFT Calculations

Two bisabolene-derived sesquiterpenoids, named artaboterpenoids A (158) and B (159 and 160), were isolated from the roots of Artabotrys hexapetalus, a medicinal plant of the Annonaceae family found in China. Bisabolene 158 was assigned a novel hydrocarbon skeleton formed by a C-2–C-10 linkage, while artaboterpenoid B (159 and 160) was in fact obtained as a pair of enantiomers representing the first example of a 1,2-seco-bisabolene-type sesquiterpene lactone [77]. The absolute configurations of 158–160 were determined by comparison of experimental and DFT calculated electronic circular dichroism (ECD) analyses. The ECD spectrum of 158

New Techniques of Structure Elucidation for Sesquiterpenes

281

showed an intense negative Cotton effect at 208 nm (ε = − 18.04 mol–1 dm3 cm−1 ), which resulted from the π x → π x * electronic transition of the 1(6) double bond of the cyclohexene ring, rather than the 11 double bond. The ECD spectrum of 158 was calculated for the four possible enantiomers of this sesquiterpenoid using the B3LYP/6-31G(d) level of theory. The experimental ECD spectrum of 158 showed a good agreement with the calculated spectrum of the (2R,4S,8S,10R)-isomer considering the sign and the intensity of the analyzed transition [77]. The mixture of artaboterpenoids B (159 and 160) was resolved on a chiral column and their absolute configuration was established as (−)-(8R)-159 and (+)-(8S)-160 by application of helicity rules [77]. Enantiomer (−)-(8R)-159 displayed cytotoxic effects toward the HCT-116, HepG2, A2780, NCI-H1650, and BGC-823 cell lines with IC 50 values between 1.38–8.19 μM. 9

8

4 5

10

3

6

2

O

14

12 13 11

10

O

9

HO

8 2

OH 15

12 11

14

O

13

4 3

5

O

6

1

O

15

O

1 7

7

158 ((2R,4S,8S,10R)artaboterpenoid A)

160 ((+)-(8S)artaboterpenoid B)

159 ((—)-(8R)artaboterpenoid B)

Two sesquiterpenoid glucosides of the guaiane-type (161 and 162) were isolated from the fruits of Gardenia jasminoides, a medicinal plant of the Rubiaceae family widely used in China [78]. The methodology for the determination of the absolute configuration of 161 was done by means of an ECD analysis. Its ECD spectrum showed a positive Cotton effect at 313.0 nm and a negative Cotton effect at 232.8 nm, suggesting that C-1 has the (R)-configuration. The DFT ECD calculation of a model compound without the sugar moiety was performed at the B3LYP/6311++G(2d,p) level. The shapes of the computed and experimental spectra were almost identical, indicating that the structure and absolute configuration of 161 corresponded to (1R,7R,10S)-11-O-β-d-glucopyranosyl-4-guaien-3-one. The structure of 162 was determined by comparison of the NOESY and ECD spectroscopic data with those of 161 [78]. 14 10

2

O

3

9

1 4

O

8

5 6

7

11

O

15 12

HO

13

1'

161

O HO

6'

5' 2'

3'

HO

4'

OH OH

HO

O

O

OH OH

HO 162

The chemical study of the fruits of Illicium simonsii, a medicinal bush of the Illiciaceae family, has revealed the presence of a unique pentacyclic structure with a 5/5/5/5/5 caged 2-oxatricyclononane ring system fused to a five-membered carbocyclic ring and a five-membered lactone [79]. This novel compound received the

282

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

trivial name of illisimonin A (163) and the structure was deduced by extensive NMR analysis. Its absolute configuration was established by comparison between experimental and theoretical ECD analytical data. The experimental spectrum of 163 showed a strong negative Cotton effect at 216 nm for the n → π* transition of the lactone ring moiety. Calculation of the theoretical ECD spectrum for the two possible enantiomers of this compound was carried out using time-dependent DFT at the B3LYP/6-31G(d) level of theory. The similarity of both spectra allowed assignment of the (1R,4R,5R,6R,7S,9S,10S) configuration to 163. A biosynthesis pathway for the formation of 163 from bisabolene through cedrane and seco-prezizaane intermediates was proposed. Interestingly, illisimonin A (163) demonstrated a neuroprotective effect against cellular injury induced by oxygen–glucose deprivation in SHSY5Y cells with an EC 50 value of 27.7 μM [79]. O HO 15 2

HO

12

11

10

9

8

1

7

O 4 3

OH

6

13

O

5 14

163 (illisimonin)

Four novel germacranolide sesquiterpene lactones named trichospirolides A– D (164–167), were isolated from Trichospira verticillata, an unexplored species collected in Alajuela Province, Costa Rica. Their structures were elucidated taking into consideration extensive NMR spectroscopic analysis and molecular modeling, including analysis of internuclear distances and coupling constants for all possible diastereomers of each compound. A detailed ECD study was undertaken for the determination of absolute configuration of 164–167 [80] on the basis of the comparison between experimental and calculated ECD and UV spectra. The excitation energies, rotatory strengths, and oscillator strengths for each transition were calculated for the first forty electronic states at the TDDFT/B3LYP/aug-cc-pVDZ level of theory including the PCM with methanol as the solvent. The study was conducted to find new antiplasmodial compounds and revealed that trichospirolide A (164) exhibited the highest activity among the four compounds against the growth of Plasmodium falciparum Dd2 strain (IC 50 value of 1.5 μM) [80].

New Techniques of Structure Elucidation for Sesquiterpenes O

O

O

O

O

H

O

H O

O

283

O

HO

165

164 O

O

O O

O

O

HO

O O

O

O

O

O

O

O 166

O

O

O

O

O

O

167

A chemical study of the leaves of Piptocoma rufescens collected in the Dominican Republic afforded six new germacranolide-type sesquiterpene lactones (168–173), together with eight known related compounds. The structure elucidation of the new lactones was determined by extensive NMR measurements, together with the singlecrystal X-ray diffraction analysis of 170 [81]. The absolute configuration of 168 was established by application of the modified Mosher ester NMR method carried out in NMR tubes, while those of the whole series of compounds were achieved by a detailed analysis of their ECD spectra. The negative Cotton effects at 232 and 264 nm for the π → π* and n → π* transitions of the lactone ring, measured in the ECD spectrum of 168, was in agreement with the (7R) absolute configuration and with previous studies of related sesquiterpene lactones, while the negative Cotton effects at 223 and 272 nm in the ECD spectrum of 170 also indicated an α-configuration for H-7. All compounds displayed a high cytotoxicity toward the HT29 human colon cancer cell line and compound 172 exhibited also NF-κB (p65) inhibitory activity [81].

284

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas O

O O

O

O

HO

O

HO

O

O O

O O

O

168

169 O

O

O

O O

O

O

O O

O O

O O

O

170

171 O

O O

O O

O

OH O

O

O

O O

O O

O

172

173

The methodology for the configurational analysis of sesquiterpenes 174–213 followed a similar strategy to the examples describes above. The sequence involved measurement of the ECD spectrum in a selected solvent, followed by calculation of the ECD curves for the most probable enantiomer, usually employing a timedependent DFT method at a confidence level of theory, and comparison between the curves. It is important to mention that if an analyzed structure has several conformers, the calculated spectra of each conformer will require Boltzmann-averaging to obtain the ECD calculated curve. If this spectrum is comparable with its experimental counterpart, the absolute configuration of the studied compound can be assigned according to the configuration of the calculated molecular model. Table 2 summarizes illustrative works that were carried out using this methodology.

Artemisia rupestris Artemisia rupestris Elephantopus scaber

Artepestrin D (189)

Artepestrin F (190)

Elescabertopin A (191)

Petasites hybridus

Artemisia rupestris

Artepestrin B (188)

8βH-Emophilanolide (194)

Artemisia rupestris

Artepestrin A (187)

Elephantopus scaber

Artemisia annua

Hydroperoxyguaienone 186

Petasites hybridus

Trichothecium crotocinigenum

Trichothecrotocin A (185)

8β-Hydroxyeremophilanolide (193)

Tanacetum sonbolii

Germacranolide 184

Elescabertopin A (192)

Illicium henryi

Illihenazulene A (183)

Curcujinone A (179)

Abies holophylla

Curcuma wenyujin

(+)-seco-Tanapartholide A (178)

Abiesesquine A (182)

Artemisia argyi

(–)-seco-Tanapartholide B (177)

Curcuma wenyujin

Artemisia argyi

Canin (176)

Carthamus oxycantha

Artemisia argyi

Peribysin Q (175)

1β,10β-Epoxydesacetoxymatricarin (181)

Periconia macrospinosa

Peribysin O (174)

Curcujinone B (180)

Source Periconia macrospinosa

Compound

Table 2 ECD analysis of sesquiterpenes using time-dependent DFT Level of theory

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31G(d)

mPW1PW91/6-311G(d)

B3LYP/6-31G(d,p)

B3LYP/6-31+G(d)

B3LYP/6-31G(d,p)

B3LYP/TZVP

B3LYP/6-311++G(d,p)

B3LYP/6-311++G(d,p)

B3LYP/6-31G(d,p)

B3LYP/6-31G(d,p)

B3LYP/6-31G(d,p)

BHLYP/def2-TZVP

BHLYP/def2-TZVP

Ref.

(continued)

[93]

[93]

[92]

[92]

[91]

[91]

[91]

[91]

[90]

[89]

[88]

[87]

[86]

[85]

[84]

[84]

[83]

[83]

[83]

[82]

[82]

New Techniques of Structure Elucidation for Sesquiterpenes 285

Source Petasites hybridus Petasites hybridus Petasites hybridus Petasites hybridus Petasites hybridus Petasites hybridus Petasites hybridus Petasites hybridus Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita Artemisia vestita

Compound

2-Angeloyloxyeremophil-7(11)-en-12,8-olide (195)

2-(3-Methylsulfanylprop-2-enoyl)oxyeremophil-7(11)-en-12,8-olide (196)

2-Methacryloxyeremophil-7(11)-en-12,8-olide (197)

2-Angeloyloxy-9-hydroxyeremophil-7(11)-en-12,8-olide (198)

2-Angeloyloxyeremophil-7(11)-en-12,8-olide (199)

2-Methacryloxyeremophil-7(11)-en-12,8-olide (200)

Isopetasin (201)

Iso-S-petasin (202)

Arvestolide D (203)

Arvestolide E (204)

Arvestolide F (205)

Arvestolide G (206)

Arvestolide H (207)

Arvestolide I (208)

Arvestonate A (209)

Arvestonate B (210)

Arvestonol (211)

Arvestolide J (212)

Arvestonate C (213)

Table 2 (continued) Level of theory

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31+G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

B3LYP/6-31G(d)

Ref.

[94]

[94]

[94]

[94]

[94]

[94]

[94]

[94]

[94]

[94]

[94]

[93]

[93]

[93]

[93]

[93]

[93]

[93]

[93]

286 J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

New Techniques of Structure Elucidation for Sesquiterpenes OH

287 OH

OH O

O OH

O 175

174

O

O O

HO O

O

O

O

O

O

O

O

OH

O 176

OH

177

178 O

O

OH

O O

O

O

O

O

O 181

180

179 O

O O

OH OH

O

HO

O

O O 183

182

184

OH

HOO

O

O

OH

O

O

HO

185

OH

186 O

O

OH OH O

O HO

HO

O 187

O 188 O OH

O

O HO

O 189

OH

HO

O O 190

288

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas O

O

O

O

OH O

OH O

O O

O

O O

O OH

O

O OH

O

191

192

O

O

O

O

O

O

O

O O

O

R

O

O O

196

195

193 R = OH 194 R = H

O

O O

O

S

O

O

O O

O

O

OH 197

198

199

O

O

O O

O

O

O

O

200

O

O

201

202

O OAc

O O

OH

HO

O

O

O O O

O 203

O

O

HO

O

204

205

O OAc

OAc

O

HO HO

O O 206

O

O

O O

O 207

208

S

New Techniques of Structure Elucidation for Sesquiterpenes

289

OAc

OH

OH O

O O

OH OH

O

OH OH

209

O

OH

210

211 O

O

O O OH

O 212

3.2.2

O

O

O

OH

O 213

The Exciton Chirality Method and Molecular Modeling

This section considers those examples that have used the exciton chirality method [95, 96] in combination with molecular modeling of sesquiterpenes using timedependent DFT calculations. It should be mentioned that the specific alignment of the chromophores involved in the transitions can be estimated and visualized in the DFT optimized molecular models for each calculated structure in a highly efficient manner. A phytochemical study of the rhizomes of Acorus tatarinowii, a Chinese medicinal plant, gave four novel acorane sesquiterpenes (214–217) together with two other new sesquiterpenes. Their structures were elucidated by extensive NMR spectroscopic analysis together with X-ray crystallography, while their absolute configurations were determined by a combined method that included calculation of the Flack parameter for the single-crystal X-ray analysis of 214 and the exciton chirality method applied to acotatarone B (217). The ECD spectrum showed a split curve with a negative band at 221 nm and a positive band at 246 nm associated with the exciton coupling between the two α,β-unsaturated carbonyl chromophores, indicating their absolute configuration of the spiro carbon at C-5 to be (R). The concordance between the experimental ECD spectrum of 217 with that calculated at the DFT B3LYP/631G(d) level of theory in methanol confirmed the absolute configuration of this compound as (1S,5R) [97]. It is important to remark that ECD calculated spectrum arise from Boltzmann averaging of individual curves of each low energy conformer. The anti-Alzheimer effects of all compounds were estimated by means of acetylcholinesterase and β-site amyloid precursor protein cleaving enzyme 1 inhibition assays [97].

290

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas 15

3

10 9 14

5

8

O 2

4

7

O

1 6 11

O

O

OR 13

12

O

214 R = H 215 R = Ac

O

216

217

A phytochemical study of Celastrus orbiculatus, an insecticidal plant of the Celastraceae family, yielded the dihydroagarofuran derivative 218, named celaspene A, together with another eight new related compounds [98]. Their structures, bearing multiple chiral centers, were elucidated based on extensive NMR spectroscopic data. The systematic conformational search of celaspene A (218), followed by structure optimization and time-dependent DFT ECD calculations, gave its calculated ECD spectrum, which was in agreement with the experimental curve that showed a characteristic Davidoff-type split with a first negative Cotton effect at 239 nm and a second positive effect at 222 nm due to the couplings of the two benzoyloxy groups, thus establishing the absolute configuration of 218 as (1S,4R,5S,6R,7R,8R,9S,10S) [98].

BzO

OAc OAc

BzO

O

OR

O

OAc

O

218 (celaspene)

219 R = H (frullanic acid) 220 R = CH3

Chemical examination of the Chinese liverwort Frullania serrata yielded two new cadinane-type sesquiterpenes, named frullanic acid (219) and its methyl ester (220). Their structures were elucidated by detailed analysis of spectroscopic data including 1D and 2D NMR and high-resolution mass spectrometry. The absolute configuration of frullanic acid (219) was determined by comparison of its experimental and calculated ECD spectra, with the latter obtained using time-dependent DFT methods and the exciton chirality method [99]. The ECD spectrum of 219 showed a negative chirality resulting from the exciton coupling between the α,β-unsaturated ketone at 238 nm, ε = − 5.90 mol−1 dm3 cm−1 , for the π → π * transition and the α,βunsaturated carboxylic acid group at 215 nm, ε = + 7.32 mol−1 dm3 cm−1 , for the π → π *, implying that the transition dipole moments of the two chromophores were oriented in a counterclockwise arrangement. These observations were also supported by the time-dependent DFT ECD calculated spectrum and the molecular models involved [99]. Sarcanolides A (221) and B (222), two novel lindenane-type sesquiterpenoid dimers, were isolated from the plant species Sarcandra hainanensis, which has been used in Chinese folk medicine to treat inflammation. Their structures, containing a peculiar nonacyclic ring system, were determined by NMR spectroscopy, while the absolute configurations were analyzed by EDC time-dependent DFT calculations

New Techniques of Structure Elucidation for Sesquiterpenes

291

at the PBEPBE/6-31G* level of theory in gas phase and in methanol [100]. The calculated ECD spectra of sarcanolides A (221) and B (222) closely matched their experimental ECD curves and the molecular models of both structures allowed the interpretation of the exciton couplings, which arose from the interaction of the α,βunsaturated γ -lactone and the α,β-unsaturated ketone chromophores [100]. O O

O O

O

O O

HO OH

O OH

O

O

O

O

HO OH

O OH

O

OH 221 (sarcanolide A)

222 (sarcanolide B)

4 Techniques Based on X-Ray Diffraction Single-crystal X-ray crystallography is one of the most powerful techniques for the elucidation of the three-dimensional molecular structures that has been extremely useful for the field of natural products. While most advances in X-ray diffraction analysis have concentrated on the improvement of speed and accuracy of data collection, some developments have focused on the generation of enhanced tools for the determination of the absolute configuration of organic compounds. These approaches have been valuable for natural products that crystallize in non-centrosymmetric space groups. The calculation of the Flack parameter is based on the comparison obtained from a complete derivation of a Bijvoet intensity ratio by way of mean-square Friedel differences [101], while the Hooft protocol includes probabilistic methodology for the determination of the absolute structure of an enantiopure compound and is also based on Bijvoet-pair intensity differences [102]. Therefore, in both cases the collection of a larger data set is required to include pairs of redundant reflections related to each other by inversion through the origin. Furthermore, a reliable absolute configuration assignment can be made by using Cu Kα radiation. In both methods, if the analyzed structure is solved in the correct absolute configuration, either the Flack and/or the Hooft parameters will be near zero. On the contrary, if the structure is being solved in the opposite absolute configuration, both parameters will be increased up to the unit. Inversion of the model structure in the crystallographic software can be applied to test this attribute. This section describes the structure elucidation of new sesquiterpenoids using X-ray crystallography for which their absolute configuration determination was achieved by calculating the Flack and/or Hooft parameters.

292

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

4.1 Calculation of the Flack Parameter Two new sesquiterpenoids with a unique structure, named dendrowardols A (223) and B (224), were isolated from the stems of Dendrobium wardianum, a species of the Orchidaceae family that mainly grows in the south of China and some Southeast Asian countries, such as Myanmar, Bangladesh, and Thailand. Their structures were established based on extensive spectroscopic analysis including the 1D and 2D NMR data [103]. The absolute configuration of 223 was assigned as (1S,4R,5R,6R,7S,10R,11S) by X-ray diffraction analysis using the anomalous scattering of Cu Kα radiation that allowed the calculation of the Flack parameter with a significant value of 0.09(19). The relative configuration of the diastereomer 224 was determined by single-crystal X-ray diffraction analysis that was carried out using Mo Kα radiation, thus limiting the Flack analysis. Dendrowardol B (224) was evaluated by an MTT assay showed a promoting effect on the proliferation of human lens epithelial cells induced by d-galactose [103]. OH

OH

14 2

OH 10

9

6

7

1

3

OH

8

5 4

11

12

13

OH

OH 224 (dendrowardol B)

223 (dendrowardol A)

Two new plant metabolites of the aristolane-type sesquiterpenoid, designated as nardoaristolone A (225) and B (226), were isolated from the underground parts of Nardostachys chinensis, a medicinal species of the family Valerianaceae that generally grows in the Himalayan mountains [104]. Nardoaristolone A (225) contains a chalcone derivative, while nardoaristolone B (226) possesses a nor-aristolane hydrocarbon skeleton with an unusual 3/5/6 tricyclic ring system. A biogenetic pathway was proposed to explain the formation of both metabolites, for which their structures were elucidated by spectroscopic measurements. The absolute configurations were established by single-crystal X-ray diffraction analysis as (1R,4R,5R,6S,7R,10S)-225 and (4R,5R,6S,7R)-226 with Flack parameters of 0.0(3) and −0.03(19), respectively [104]. OH O 6''

7'

9'

5''

1'' 2''

O

4''

10

O

4

O

10

7

11

14 15

12 6

4 12

14

9

2 3

7 6

O

1

O

5

8

5

3

4' 5'

9 1

2

3'

6'

OO

3''

2' 1'

8'

11 13

15 13

225 (nardoaristolone A)

226 (nardoaristolone B)

New Techniques of Structure Elucidation for Sesquiterpenes

293

Three new dihydro-β-agarofurans named bilocularins A–C (227–229) were isolated from the dichloromethane extracts of the leaves of the Australian rainforest tree Maytenus bilocularis. Their structural elucidation was carried out through the analysis of NMR and mass spectrometric data, while the relative configuration was proposed from ROESY experiments. The absolute configurations of bilocularins A (227) and B (228) were determined as (1S,4R,5S,6R,7R,8R,9S,10S) and (1S,4R,5S,6R,7R,9S,10S), respectively, using the anomalous dispersion effects measured in their single-crystal X-ray diffraction analyses [105]. The Flack parameter of 5 was 0.02(6), by considering of a total of 1955 Bijvoet pairs, while that for 6 was 0.05(6) with a total of 1825 Bijvoet pairs. The absolute configuration of bilocularin C (229) was assigned as (1S,4R,5S,6R,7S,8R,9S,10S) based on biosynthesis considerations and comparison of chiroptical data. Bilocularins A (227) and B (228) each displayed a biological effect comparable to the drug efflux pump inhibitor verapamil in reversing drug resistance of a the CEM/VCR R human leukemia cell line [105]. OAc OBz

AcO 15 1

OAc OBz

OR

9 5

3

AcO 2

O

7 12

O

O

11

OR1

14

13

227 R = Ac, R = H (bilocularin A) 229 R1 = H, R2 = Ac (bilocularin C) 1

2

OAc 228 (bilocularin B)

The chemical examination of extracts of Eupatorium chinense, a shrub of the Asteraceae family collected at Changyang, People’s Republic of China, gave ten new germacrane-type sesquiterpene lactones (230–239) [106]. The absolute configuration of compound 230, named eupachinsin, was determined as (2R,3R,6R,7R,8R) by single-crystal X-ray diffraction analysis and the calculation of the Flack parameter 0.07(7) determined with 1701 quotients. The relative stereochemistry for compounds 231–239 was determined by the similarity of the ROESY correlations and coupling constants as compared to those of 230. The ECD spectra of 230–239 were found to be very similar, showing a weak positive Cotton effect around 250 nm and an intense negative effect around 210 nm, mainly attributed to the α,β-unsaturated lactone chromophore containing the exocyclic double bond. The resemblance observed in the ECD spectra of these compounds indicated that compounds 231–239 have the same absolute configuration at C-6 and C-7 as compound 230. Compounds 232 and 233 showed cytotoxicity against the MDA-MB-231 human breast cancer cell line, with IC 50 values of 0.8 and 3.4 μM, respectively, while 232–234 showed cytotoxic activity against a human hepatocellular carcinoma cell line (HepG2), with IC 50 values ranging from 3.6 to 7.6 μM [106].

294

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas HO O

1'

8

O

1

RO

10 3

14

AcO 15

2'

O 12

11

O

O O

4' 13

6

HO

5'

AcO O

R

O

230 R = H 231 R = Ac

232 R = H 233 R = OH

HO

R2

O

O O

AcO

O AcO

O O

O

R1O

O

235 R1 = H, R2 = H 236 R1 = Ac, R2 = OH

234

HO OH

HO

O

OH

O

O AcO

AcO R

O

O

O O 237 R = H 238 R = OH

O 239

Three new nor-cinalbicane (240–242) and one eremophilene (243) sesquiterpenoid were isolated from the stems of Jasminum officinale collected from Nyingchi County, Tibet Autonomous Region, China. The structures of jasminols A (240), G (241), H (242), and B (243) were elucidated on the basis of NMR and mass spectrometric analysis. The absolute configurations of 240–242 were determined as (4S) and that for 243 as (4S,5R) by single-crystal X-ray diffraction using the anomalous dispersion of Cu Kα radiation. The reported values for the Flack parameters were − 0.02 for 240, 0.00 for 242, and 0.03 for 243. The four compounds (240–243) showed moderate anti-inflammatory activity, as evaluated by means of their inhibition of lipopolysaccharide-induced nitric oxide production in RAW264.7 macrophage cells [107].

New Techniques of Structure Elucidation for Sesquiterpenes O 1 2 3 4 15

OH

295 O

9

10

8 7

5

12

OH

OH

240 (jasminol A)

241 (jasminol G)

6 14

11 13

O

OH

OH

O

O 242 (jasminol H)

243 (jasminol B)

Four new cadinane-type sesquiterpenoids (244–247) were isolated from the aerial parts of Heterotheca inuloides, which are used commonly in Mexican traditional medicine. Their structures were determined by extensive NMR spectroscopic analysis and confirmed by X-ray crystallographic studies of 244–246 [108]. The absolute configurations of metabolites 245–247 were assigned by calculation of the Flack parameters using anomalous X-ray scattering from the oxygen atoms and by comparison between experimental and calculated ECD spectra. The antiinflammatory potential of the four cadinane derivatives was studied using the 12-Otetradecanoylphorbol-13-acetate-induced mouse ear edema model. Cadinane 244 showed a moderate anti-inflammatory activity of 43.14 ± 8.09% at a dose of 228 μg/ear [108]. O HO HOOC 244 (cadinane)

HOOC

H 245

HOOC

246

247

Chemical examination of the medicinal plant Salvia scapiformis, which has been used for long time for the treatment of colds, bronchitis, tuberculosis, and traumatic hemorrhage in southwestern China, led to three new germacrane derivatives named glechomanamides A–C (248–250), bearing an unusual 8,11 -7,12-lactam functionality, together with two pairs of 7,12-hemiketal sesquiterpenoid epimers (251/253 and 252/254) [109]. Their structures were determined by extensive spectroscopic methods and verified by single-crystal X-ray diffraction analysis of 248–252 and 254. All crystallographic data were collected using Cu Kα radiation, which permitted

296

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

the calculation of the Flack parameters, and therefore the assignment of the absolute configuration for all compounds. It is relevant to comment that, when dissolved, the crystals of hemiacetal esters experienced a tautomeric equilibrium inverting the chiral center at C-7. Glechomanamide B (249) showed antiangiogenic activity by suppression of vascular endothelial growth factor and a significant suppression of mRNA expression associated with glycolysis and angiogenesis biomarkers, both effects in human umbilical vascular endothelial cells [109]. 2 3 4

O 5 15

O 1 6

10 14 7

9

O

R1 11

R1

O

O

8

O

R1

13

R2 N 12 H O

R2 O

R2 O O

O

248 R1 = OMeBu, R2 = OH (glechomanamide A) 249 R1 = OTig, R2 = OH (glechomanamide B) 250 R1 = OTig, R2 = OMe (glechomanamide C)

251 R1 = OMeBu, R2 = OH 252 R1 = OTig, R2 = OH

OMeBu = ( O

253 R1 = OMeBu, R2 = OH 254 R1 = OTig, R2 = OH

OTig = O

( O O

A new sesquiterpenoid dimer named chloraserrtone A (255) was isolated from Chloranthus serratus, a medicinal plant of the family Chloranthaceae collected from Wenzhou, People’s Republic of China. The structure of this sesquiterpenoid dimer, formed by two lindenane monomers was elucidated considering NMR, mass spectrometry and X-ray diffraction data [110]. This was the first report of a sesquiterpene dimer with a unique carbocyclic system linked by C-15–C-15 , C-4–C-6 , and C-6–C-11 bonds to form two six-membered rings between the monomeric units. The absolute configuration of 255 was determined as (1R,3S,4S,5S,6R,9R,10R,1 R,3 S,9 R,10 S,11 R), taking into account the Flack parameter that gave a satisfactory value of −0.07(15) [110]. 14

OH 9

1

2

10 15 3 4 15' 4'

6'

5'

3'

COOMe 7 12 6 OH 11' 11 13 COOMe 7'

10' 2'

1'

9' 14'

O 8

5

13' 8'

12'

O

OH

255 (chloraserrtone A)

The sesquiterpene derivative nicotabin A (256) was isolated from the leaves of the widely distributed plant Nicotiana tabacum. An investigation of the chemical constituents of the leaves of this species, obtained from Yunnan Province in China and carried out by a dereplication procedure, resulted in the isolation of this novel compound, which is formed presumably by a biosynthetic combination of a sesquiterpenoid with a C6 unit probably related to citric acid. Its structure elucidation was achieved by spectroscopic methods and single-crystal X-ray diffraction analysis

New Techniques of Structure Elucidation for Sesquiterpenes

297

using Cu Kα radiation. The absolute configuration of 256 was confirmed by calculation of the Flack parameter with a value of 0.05(5). This compound possesses a new hydrocarbon skeleton containing a 5/6/5/5/5 highly oxygenated ring system fused with the spirocyclic centers at C-2 and C-5 [111]. O

OH

19 14

O

3 4

13

2

6 11

18

10 12

8

1

OH

OH

20

O

17

5

7

21

O

16

9 15

256 (nicotabin A)

The rhizomes and roots of the medicinal plant Nardostachys chinensis yielded nardochinoids A–C (257–259), three new rare sesquiterpenoid dimers [112]. Nardochinoid A (257) represented an unprecedented pattern of dimerization composed of two types of sesquiterpenes defined by the presence of a new fused 3,8-dioxatricyclo[7.2.1.01,6 ]dodecane-11-one ring system, while nardochinoid B (258), a unique sesquiterpene dimer with a pyridine core, was the first report of a nitrogen-containing nornardosinane-aristolane sesquiterpene conjugate. Nardochinoid C (258) was the first sesquiterpene dimer fusing a nornardosinane sesquiterpene with a nardosinane sesquiterpene through a five-membered Oheterocyclic ring to generate a 6/6/5/6/6 polycyclic ring system. Their structural elucidation was determined by NMR spectroscopic methods and X-ray singlecrystal diffraction analysis. The absolute configuration of 257 was determined as (1S,4R,5S,7R,10R,4 R,5 R,6 S,7 R) by a Flack parameter of −0.01(23). The structure elucidation of 258 was difficult to establish by NMR spectroscopy due to the multiple attached quaternary carbons, but an X-ray diffraction study allowed the structure and absolute configuration to be determined by obtaining a Flack parameter with a value of −0.02(5). Likewise, the X-ray diffraction of 259 confirmed its structure and absolute configuration as (4R,5S,4 R,5 R,6 S), which was supported by a reliable Flack parameter of 0.06(9). Potential anti-inflammatory activities of 257–259 were evaluated in lipopolysaccharide-activated RAW 264.7 cells, of which 258 and 259 showed significant activity. A biosynthetic route was proposed for these interesting structures [112].

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J. C. Pardo-Novoa and C. M. Cerda-García-Rojas 14

O 3 O 13

5

11

3'

15'

13' 7'

9

11'

9'

O 15

O

1

5'

1

7

12

O

14' 1'

9 5

3

12'

7

N

14

O

O

15

257 (nardochinoid A)

258 (nardochinoid B) 13' 15' 14'

12'

3'

11' 5'

O

7' 1' 9'

O O 259 (nardochinoid C)

The new ainsliatriolides A (260) and B (261), guaianolide sesquiterpenoid trimers, were isolated from Ainsliaea fragrans, a medicinal plant distributed in the southern part of China [113]. These structures possess a unique hydrocarbon skeleton through two different C–C linkages of the type A (4−2 /15−14 ) and B (15 /15 ). In this work, the structural elucidation was carried out by an extensive analysis of their spectroscopic data, while the determination of absolute configuration was performed by single-crystal X-ray diffraction experiments using the Flack parameter that provided values of 0.12(4) for 260 and 0.17(7) for 261. Ainsliatriolide A (260) showed potent cytotoxicity toward the A549, HT-29, BEL-7402, and HL-60 cancer cell lines with an averaged IC 50 value of 1.17 μM [113]. 13'' 8'' 11'' 12''

9'' 7'' 14''

10''

6'' 5''

1'' 2''

4'' 3''

O

15'

3'

2

1 5

15

10'

7'

10

4

1'

8' 13'

3

2'

6' 12' 11'

O O

5'

O

O

14 15''

4'

O

O

O

9'

14'

O

8 6

O

OH O O O

9

HO O

7

O

OH

O OH

O

260 (ainsliatriolide A)

O 261 (ainsliatriolide B)

4.2 Calculation of the Flack and Hooft Parameters Two new sesquiterpenoids derived from the eudesmane skeleton were isolated from the ethanol extracts of the fruits of Daucus carota. The new compounds 262 and 263 were elucidated taking into account an evaluation of NMR and mass spectrometric

New Techniques of Structure Elucidation for Sesquiterpenes

299

data [114]. The absolute configuration of 262 was assigned in light of a single-crystal X-ray diffraction study. The data collection was achieved using Cu Kα radiation allowing calculation of the Flack x = 0.07(19) and Hooft y = 0.06(7) parameters based on 1125 Friedel pairs. Given that both parameters were close to zero, the absolute configuration of 262 was assigned as (1S,3R,4R,5S,7R,10S). In view of the close structural relationship between eudesmane derivatives 262 and 263, the relative configuration of the latter compound was also determined as (1S,3R,4R,5S,7R,10S). The position of the d-glucose moiety was confirmed through an HMBC experiment [114]. OH 15

OH 9

1 2 3

HO 14

8

10 4

5

HO

12 7

OH

6

11

OH

HO

O

OH

13

262

O

OH OH

HO 263

A study to explore the diastereoselectivity in the 1,3-dipolar cycloaddition of diazomethane to the exocyclic double bond of α,β-unsaturated sesquiterpene lactones was undertaken [115]. A cycloaddition reaction between zaluzanin A (264), isolated from Zaluzania augusta (Fig. 3), and diazomethane yielded spiropyrazoline 265 with high diastereoselectivity (Fig. 4) [115]. Further cycloaddition reactions performed on parthenin, coronopilin, and psilostachyin gave their corresponding spiropyrazolines (266–268) with total chemoselectivity, while the diastereoselectivity toward the (11S)-stereoisomer ranged between 86–100%, while that of mexicanin I acetate, helenalin, and helenalin acetate yielded the (11R)-diastereoisomer (269–271). Treatment

Fig. 3 Inflorescence of Zaluzania augusta (Lag.) Sch.Bip. Photograph courtesy of Dr. J. Martín Torres-Valencia

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J. C. Pardo-Novoa and C. M. Cerda-García-Rojas

Fig. 4 1,3-Dipolar cycloaddition of diazomethane to zaluzanin A (264) to yield the spiropyrazoline 265

CH2N2

OH

HO O

OH

HO O

Et2O/MeOH

O

O

N

N

265

264 (zaluzanin A)

of helenalin and its acetate with an excess of diazomethane provided dipyrazolines (272–273) with 98% of diastereoselectivity toward the (2R,3S,11R) diastereoisomer. The absolute configurations of all compounds were verified by X-ray diffraction analysis by means of the calculation of the Flack and Hooft parameters [116]. O O

HO

HO

HO O

O

O

O

N N

266

267

268

269

N N

N N

O O

O

RO

N N O

O O AcO

O

N N O

O

O

O O

N N

270 R = H 271 R = Ac

RO

O N N

272 R = H 273 R = Ac

The phytochemical analysis of the aerial parts of Stevia tomentosa (Fig. 5), collected in Epazoyucan, Hidalgo, Mexico, afforded a new sesquiterpene lactone that was accorded the trivial name of epazoyucin (274) [117]. This new guaianolide was characterized by its physical and spectroscopic data. The absolute configuration of this rare product having two β-oriented epoxides in the five-membered ring was determined as (–)-(1S,3S,4S,5R,7R,8R,11S)-1,5:3,4-diepoxyguaia-10(14)en-12,8-olide (274) by single-crystal X-ray diffraction analysis that allowed evaluation of the Flack and Hooft parameters. In addition, a vibrational circular dichroism study accomplished with DFT calculations complemented the streochemical study of 274 [117].

O

O

O

O 274 (epazoyucin)

Chemical analysis of the roots of Lasianthaea aurea, collected near Morelia, Mexico, yielded a sesquitepenoid known as podocephalol (275) [118]. The absolute

New Techniques of Structure Elucidation for Sesquiterpenes

301

Fig. 5 Inflorescence of Stevia eupatoria Willd. Photograph courtesy of Dr. J. Martín TorresValencia

configuration of this ar-himachalene derivative was analyzed by X-ray diffraction analysis of its acetyl derivative 276, for which the structure was solved by direct methods and refined to a discrepancy index of 4.7%. Calculation of the Flack parameter gave an x = 0.10(2) value, while estimation of the Hooft parameter provided y = 0.11(11). Inversion of the absolute configuration of the X-ray structure increased these parameters to x = 0.9(2) and y = 0.89(11), respectively, confirming that podocephalol acetate (276), and therefore podocephalol (275) had the (7R) configuration. The crystal data also allowed the identification of unusual supramolecular layers in the cell package of 276. In addition, vibrational circular dichroism measurements of 276 reinforced the assignment of the absolute configuration of these molecules [115]. OR

275 R = H (podocephalol) 276 R = Ac (acetyl-podocephalol)

The tricyclic sesquiterpenoid rasteviol (277), obtained by reduction with sodium borohydride in methanol of the natural product rastevione isolated from the roots of Stevia serrata collected near Morelia, Mexico, was transformed into the novel sesquiterpenoids 278–283 by means of multiple Wagner-Meerwein rearrangements and hydride shifts that occurred when 277 was treated with Et2 O:BF3 (Scheme 1). Four of these new compounds (279, 281–283) possess new hydrocarbon skeletons

302

J. C. Pardo-Novoa and C. M. Cerda-García-Rojas O OH

284

OH

O OTs

OAng

O

OH

OH OH 287

278

285

OH OAng

O OAng

OAng

HO

O OAng

OAng OAng 277 (rasteviol)

283

OH

OAng O

282

279

OAng O

OAng

OAng

OAng 280

281

OH O

OH

286

Scheme 1 Transformation of rasteviol (277) into novel sesquiterpenoid structures by Wagner– Meerwein molecular rearrangements

[119]. All structures were determined by NMR spectroscopy and mass spectrometry in combination with single-crystal X-ray diffraction analyses of 284–287, which supported the structural assignments and confirmed their absolute configurations by calculation of Flack and Hooft parameters. The reaction mechanisms involved in the molecular rearrangements were studied by deuterium-labeling experiments [119]. The bicyclic sesquiterpenoid parvifoline (288) was isolated in good yield from the roots of Acourtia humboldtii, collected near Morelia, Mexico. This benzocyclooctene was modified chemically to generate derivatives 289–299, which were studied as tubulin polymerization inhibitors [120]. The structural analysis and characterization of the new compounds 292–294 and 296–299 were achieved by NMR spectroscopy,

New Techniques of Structure Elucidation for Sesquiterpenes

303

mass spectrometry, and X-ray diffraction analysis. The absolute configurations of epoxide 293 and that of compound 296, which involved a molecular rearrangement of the epoxide group with a transannular 1,5-hydride shift, were verified by calculation of the Flack (x) and Hooft (y) parameters that gave x = −0.03(19) and y = − 0.00(6) for 293 and x = 0.10(3) and y = 0.07(14) for 296. Inversion of the absolute configuration of the X-ray structures increased these parameters to x = 1.03(19) and y = 1.00(5) for 293 and x = 0.90(3) and y = 0.93(14) for 296. Confirmation of the absolute configuration of the parvifoline derivatives was an important task to be achieved considering their biological interactions with tubulin, which were studied by using in vitro experiments and by docking analysis. Compounds 288, 292, and 295 inhibited the polymerization of the α,β-tubulin heterodimer by 24%, 49%, and 90% as compared to colchicine. Competitive inhibition of 288, 292, and 295 with colchicine were also investigated to support their binding at the colchicine secondary site in α-tubulin, while evaluations of their cytotoxicity on cancer cell lines showed that 295 was active against the HeLa cervix carcinoma and HCT 116 colon carcinoma lines with IC 50 3.3 ± 0.2 and 5.0 ± 0.5 μM, respectively [120]. RO

RO

RO

O 288 R = H (parvifoline) 289 R = Bz RO

290 R = H 291 R = Bz RO

RO

O 294 R = H 295 R = Bz

292 R = H 293 R = Bz

O

O

296 R = H 297 R = Bz

298 R = H 299 R = Bz

5 Conclusions Although research on many aspects of sesquiterpenes has been developed extensively during this present century, the chapter focuses on summarizing reported laboratory investigations that have incorporated new successful techniques for the structure determination of this representative group of organic natural products. The information contained in this contribution has allowed a presentation of the panorama of several new tools that are accessible to natural product researchers currently. It is relevant to point out that the structure elucidation strategies applied in the articles that have been cited can be extrapolated to other groups of naturally occurring and synthetic compounds. The importance of having precise three-dimensional structures through validated molecular models of sesquiterpenes is of high relevance, because they are very useful

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in understanding the chemical reactivity of these organic natural products, as well as in later stages of their research, in explaining their biological properties in terms of molecular interactions with more complex structures, such as the proteins of a biological system. The availability of these new techniques of structure elucidation rapidly produces the generation of large amounts of data in ever shorter times, with increasingly finer precision, with data stored in large information repositories. In future, this will allow the generation of faster and automated scenarios, including artificial intelligence systems, with the aim of solving more complex compound structural problems or the discovery of new molecular phenomena. Acknowledgements We are grateful to Conacyt-Mexico for financial support through grant CB 241053 and for a postdoctoral fellowship awarded to J.C.P.N. The authors wish to thank Dr. J. Martín Torres-Valencia, Universidad Autónoma del Estado de Hidalgo, Pachuca, Hidalgo, Mexico for kindly providing the plant photographs.

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Julio C. Pardo-Novoa was born in Villamar, Michoacán, Mexico on October 2, 1987. He undertook undergraduate studies in Chemistry, obtained his B.Sc. degree from Universidad Michoacana de San Nicolás de Hidalgo, Mexico in 2011, and continued graduate work at the same institution to obtain a M.Sc. degree in 2014 and a Doctor of Science degree in 2018. He is currently carrying out postdoctoral studies in the Department of Chemistry, Cinvestav-IPN in Mexico City. His research interests are focused on the field of natural products, including isolation, structure elucidation, and chemical transformations. During his scientific career, he has published four research articles.

Carlos M. Cerda-García-Rojas was born in Uruapan, Michoacán, México on February 11, 1963. He obtained his B.Sc. degree from Universidad Michoacana de San Nicolás de Hidalgo, in Morelia, Mexico in 1984. He obtained his M.Sc. degree in 1988 and a Doctor of Science degree in 1992 both from Centro de Investigación y de Estudios Avanzados del Instituto Politécnico Nacional (Cinvestav-IPN) under the guidance of Professor P. Joseph-Nathan. After a postdoctoral fellowship at Scripps Institution of Oceanography, University of California, San Diego in La Jolla, California, USA from 1993 to 1994 under the supervision of Professor D. John Faulkner, he became a faculty member at the Department of Chemistry, Cinvestav-IPN, Mexico City, where he has been promoted to Professor of Chemistry. He has developed research on the determination of the absolute structure and molecular conformation of bioactive natural products with medicinal and industrial relevance by molecular modeling using quantum mechanical calculations and nuclear magnetic resonance. His research interests also focus on the chemical transformation of natural compounds, including sesquiterpenes and diterpenes, to explore their biological activities as anticancer agents. Derived from interdisciplinary collaborative work, he has coauthored 150 research publications and has supervised 14 Master of Science and nine doctoral theses. He has been honored with the 2012 Martín de la Cruz Prize in Chemistry and Biology, awarded by the General Health Council on World Health Day, Mexico City, for his scientific research applied to the development of medicines from natural products.

Human Endogenous Natural Products Yingjie Bai, Liyun Zhang, and Xiaoguang Lei

Contents 1 2

3

4

5

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Endogenous Natural Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Endogenous Amines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Steroid Natural Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Fatty Acid-Derived Natural Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Host–Microbe Co-metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Presence and Function of the Human Microbiota . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Co-Metabolism of Tryptophan and Other Aromatic Amino Acids . . . . . . . . . . . . 3.3 Co-Metabolism of Bile Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Co-Metabolism of Choline . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Co-Metabolism of Cysteine and Other Sulfur-Containing Compounds . . . . . . . . Human Natural Products and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Steroidogenic Inherited Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Trimethylamine Oxide and Atherosclerosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Techniques for the Identification of Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Y. Bai · L. Zhang Beijing National Laboratory for Molecular Sciences, Key Laboratory of Bioorganic Chemistry and Molecular Engineering of Ministry of Education, Synthetic and Functional Biomolecules Center, College of Chemistry and Molecular Engineering, Peking University, 282 Chengfu Road, Haidian District, Beijing, China e-mail: [email protected] L. Zhang e-mail: [email protected] X. Lei (B) Beijing National Laboratory for Molecular Sciences, Key Laboratory of Bioorganic Chemistry and Molecular Engineering of Ministry of Education, Synthetic and Functional Biomolecules Center, College of Chemistry and Molecular Engineering, College of Chemistry and Molecular Engineering and Peking-Tsinghua Center for Life Science, Peking University, 282 Chengfu Road, Haidian District, Beijing, China e-mail: [email protected] © The Author(s), under exclusive license to Springer Nature Switzerland AG 2021 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 114, https://doi.org/10.1007/978-3-030-59444-2_4

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5.2 Nuclear Magnetic Resonance Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 333 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 334

1 Introduction Natural products, also known as “secondary metabolites,” are a class of chemical compounds that are produced by certain living organisms. Not only those compounds generated from living organisms (such as bacteria, fungi, or plants) are natural products, but many human endogenous biochemicals, including trace amines and fatty acid derivatives, are also natural products based on their definition. Besides these endogenous natural products produced by human cells, the human microbiome also produces natural products, as elucidated by the Human Microbiome Project (HMP) [1]. As a result of recent achievements made in metabolomics research and from the HMP, some previously accepted information has been corrected regarding endogenous natural products. Therefore, this contribution will describe the current understanding and the recent progress made on endogenous natural products that are produced by human cells and symbiotic microbes.

2 Endogenous Natural Products 2.1 Endogenous Amines 2.1.1

Catecholamines

Neurotransmitters are essential for the functioning of the nervous system, and many endogenous amines (e.g., catecholamines) work as neurotransmitters [2]. Catecholamines, including dopamine, norepinephrine, and epinephrine, are key chemical neurotransmitters and hormones that regulate physiological processes. The biosynthesis of catecholamines starts from tyrosine. First, tyrosine is 3-hydroxylated to form l-DOPA. Next, aromatic acid decarboxylase (AADC) transforms lDOPA to the first catecholamine (dopamine). The second catecholamine (norepinephrine) is the β-hydroxylation product of dopamine. Finally, phenylethanolamine N-methyltransferase (PNMT) converts norepinephrine into epinephrine [3] (Fig. 1). Although related metabolic research is extensive, misunderstandings in the area occur because of limitations in the experimental protocols used previously. Studies in the last 30 years have permitted the construction of a more accurate atlas for the disposition and metabolism of catecholamines than before. The deamination of norepinephrine and epinephrine by monoamine oxidase (MAO) yields the reactive aldehyde 3,4-dihydroxyphenylglycolaldehyde (DOPEGAL), which is reduced to form

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O HO

OH

AADC

NH2

HO

HO

hydroxylase

HO

NH2 dopamine

DOPA

OH

HO PNMT HO

N H

NH2 OH norepinephrine

OH OH epinephrine

MAO

HO HO CHO

HO DOPAL

CHO

HO

OH DOPEGAL

HO

HO HO COOH

HO

OH

HO

MeO

OH OH

OH DOPAC

DHPG

DHPG

Fig. 1 Metabolism of catecholamines

dihydroxyphenylglycine (DHPG) rather than oxidized to 3,4-dihydroxymandelic acid (DHMA) (Fig. 1). The homeostasis of catecholamines in neurons is also different from our earlier understanding of this process. The neurons store catecholamines dynamically, and the leaked cytoplasmic catecholamines are oxidized intraneuronally. It is worth noting that the catecholamine turnover in the neurons depends upon vesicular leakage rather than neuronal activity [3]. Besides the central nervous system, important sources for catecholamines and catecholamine metabolites include the peripheral nervous system, the gastrointestinal tract, and the urinary system. In fact, the catecholamine metabolite 3-methoxy-4hydroxyphenylglycol (MHPG), is synthesized mainly by the peripheral sympathetic nerves from the O-methylation of DHPG. A related metabolite, MHPG sulfate, has been correlated with norepinephrine oxidation in the gastrointestinal tract, possibly by the liver. About 50% norepinephrine and 45% dopamine in the body is synthesized in the mesenteric organs. Other dopamine metabolites (mainly homovanillic acid) are derived in the gastrointestinal tract efficiently. The urinary system is also capable of producing catecholamine, and urinary dopamine is synthesized by plasma DOPA decarboxylation in the kidney parenchyma [3].

2.1.2

Trace Amines

Another family of endogenous amines besides the neurotransmitters are called the trace amines, which include β-phenylethyalmine (PEA), p-tyramine (TYR), tryptamine, octopamine, and some of their metabolites. The biosynthesis of these trace amines is similar to the catecholamines, but additional amino acid decarboxylases are involved and the decarboxylation undertaken is not quite specific. For example, leucine can be decarboxylated by valine decarboxylase and methionine decarboxylase, while no enzyme is termed “leucine decarboxylase.” In fact, nearly all of the decarboxylated amines (except for asparagine, glutamine, and glycine)

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of proteinogenic amino acids are known, while many transformations are related to microbiota. There are also many trace amine derivatives derived from aromatic decarboxylated amines. For example, dopamine-β-hydroxylase converts p-tyramine into p-octopamine, while catechol-O-methyl transferase converts dopamine into 3methoxytyramine. The chemical diversity of aromatic trace amines is increased further by methylation to form N-methylphenylethylamine, N-methyltyramine, synephrine, and N-methyltryptamine [4] (Fig. 2). The function of the trace amines remains largely unknown. Many trace amines are proven or putatively proven to be endogenous neural active compounds. Three trace amines, β-phenylethylamine, p-tyramine, and tryptamine, have been demonstrated as neuron modulators, although their action is complex. High-affinity mammalian receptors for the archetypal trace amines have been investigated for a long time, and these turned out to be a family of G-protein-coupled receptors (GPCRs) named trace amine-associated receptors (TAARs). Humans have six functional TAAR genes, of which most are orphan receptors expressed in the olfactory system, except for TAAR1. As the target of β-phenylethylamine and p-tyramine, TAAR1 plays a role in trace amines secondary amines

primary amines O OH NH2

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Fig. 2 Metabolism and generation of some archetypal trace amines

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the central and peripheral nervous systems. In the TAAR1 variant, the syntheses of β-PEA and TYR occur in dopaminergic terminals. Unlike dopamine, which is stored within synaptic vesicles, trace amines readily diffuse across the plasma membranes. It is known already that the organic cation transporter OCT2 plays a role in the reuptake of trace amines into the presynaptic terminals, while the postsynaptic importer of trace amines is still unknown. Since TAAR1 is located predominantly in the cytoplasm, both pre- and postsynaptic effects are plausible. The heterodimerization of TAAR1 with the dopamine D2 receptor D2R may trigger translocation of TAAR1 onto the cytoplasmic membrane, which subsequently promotes biased D2R signaling through the Gi cascade instead of the arrestin pathway. Unlike D2R, TAAR1 does not interact with D1. Dopamine and its metabolites also play multiple roles in the TAAR1 pathway. Dopamine itself activates presynaptic D2R and inhibits the synthesis of PEA/TYR via the Gi signal transduction pathway. An extracellular catechol-Omethyl transferase transforms dopamine into a trace amine, 3-methoxytyramine (3MT), which activates both pre- and postsynaptic TAAR1 as the archetypal trace amines (Fig. 3). Nutrient-induced hormone secretion is also regulated by TAAR1 in the peripheral nervous system, and this could be developed as a novel therapeutic target for diabetes and obesity [4].

2.2 Steroid Natural Products Besides the endogenous amines, steroids are also a family of endogenous secondary metabolites. The most important chemicals in this family are the steroid hormones, which regulate a number of physiological and developmental processes from birth to death. The biosynthesis of steroid hormones starts from the cleavage of cholesterol by P450scc to form pregnenolone, the last common intermediate of steroid hormone biosynthesis. The 3-dehydrogenation of pregnenolone yields progesterone, which can be transformed further to aldosterone. Progesterone can also undergo 17hydroxylation to form 17-hydroxyprogesterone, which is the progenitor of cortisone and cortisol. The C-17–C-20 lysis of pregnenolone gives dehydroepiandrosterone, which is the common intermediate of androstane-type and estrane-type steroids. The 3-dehydrogenation and 17-dehydrogenation of dehydroepiandrosterone yields testosterone as the final product via androstenedione or androstenediol. Further demethylation of testosterone or androstenedione gives estradiol or estrone. Testosterone is also the precursor of 5α-steroid hormones such as dihydrotestosterone and androsterone. The majority of their biosynthetic enzymes are either cytochrome P450s (CYPs) or hydroxysteroid dehydrogenases (HSDs), and some transformations are not strictly order-dependent, because of the promiscuity of these enzymes. Moreoverever, human P450c17 does not prefer 17-hydroxylprogesterone and the C-17– C-20 lysis of 17-hydroxylpregnenolone is the preferred pathway to androstane-type steroids in human beings [5] (Fig. 4). Steroid hormones are important in homeostasis. Cortisone and cortisol are glucocorticoids that inhibit inflammation and dehydration while increasing blood sugar.

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O

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Fig. 3 Current understanding of TAAR1 activation by the archetypal trace amines. Hollow arrows: Chemical transformations. Solid arrows: Stimulating signal transductions. Blocked arrows: Inhibitory signal transductions [4]

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cholesterol P450scc O

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Fig. 4 Major metabolism pathways of human steroid hormones

Aldosterone is a mineralocorticoid that stimulates active reabsorption of sodium and associated passive reabsorption of water in the kidney. Androstenedione, testosterone, estrone, estradiol, progesterone, and some other related compounds are sex steroids that govern the development and maintenance of secondary sex characteristics such as the menstrual cycle. The steroid hormones usually function by activating the corresponding nuclear receptor, and the genetic disorders related to steroid hormones will be discussed in Sect. 4. It should be noted that some of the non-hormone steroids play very important roles in life, for example, cholesterol modification is essential for Sonic hedgehog signaling, which, for example, is important for the evolution of radio- and chemoresistance of several types of tumors [6].

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2.3 Fatty Acid-Derived Natural Products The metabolism of fatty acids also leads to a rich source of endogenous natural products. Among them, the polyunsaturated fatty acid-derived (PUFA) metabolites are well studied both in terms of their metabolism and function. Polyunsaturated fatty acids are essential fatty acids that can be categorized as ω-6 fatty acids (e.g., arachidonic acid) or ω-3 fatty acids (e.g., eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA)). The metabolism of arachidonic acid (ARA) starts from the hydrolysis of various membrane arachidonyl phospholipids. The incomplete hydrolysis of arachidonyl phospholipids gives endocannabinoids including 2-arachidonoyl glycerol and anandamide, while the complete hydrolysis releases free ARA. Oxidation of ARA by cyclooxygenase, lipoxygenase, or cytochrome P450 enzymes gives various classes of metabolites including thromboxanes, lipoxins, and most of the prostaglandins and leukotrienes, while transcellular biosynthesis is common in arachidonic acid metabolism [7] (Fig. 5). It should be noted that a few prostaglandins and leukotrienes like PGD3, PGE3, and LTB5 are transformed from eicosapentaenoic acid (EPA) rather than arachidonic acid. Besides ω-3 fatty acids, functional metabolites of ω-6 fatty acids also are known from the metabolism of EPA and give various EPAoriginating resolvins, while metabolism of DHA generates maresins and protectins together with the DHA-originating resolvins [8] (Fig. 6). thromboxanes, lipoxins, prostaglandins, and leukotrienes

O OH arachidonic acid

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OH

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Fig. 6 Metabolism of DHA and EPA

The ARA metabolites, including PGE2, other 2-series PGs, and the 4-series LTs, usually bind to a specific receptor (typically, a GPCR) and they mediate or upregulate inflammation. In contrast, extensive animal models of inflammation indicate that the ω-3 fatty acid metabolites including the resolvins, protectins, and maresins are anti-inflammatory or inflammation resolving. The biosynthesis and receptors of ARA metabolites are targets of various anti-inflammatory drugs. Interestingly, a

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recent discovery has indicated that aspirin, a very widely used anti-inflammatory drug, triggers the production of novel anti-inflammatory resolvins and lipoxins after binding to its target, COX-2, which may be an explanation for its well-known painkilling action [8]. Besides PUFAs, platelet-activating factor (PAF) is another multifunctional endogenous compound. There are two pathways for PAF biosynthesis, the remodeling and the de novo pathway. The remodeling pathway, in which PAF is synthesized by substitution of an arachidonoyl moiety with an acetyl in 1-O-alkyl-2arachidonoylglycerophosphocholine, is believed to be the primary source of PAF, especially during inflammation. The de novo pathway, in which 1-O-alkyl-snglycero-3-phosphate is acetylated, hydrolyzed to 1-O-alkyl-2-acetyl-sn-glycerol, and converted to PAF by a dithiothreitol-insensitive CDP-cholinephospho-transferase, has been proposed as a source of PAF for housekeeping physiological roles. Besides the activation of platelets, as its name indicates, PAF initiates inflammation by action on the PAF receptor [9]. Additionally, it has been shown that PAF receptor activation is neuroprotective and that PAF induces neuronal apoptosis independent of the PAF receptor [10]. The platelet-activating factor is also believed to play a non-pathological role in stimulating uterine contraction at the end of pregnancy.

3 Host–Microbe Co-metabolism 3.1 Presence and Function of the Human Microbiota As the human commensal microorganisms (i.e., the microbiota) are estimated to be more than ten times greater compared to those of human somatic and germ cells, they are believed to explain the individual differences of normal physiology and predisposition to disease. Therefore, a project named the human microbiome project (HMP) was launched after the human genome project [1]. The human microbiome project led to the discovery that the diversity and abundance of each habitat’s signature microbes vary largely, but the microbiota of the specific niche of an individual is quite stable [11]. A further data study named the expanded HMP has revealed a more complicated atlas of personalized microbiome composition, function, and dynamics [12]. According to the HMP, a “long tail” of low-abundance genes and pathways probably encodes many uncharacterized bioactivities and metabolites in the microbiomes. Therefore, host–microbe co-metabolism is believed to be essential in elucidating the roles of the microbiota in health and diseases [11]. The co-metabolism of the chemicals of host and its microbiota can be categorized roughly into three patterns: the first pattern involves the chemicals produced solely by microbes, like the production of vitamins, siderophores, and short-chain fatty acids [13]. The second pattern involves those chemicals produced solely by the host, like the production of primary bile acids [14] and most compounds included in Sect. 2. The third pattern involves the chemicals produced collaboratively by the

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host and the microbes. Specifically, some metabolites are synthesized by the microbes and are biotransformed by the host such as trimethylamine oxide [14], while other metabolites are synthesized by the host and biotransformed by the microbes, as exemplified by unconjugated secondary bile acids [14]. The remaining metabolites are generated endogenously and are biotransformed at least once by both the host and its commensal microbes, such as conjugated secondary bile acids [14]. There are also some miscellaneous metabolites for which their categorizations are troublesome, such as spermidine and tyramine, which are produced by both the host and its commensal microbes [4]. Many microbiota-related metabolites are bioactive. By affecting differentiation, migration, proliferation, apoptosis, and other processes of the host cell, they have multiple physiological or pathological effects on the eukaryotic host or other symbiotic bacteria in the environment. As shown in Fig. 7, the action of short-chain fatty acids produced by commensal bacteria on the GPR43/GPR109A of dendritic cells will inhibit food allergies, while the action of the same compounds on the GPR43 of T cells will inhibit the onset of allergic bowel syndrome (IBD). When targeting GPR109A of microglial cells, short-chain fatty acids may induce Parkinson’s disease. The function of endogenous natural products is also diverse: trimethylamine oxide can cause cardiovascular disease and arteriosclerosis; bacterial lipopolysaccharides may activate TLR4/9 and induce non-alcohol fatty liver disease (NAFLD), and 4ethylphenol sulfate, as secreted by B. fragilis, may lead to autistic spectrum disorders (ASD) [15].

Fig. 7 Diverse and multiple functions and targets of symbiont bacterial natural products [15]

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3.2 Co-Metabolism of Tryptophan and Other Aromatic Amino Acids Tryptophan is a common substrate for many host microbe co-metabolites, which can be categorized into kynurenine-, serotonin-, and indole-related metabolites. Co-metabolism of tryptophan relates to a variety of neural and immune pathological states, including ASD, depression, Alzheimer’s disease, and many autoimmune diseases. To study their origin, Olson and coworkers developed a quantitative ex vivo assay using 6-fluorotryptophan and 19 F-NMR to study the origin of 30 tryptophan metabolites (Fig. 8). According to their results, some metabolites only emanate from the liver, while others only come from the feces (Fig. 8). Therefore, the tryptophan metabolic pathways in the liver and gut microbiota are quite different. In addition, it has been found that the gut microbiome metabolism is disrupted by antibiotic treatment, as may be expected [16]. Tryptophan also shares some common biotransformation pathways with other aromatic amino acids including phenylalanine and tyrosine. A gene cluster in the gut symbiont Clostridium sporogenes has been observed to transform these aromatic amino acids into arylpyruvates, which are further oxidized to arylacetates or reduced to aryllactates, arylacrylates, and arylpropionates. The gene cluster is also found in C. botulinum, Peptostreptococcus anaerobius, and C. cadaveris, while these metabolites were found further to affect intestinal permeability and systemic immunity in a mouse model [17].

3.3 Co-Metabolism of Bile Acids Another good example of host microbe co-metabolism is the generation and transformation of bile acids. Bile acids are synthesized originally from cholesterol in hepatocytes. After efficient conjugation to either taurine or glycine, bile acids are transported out of the hepatocytes and secreted into the small intestine as one of the principal constituents of bile. In the gut, a ubiquitous microbial enzyme named bile salt hydrolase cleaves the amide bond and releases the unconjugated bile acid, which further can undergo various biotransformation steps in different bacteria, including oxidation, epimerization, 7-dehydroxylation, esterification, and desulfation. At the terminal ileum, the processed bile acids are absorbed into the blood by passive diffusion and active transport and recycled to the liver via the portal vein, a phenomenon known as enterohepatic circulation. The co-metabolized bile acids such as DCA and LCA are conjugated to taurine or glycine along with the host-synthesized bile acids in the hepatocytes, which has further increased the chemical diversity of bile acids (Fig. 9) [18]. In classical biochemistry, bile acids are believed to be the solubilizers of lipidsoluble nutrients, which thus facilitate their assimilation. Nevertheless, recent target identification of bile acids has revealed their systemic endocrine functions in versatile

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kynurenine pathway metabolites O

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Fig. 8 Diverse tryptophan metabolites in the gut that are produced by different pathways. The compounds marked in blue are produced by two liver pathways including the kynurenine pathway and the serotonin pathway, while the compounds marked in red come from the feces. The circled compound, 6-F-tryptophan, is used as a labeled tryptophan to study the origin of indole-related metabolites [16]

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Fig. 9 Targets of the bile acids

signaling pathways. The first discovered bile acid-activated nuclear receptor, FXR, regulates genes involved in the enterohepatic recycling and detoxification of bile acids to provide a negative feedback signal for bile acid biosynthesis. It is worth stressing that besides FXR, the pregnane X receptor (PXR) and vitamin D receptor (VDR) are also bile acid-activated nuclear receptors. Another well-known bile acid receptor is TGR5, which is activated by secondary bile acids and trigger metabolic actions including energy homeostasis, thermogenesis, and insulin signaling through the cAMP axis. In addition to TGR5, bile acids are also modulators for other GPCRs including muscarinic receptors and formyl peptide receptors (FPRs) [14, 18] (Fig. 10). Recently, MRGPRX4, a GPCR related to itching, was found to be targeted by bile acids [19]. The co-metabolism of bile acids is also a good example that the metabolism of endogenous compounds can be affected by many factors. It is well known that species variation is an important consideration in the diversity of endogenous compounds. For example, the mouse but not the human synthesizes 6-hydroxylated bile acids (e.g., muricholate) efficiently, while their recycled 7-dehydroxylated bile acids undergo

Fig. 10 Bile acid metabolism

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Fig. 11 Rare bile acids discovered in other mammals or under stress conditions [22]

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7-hydroxylation in hepatocytes [18]. In other mammals, besides humans and mice, additional types of bile acids have been discovered, including the 1-hydroxy bile acids (e.g., vulpecholate) and the 23-hydroxy bile acids (e.g., bitocholate) [20, 21]. The metabolic state of bacteria is also a factor for chemical diversity. For instance, stress is believed to be a reason for the production of 5α-bile acids (e.g., allocholic acid), which are known shunt metabolites in the 7-dehydroxylation pathway [22] (Fig. 11).

3.4 Co-Metabolism of Choline It is well known that choline is an essential moiety in the makeup of many phospholipids in the cell membrane, but it is less understood that choline is involved in host microbe co-metabolism. In fact, choline is metabolized to trimethylamine by gut microbiota, which was first reported in 1910. The free trimethylamine (fishy odor) can either be oxidized by the liver enzyme FMO3 to form the atherosclerosis-inducing trimethylamine oxide or be reduced by methanogenic archaea to the powerful greenhouse gas methane. However, the gene cluster and enzymes responsible remained unknown until recently, when Balskus and coworkers reported that Desulfovibrio desulfuricans uses a glycyl radical enzyme CutC (Ddes_1357) and a glycyl radicalactivating protein CutD (Ddes_1357), to cleave choline into trimethylamine and acetaldehyde [23] (Fig. 12).

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Fig. 12 Bacterial metabolism of choline

3.5 Co-Metabolism of Cysteine and Other Sulfur-Containing Compounds The catabolism of methionine, cysteine, and taurine is also an important source of many low-molecular-weight endogenous compounds, especially sulfur-containing endogenous compounds in the gut. Methionine is the precursor of the gut sulfides including methylthiol and dimethyl sulfide, although the microbial pathway for dimethyl sulfide formation is not established. Methionine and cysteine are also precursors for the thioether-bearing amino acids, such as lanthionine and homolanthionine in microbes. The complete breakdown of these sulfur-containing compounds and the metabolism of the sulfate-reducing bacteria yield endogenous hydrogen sulfide, which may increase bacterial antibiotic resistance, protect them from reactive oxygen species, and function as a gaseous neurotransmitter for the host.

4 Human Natural Products and Disease 4.1 Steroidogenic Inherited Diseases Dysfunction of human natural products may be related to many diseases, including inherited conditions. The dysfunction of the steroidogenic pathway has a direct causal relationship to a range of inherited diseases related to sexual differentiation, reproduction, fertility, hypertension, and obesity. For example, recessive genetic mutations to P450c17 and the resulting absence of 17α-hydroxylase and 17,20-lyase activities block the synthesis of adrenal and gonadal sex steroids. For genetic males, the development of external genitalia is absent, which is termed as male pseudohermaphroditism or “46, XY disorder of sex development”. For genetic females, they are phenotypically normal but fail to undergo adrenarche and puberty, which results in teenage hypertension and sexual infantilism [5].

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4.2 Trimethylamine Oxide and Atherosclerosis While the deficiency of hormone steroids will cause steroidogenic inherited diseases, the abnormal production of trimethylamine oxide is known to be a reason for the increased risk of atherosclerosis as well. One of the best examples of host–microbe co-metabolism-related disease is atherosclerosis. As previously described, the cometabolism of choline by both the liver and the gut microbiota generates trimethylamine oxide (TMAO). Using metabolomics, it has been discovered further that high TMAO levels correlate with a high risk for cardiovascular diseases, and the mechanism involved is that TMAO promotes the upregulation of multiple macrophage scavenger receptors as atherosclerosis biomarkers. In addition, antibiotics and TMAOmimetic enzyme inhibitors are bioactive in terms of reversing atherosclerosis, which has the potential to be developed as translational therapy for the gut microbiota [24].

5 Techniques for the Identification of Metabolites Our human metabolomic profile is far more complex when compared to that of the human genome: the human genome contains approximately 20,000 protein-coding genes, but the human metabolomic profile consists of more than 500,000 compounds (identified so far) and each class of compounds has a high level of diversity [25]. Metabolomics studies are often combined with genomics or proteomics toward a systems biology approach, to provide biochemical insight into the organism being studied. An integrative analysis of an organism’s response to a certain treatment on the transcriptome, proteome, and metabolome levels will lead to a better understanding of the physiological conditions and biological mechanism in complex systems (Fig. 13). Metabolites that represent a diverse group of low-molecular-weight structures include lipids, amino acids, peptides, nucleic acids, organic acids, vitamins, thiols, and carbohydrates, which make their global analysis difficult. The compositional diversity of metabolites provides wide ranges of physiochemical properties, including molecular weight, hydrophobicity/hydrophilicity, acidity/basicity, and boiling point [27]. Not only are human endogenous metabolites very complex, but the sources that produce metabolites are diverse and complicated. Nearly all microbes produce metabolites, including secondary metabolites (natural products) and small-molecule

Fig. 13 The genome–transcriptome–proteome–metabolome cascade [26]

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virulence factors [25]. These metabolite molecules drive complex physiological cycles in humans. There are generally two complementary approaches used for metabolomic investigations: targeted analysis (i.e., metabolic profiling) and untargeted analysis (i.e, metabolic fingerprinting) [28]. The targeted analysis normally focuses on certain kinds of metabolites and this kind of selective metabolite identification is often termed “targeted” metabolomics. The other metabolomics analytical procedure is often termed “untargeted analysis” or “global metabolomics,” which aims to detect all the metabolites in a given sample, regardless of their chemical class or character. For either approach, the identification of novel compounds within a complex, unpurified biological mixture is technically challenging. A typical metabolomics workflow is summarized in Fig. 14: first, factors should be considered such as the number of subjects, the appropriate controls to be used, the types of samples, and how frequently samples will be taken. Second, the collected samples must undergo preparation for either LC–MS or NMR analysis, and the resultant spectra are recorded based on standardized protocols. The datasets are then subjected to statistical analysis to identify the levels of variance between the test groups. Once key metabolites are identified, a univariate analysis can be performed to establish trends in metabolite levels as a function of a certain perturbation (i.e., dose, time) or to quantify differences between the samples. After significance has been established, metabolic pathways are examined to understand the biochemical basis for a given effect. Multiple strategies may be employed to provide a wide range of metabolite identifications. These include the use of mass spectrometry (MS, often coupled to a chromatographic procedure) and nuclear magnetic resonance (NMR) spectroscopy, and these two are the most prevalent analytical approaches. A comprehensive review of metabolomics analysis is beyond the scope of this contribution, although in the paragraphs below an overview is provided of recent work describing key strategies and

Fig. 14 An outline protocol of metabolomic analysis [29]

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applications of targeted and untargeted analyses of human endogenous metabolite identification.

5.1 Mass Spectrometry Mass spectrometry (MS) plays a major role in metabolite profiling due to its high sensitivity. The key objective is to enhance sample throughput and facilitate comprehensive metabolome identification. One approach is to perform rapid LC–MS runs to increase sample throughput, but this comes at the expense of sensitivity and overall metabolome coverage. The second option is to obtain a more comprehensive metabolome coverage through long analytical runs, for example, by making the chromatographic peaks narrower to accommodate more peaks within a defined time window. Narrower chromatographic peaks can be obtained by (1) increased peak capacity; (2) lower ionization suppression; and (3) enhanced signal-to-noise ratio, thus leading to increased sensitivity [30]. The current state of the art in LC–MSbased metabolic profiling is to use reversed-phase HILIC (hydrophilic interaction chromatography), coupled to high-resolution mass spectrometry. A key strategy for metabolomic profiling is to employ only a few methodologies in order to cover as wide a range of metabolite classes as possible. For each compound class, the correct analytical methodologies need to be selected and developed. The group of Wilson has summarized the following required LC–MS conditions for a specified compound class: (1) RPLC–MS for medium-polarity analytes; (2) HILIC– MS for polar metabolites; (3) targeted HILIC–MS/MS or CE–MS/MS for targeted primary metabolites; (4) GC/MS or RPLC/MS for lipidomic analysis, and (5) GC– MS for volatile components [31]. There is now a number of applications emerging employing LC–MS in metabolomics. Biomarker discovery often starts with the study of animal models, for example, obesity, cancer hepatopathy, and cancer nephrotoxicity [31]. Metabolic profiling has been used successfully to characterize metabolic pathways disrupted in mouse models of human diseases, including cardiac disease and type 2 diabetes. The implementation of metabolomics as a screening procedure in large-scale mutagenesis programs has been successful in identifying mutants that possess clinically related phenotypes. Using this approach, models of various human metabolic diseases have been identified, such as a model for branched-chain ketoaciduria, and a model of lipotoxic cardiomyopathy. A substantial amount of work has focused on understanding the role of lipotoxicity and insulin resistance in humans. Liquid chromatographyMS lipidomics has been used to investigate inflammation in the adipose tissue of obese women, suggesting that the content of ceramides and long-chain fatty acids in triglycerides in the tissues correlates with the degree of fatty liver by comparing women with a similar body mass index but with different degrees of hepatic steatosis. Additionally, LC–MS-based metabolite profiling has been used to identify serum metabolic biomarkers of heart failure, where pseudouridine and 2-oxoglutaric acid were found to be potential markers that are being assessed in further targeted work

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to define whether these differences are the cause or effect of the pathophysiology of heart failure [32]. Newgard’s research group has applied target analysis of up to 250 metabolites to study heart-related diseases including myocardial ischemia and planned myocardial infarction [33].

5.2 Nuclear Magnetic Resonance Spectroscopy The sensitivity of NMR spectroscopy is not comparable to that of mass spectrometry, but it offers advantages for compounds that are difficult to be ionized or else require derivatization for MS. Using stable isotope labeling, NMR spectroscopy offers several advantages in elucidating the dynamics and mechanisms of metabolite transformations and for exploring the compartmentalization of metabolic pathways by its inherent capability to monitor isotope enrichment. Nuclear magnetic resonance spectroscopy can be used to measure a wide variety of spin 1/2 nuclei, including the biologically relevant 1 H, 13 C, 31 P, and 15 N as well as the pharmaceutically relevant 19 F. Among these, 1 H, 19 F, and 31 P are at high abundance, while the lower natural abundance of 13 C and 15 N can be useful in measuring enrichment due to catabolism of labeled precursors as a function of specific biological processes or pathways. The positional enrichment of a molecule in both 13 C and 15 N NMR spectroscopic methods may be used to establish which pathways are being activated or inactivated. Samples for NMR metabolomic profiling are collected in a uniform way to minimize variability and are analyzed in terms of their NMR profiles to collect data on all metabolites potentially present in the sample. Using pattern recognition (e.g., principal component analysis and partial least squares discriminant analysis), the potential biomarkers usually can be identified using visualization tools. The identified markers eventually will be placed in a metabolic pathway to provide insight into the biochemical phenomena are observed. Two-dimensional (2D) NMR methods offer an improved resolution for unambiguous identification of metabolites in a mixture, including COSY (correlated spectroscopy, TOCSY (total correlation spectroscopy), and HSQC (heteronuclear single-quantum correlation). Automation of metabolite identification can be achieved using software with combined TOCSY and HSQC data. By setting tolerance levels for the matching of 1 H and 13 C NMR signals, one can maximize compound identification efforts while minimizing false positives [35]. The use of 1 H NMR spectroscopy is well documented, and its application has been used in a wide range of fields and sample matrices, such as biofluids, plants, and natural product mixtures. Biofluids NMR metabolomics offers the potential for a thorough understanding of disease pathogenesis and the identification of disease biomarkers. For a metabolic biomarker to be useful clinically, its level must clearly associate with the disease risk or its progression and should be insensitive to variables (e.g., ethnicity, diet, and location). Blood plasma or urine 1 H NMR spectra usually are measured on samples obtained from both healthy and unhealthy volunteers. In a recent investigation to catalog the metabolic components of urine, a comprehensive study of the human

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urine metabolome using NMR, GC/MS, ICPMS, NMR, GC/MS, ICPMS, direct infusion, and LC/MS/MS was conducted, and 445 unique urine metabolites were identified. The potential of 1 H NMR metabolic profiling to identify the trace level of body fluids was demonstrated and validated as a rapid and non-destructive tool for forensic analysis. Plant metabolomic studies have addressed the effects of genotype, ecotype, and environmental stressors, such as drought and submergence, which can provide metabolic phenotype information for chemical genomics studies in plants. In addition to the primary metabolome, plants also produce an immense number of secondary metabolites (ca. 200,000) that can interact with beneficial or harmful organisms. Nuclear magnetic resonance metabolomics is also useful in discovery-oriented natural products chemistry. 2D-NMR spectra of unfractionated sample extracts can facilitate the identification of novel compounds that might be lost during chromatographic fractionation due to the low abundance or instability of the natural products. A common approach, activity-guided fractionation in natural product research, may fail to characterize synergistically interacting biogenic small molecules. Using NMR metabolomics, unique spectral features can be identified and correlated to a phenotype of the biological property of interest. In summary, efficient metabolite identification is one of the central challenges in metabolomics. Given the chemical diversity of human metabolites, new techniques that make metabolite identification easier and more robust are still necessary. Over the past 10 years, significant advances in the methodology and software/databases used hat are associated with these platforms have greatly improved compound identification.

6 Conclusions This contribution provides an overview of the research progress made on human endogenous compounds. In summary, the natural products produced by humans are relatively well understood and only minor enhancements in the laboratory techniques used for their study are required for their complete understanding. In contrast, those natural products produced from microbes are not well understood and further information on their functional and causal relationships is needed. Considering the complexity of microbiota, a better comprehension of their variations due to sampling sites, health conditions, sampling times, and ethnic group diversity are all required. Proper animal models, precise experiment design, and appropriate biochemical characterization are needed as well. Many non-canonical mechanisms, including the gut brain axis and microbiota microbiota interactions also occur due to the action of microbiota natural products, and these also require further investigation. A better understanding of the “dark matter” will not only reveal more biological mechanisms that can be described with known methods and well-studied factors as “known knowns” but also elucidate even more biological mechanisms that cannot be described with either known methods or known factors as “unknown unknowns.”

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Acknowledgements Financial support from the National Natural Science Foundation of China Grant (21625201, 21961142010, 21661140001, 91853202, and 21521003), the National Key Research and Development Program of China (2017YFA0505200), the Beijing Outstanding Young Scientist Program (BJJWZYJH01201910001001) is gratefully acknowledged.

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Y. Bai et al. Yingjie Bai received his B.Sc. degree from Peking University in 2017. He is currently conducting his Ph.D. research under the supervision of Prof. Xiaoguang Lei at Peking University. His research focuses on the biological functions of human endogenous lipids.

Liyun Zhang obtained her Ph.D. degree from the University of Minnesota, Twin Cities in 2016. Her Ph.D. thesis focused on the identification and characterization of bioactive compounds from complex matrices, under the direction of Prof. Devin G. Peterson. After graduation, she worked as a postdoctoral research scientist on the topdown proteomics analysis of large molecules under the guidance of Emeritus Prof. Jack Henion of Cornell University. In 2019, she moved back to China and became an Associate Professor in the research group of Prof. Xiaoguang Lei at Peking University. Currently, her main research interest is chemoproteomics and metabolomics.

Xiaoguang Lei obtained a B.S. in chemistry from Peking University in 2001, and a Ph.D. in organic synthesis from Boston University in 2006. Then, he conducted postdoctoral work at Columbia University. In early 2009, he returned to China and started his independent career as a Principal Investigator and the Director of Chemistry Center at National Institute of Biological Sciences (NIBS) in Beijing. In early 2014, he received a tenured professorship from Peking University and moved to the College of Chemistry at Peking University. Now, he is a Full Professor of Chemistry and Chemical Biology and also a senior PI of the Peking-Tsinghua Center for Life Sciences. His major research areas are chemical biology, natural product synthesis, synthetic biology, and drug discovery. To date, he has published more than 100 original research papers, including in “Cell”, “Nature”, and “Science”, and has obtained more than ten approved patents for new drug discovery. He has received many prestigious awards including the 2018 David Ginsburg Award in Israel, the 2017 Tetrahedron Young Investigator Award, the 2017 Swiss Chemical Society Distinguished Lectureship Award, the 2015 Chemical Society of Japan Distinguished Lectureship Award, the 2014 Roche Young

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Investigator Award, the 2013 International Chemical Biology Society (ICBS) Young Chemical Biologist Award, and the 2010 IUPAC Young Chemist Award. Since 2017, he has served as an editor “Bioorganic & Medicinal Chemistry”.