1 Identification and Isolation of Secondary Metabolites From

Identification and Isolation of Secondary Metabolites from Podocarpus neriifolius

Using Bioactivity-Guided and 1D-NMR-Based Dereplication Approaches

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Paule Annécie Benatrehina

Graduate Program in Pharmaceutical Sciences

The Ohio State University

2018

Dissertation Committee

A. Douglas Kinghorn, Advisor

Liva Harinantenaina Rakotondraive, Co-Advisor

James R. Fuchs 1

Copyrighted by

Paule Annécie Benatrehina

2018

Abstract

In a continued effort aimed at the discovery of potential new anticancer leads of natural origin, a root sample of Podocarpus neriifolius D. Don, collected in the Vietnam rainforest, was selected as a candidate species for phytochemical and biological investigation of its bioactive secondary metabolites. An initial small-scale bioactivity- guided isolation of this plant sample was conducted, and yielded one new (83) and four known podolactones (77-79, and 84) together with three totarane-type diterpenes (80-82), with a structural revision carried out on 79. A larger scale extraction of the remaining sample was performed to isolate a greater amount of 83 for its complete structural characterization, as well as to investigate additional secondary metabolites from this plant.

A 1D-NMR spectroscopy-guided fractionation method using both 1H and selective 1D-

TOCSY NMR spectroscopy was developed as a dereplication procedure in the screening of the extracts and their fractions. This method led to the detection and isolation of further known compounds (85-92) from the hexanes, EtOAc, and aqueous extracts of P. neriifolius root sample, in addition to that of the targeted compound (83). Moreover, the 1H NMR profile of the aqueous extract revealed the presence of a major compound (89), corresponding to the glucoside derivative of the cytotoxic 78, and this compound by means of its extract of origin, was subjected to fungal-assisted biotransformation procedures using ii

two Penicillium strains, namely, P. concentricum and P. expansum. A previously reported hydrolysis of compound 89 under harsh chemical conditions led to the A-ring epoxide unit opening of this compound, resulting in an inactive product. The fungal biotransformation proved to be a useful method for the successful hydrolysis of 89 to form 78, and the present study is the first report of a podolactone chemical modification in fungal fermentation cultures. The progress of the biotransformation reaction was monitored periodically using the newly developed 1D-NMR spectroscopic dereplication method, from which both the starting material (89) and product (78) were identified. The obtained isolates were evaluated for their antiproliferative activities against four human cancer cell lines, namely,

HT-29 (colon), MDA-MB-231 (breast), MDA-MB-435 (melanoma), and OVCAR3

(ovarian). In addition, the bioactive and highly abundant inumakilactone A (78) was further evaluated in vivo in a murine xenograft model through a hollow fiber assay. Only compounds 78 and 79 were active against the cell lines used, while 78 did not show significant activity in vivo. Both 78 and 89 were tested in an insect anti-feedant assay, but neither were active.

This dissertation study has opened new avenues for the dereplication in natural product discovery, as well as an additional method for the derivatization of the podolactones, using the 1D-NMR spectroscopic and fungal biotransformation experiments mentioned above, respectively. Finally, although the major compound, 78, did not exhibit in vivo activity against the human cancer subtypes tested, this information constitutes a new contribution to the published literature regarding the podolactone class of compounds.

iii

Dedication

This dissertation is dedicated to both my late father, Telon Jacques Benatrehina (1951-

2008), and my grandmother, Monique Tombo (1940-2017), and to my family, mentors and teachers.

Acknowledgments

Achieving the work described in this dissertation and fulfilling all the prerequisites toward the completion of this doctoral degree have required significant effort, time, and most importantly significant support of various nature, including academic, research, professional, administrative, and personal, from a number of individuals. I would like to include a few words to acknowledge the various parties involved in making this journey a possibility and a success.

First and foremost, I express my deepest gratitude to both my advisors, Prof. A.

Douglas Kinghorn, and Dr. L. Harinantenaina Rakotondraibe, for the tremendous support and understanding they have provided me over the course of my graduate career. Prof.

Kinghorn graciously welcomed me into his research group, and through the years he has entrusted me with various responsibilities within and outside the laboratory, notably as an assistant in the Journal of Natural Products office, all of which have greatly contributed to my scientific and professional growth. I also thank him for his continuous assistance and endorsement for my professional endeavors, including fellowship applications and conference participation. Dr. Rakotondraibe is to be thanked for willingly assuming the role of my co-advisor, and as such for patiently providing me with assistance and direction through my research projects with hands-on coaching and constructive criticism. v

Moreover, I thank him for continuously encouraging me to keep challenging myself and think critically, while stimulating self-confidence. Both Prof. Kinghorn and Dr.

Rakotondraibe have been and continue to be exemplary role models of hard work, integrity, patience and mentorship, making the success of their students a main priority.

I would like to also thank Dr. James R. Fuchs, for his contribution as the third member on both my candidacy and dissertation committee. I am grateful for his valuable advice and constructive feedback regarding my research projects. Additionally, I thank Dr.

Esperanza J. Carcache de Blanco for also being part of my candidacy committee.

I thank the College of Pharmacy Graduate Research Committee and the Division of Medicinal Chemistry and Pharmacognosy, first for my admission into this graduate program, and for the administrative and professional support through career services, workshops and funding opportunities such as the Jack L. Beal and Chang Ahn travel awards, of which I have been privileged to be a recipient. I am also grateful for the many support services offered by The Ohio State University and from which I have greatly benefited, including professional and academic opportunities and coaching through the

OSU Career and Counseling Service.

The U.S. National Cancer Institute, NIH, Bethesda, MD is acknowledged for providing funding for this dissertation research through a program project grant P01

CA125066 awarded to Prof. A. D. Kinghorn as the principal investigator. The interdisciplinary nature of the work presented in this dissertation would not have been feasible without the involvement of many devoted collaborators, to whom I am indebted.

Thus, I would like to acknowledge the valuable contribution of colleagues at the University of Illinois at Chicago (UIC) College of Pharmacy, namely, Dr. Djaja D. Soejarto for leading field collection of plant material, Dr. Joanna E. Burdette, Dr. Wei-Lun Chen, Mr.

Austin Czarnecki, and Mr. Daniel Lantvit, for conducting the biological evaluation data. I thank Dr. Xiaoli Zhang (OSU) for providing her biostatistics expertise on the in vivo bioassay data. Moreover, I thank Dr. Craig McElroy for providing access, training and assistance for the use of various instruments within the College of Pharmacy, as well as

Drs. Arpad Somogyi and Chunhua Yuan of the OSU Campus Chemical Instrument Center

(CCIC) for assistance in acquiring mass spectrometry and NMR spectroscopy data, respectively.

I am thankful for the many colleagues from various laboratories within the College of Pharmacy, including my fellow graduate students, many of whom I now consider good friends, for their help and support, making the challenges of the graduate program more manageable. First, I thank former and current members of the Kinghorn group, namely

Drs. Patrick C. Still, Lynette Bueno Perez, and Yulin Ren, as well as Ms. Andrea L. Rague,

Ms. Garima Agarwal, Mr. Ermias Mekuria Addo, and Mr. Peter J. Blanco-Carcache.

Furthermore, I am grateful to Drs. Hee-Byung Chai, Li Pan, Jie Li, and C. Benjamin

Naman, for being great research mentors, and for providing bioassay data (H.-B. Chai). I also thank former and current colleagues from the Rakotondraibe group, namely, Drs.

Tehane Ali and Masanori Inagaki, Mr. Preston Manwill, and Mr. Choon Yong Tan and

Ms. Fengrui Wang who have conducted biological testing for part of the dissertation

vii

project. In addition, I thank Mrs. Nicole Woodard from the Carcache de Blanco group, as well as Drs. Pratiq Patel and John Woodard, and Ms. Chido Hambira from the Fuchs group.

I am grateful to Mrs. Alice Gardner and Mrs. Jennifer Bartlett, of the College of Pharmacy for their professional assistance and friendship over the years.

I would also like to acknowledge the many mentors from my alma mater, Lipscomb

University, including Drs. Jim Thomas, Kent Gallaher, Kent Clinger, and Susan Mercer, who have encouraged me to pursue graduate study in natural products research.

I am mostly grateful to my family and friends near and far, who have been a great source of encouragement in this long journey. I thank my parents, Telon and Annie

Benatrehina, for their unconditional love and support, and for inspiring me and my siblings, to always strive for excellence through education, ethics and humanity. I also thank the rest of my family, notably my sisters, Annelie and Tellie Benatrehina, for their love and support. Finally, I would like to thank my close friends Estrella and Christopher Shay for their continued support, as well as the Braden, Huston, Haynes, and Moore families, whom have given me a home away from home, for their hospitality, and generosity.

Above all, I thank God, for my life, for the aforementioned, and so much more. I close with the hope and the motivation to contribute to the betterment of life, with the many lessons learned from this experience, science and natural products research, as well as with continuous learning, and sound judgement.

viii

Vita

2007...... Lycée Mixte de Nosy-Be, High School,

Nosy-Be, Madagascar

2008-2012 ...... B.S. Biochemistry, Lipscomb University,

Nashville, TN

2012-2018 ...... Ph.D. Graduate Student, Division of

Medicinal Chemistry and Pharmacognosy,

The Ohio State University, Columbus, OH

Publications

1. Benatrehina, P. A.; Naman, C. B.; Li, J.; Pan, L.; Kinghorn A. D. Uses, biological activity, and potential toxicity of selected botanical dietary supplements consumed in the United States. J. Tradit. Complement Med. 2018, 8, 267-277 (PMCID: PMC5934707) (Invited Review).

2. Li, J.; Yuan, C.; Pan, L.; Benatrehina, P. A.; Chai, H-B.; Keller, W. J.; Naman, C. B.; Kinghorn, A. D. Bioassay-guided isolation of antioxidant and cytoprotective constituents from a Maqui berry (Aristotelia chilensis) dietary supplement ingredient as markers for qualitative and quantitative analysis. J. Agric. Food Chem., 2017, 65, 8634-8642. (PMCID: PMC568550a)

3. Kinghorn, A. D.; Benatrehina, P. A.; Agarwal, G. Natural products. In: Schwab, M. (Ed.) Encyclopedia of Cancer, 4th edition, Springer: Heidelberg, 2017, 1, 3025-3028 (Invited Book Chapter).

4. Naman, C. B.; Benatrehina, P.A.; Kinghorn, A. D. Secondary products: pharmaceuticals, plant drugs, in Encyclopedia of Applied Plant Sciences, 2nd Ed., eds. Thomas B.; Murphy D. J.; Murray B. G., Academic Press: Waltham, MA, 2017, 93-99 (Invited Book Chapter).

5. Ren, Y.; Benatrehina, P. A.; Muñoz Acuña, U.; Yuan, C.; Chai, H-B.; Carcache de Blanco, E. J.; Ninh, T. N.; Soejarto, D. D.; Kinghorn, A. D. Bioactive rotenoids and isoflavonoids from the fruits of Millettia caerulea. Planta Med. 2016, 82: 1096-1104. (PMCID: PMC4956498)

6. Naman, C. B.; Li, J.; Moser, A.; Hendrycks, J. M.; Benatrehina, P. A.; Chai, H-B.; Yuan, C.; Keller, W. J.; Kinghorn, A. D. Computer-assisted structure elucidation of black chokeberry (Aronia melanocarpa) fruit juice isolates with a new fused pentacyclic flavonoid skeleton. Org. Lett. 2015, 17, 2988–2991. (PMCID: PMC4690212)

Fields of Study

Major Field: Pharmaceutical Sciences

Table of Contents

Abstract ...... ii

Dedication ...... iv

Acknowledgments...... v

Vita ...... ix

Table of Contents ...... xi

List of Tables ...... xx

List of Figures ...... xxii

List of Abbreviations ...... xxxii

Chapter 1. Anticancer Agents of Plant Origin ...... 1

A. Cancer statistics ...... 1

B. Relevance of natural products as source of medicine since ancient times ...... 3

C. Overview of the relevance of higher plants in anticancer drug discovery ...... 9

1. The four main classes of anticancer agents ...... 9

1.1. The vinca alkaloids ...... 10

1.2. The podophyllotoxins ...... 12

1.3. The taxanes ...... 14

1.4. The camptothecins ...... 16

2. Additional plant-derived anticancer agents ...... 19

3. The continued role of higher plants as sources of potential anticancer agents ..... 21

Chapter 2. Bioactivity-guided isolation of Podocarpus neriifolius secondary metabolites

...... 23

A. Background on Podocarpus neriifolius D. Don ...... 23

1. Phylogenetic background and ethnobotanical uses of Podocarpus neriifolius..... 23

1.1. The family Podocarpaceae and the genus Podocarpus ...... 23

1.2. Phylogeny of Podocarpus neriifolius ...... 25

1.3. Uses of Podocarpus species...... 28

1.4. Uses of Podocarpus neriifolius ...... 29

2. Phytochemical background of P. neriifolius and the genus Podocarpus ...... 29

2.1. Secondary metabolites isolated from Podocarpus neriifolius ...... 30

2.1.1. Diterpenes isolated from Podocarpus neriifolius ...... 31

2.1.2. Podolactones isolated from Podocarpus neriifolius ...... 32

2.1.3. Flavonoids isolated from Podocarpus neriifolius ...... 35

2.1.4. Additional secondary metabolites from Podocarpus neriifolius ...... 36

2.2. Secondary metabolites isolated from the genus Podocarpus ...... 37

2.2.1. Diterpenes from the genus Podocarpus ...... 37

2.2.2. Podolactones from the genus Podocarpus ...... 40

3. Bioactivity of secondary metabolites from Podocarpus species ...... 44

xii

B. Statement of problem ...... 46

C. Experimental ...... 49

1. General experimental procedures ...... 49

2. Plant material ...... 50

3. Cytotoxicity-guided isolation ...... 51

4. Characterization of the isolated secondary metabolites ...... 54

4.1. Makilactone E (77) ...... 54

4.2. Inumakilactone A (78) ...... 54

4.3. 3-Deoxy-2β-hydroxynagilactone E (79) ...... 55

4.4. Inumakiol D (80)...... 55

4.5. Inumakiol E (81) ...... 55

4.6. 4β-Carboxy-19-nortotarol (82) ...... 62

4.7. Makilactone G (84) ...... 63

5. Biological evaluation ...... 64

5.1. Cytotoxicity against the HT-29 cell line (OSU) ...... 64

5.2. Cytotoxicity against the HT-29, MDA-MB-231, MDA-MB-435, and

OVCAR3 cell lines (UIC) ...... 65

5.3. In vivo assay ...... 65

D. Results and Discussion ...... 65

1. Structure determination of the isolated compounds ...... 66

1.1. Makilactone E (77) ...... 66

xiii

1.2. Inumakilactone A (78) ...... 70

1.3. 3-Deoxy-2β-hydroxynagilactone E (79) ...... 73

1.4. Inumakiol D (80)...... 77

1.5. Inumakiol E (81) ...... 80

1.6. 4β-Carboxy-19-nortotarol (82) ...... 84

1.8. Makilactone G (84) ...... 88

2. Biological activity of the isolated compounds ...... 90

E. Conclusion ...... 92

Chapter 3. 1D-NMR-guided isolation of further constituents from Podocarpus neriifolius and monitoring of podolactone biotransformation in fungal cultures ...... 93

A. Overview of dereplication methods in natural products research ...... 93

1. General dereplication methods in natural products drug discovery ...... 93

1.1. Challenges in natural products drug discovery research and the need for

dereplication methods ...... 93

1.2. Definitions of the term “dereplication”...... 96

1.3. Overview of the methods used in dereplication procedures and types of

dereplication ...... 98

1.4. NMR spectroscopy as a tool for dereplication ...... 100

2. Overview of dereplication methods for plant secondary metabolite isolation.... 101

3. Selective 1D-TOCSY as a tool for natural products dereplication ...... 103

B. Overview of the use of fungal biotransformation in natural product research ...... 104

xiv

C. Statement of problem ...... 107

1. 1D-NMR dereplication-guided purification of P. neriifolius ...... 107

2. Monitoring fungal biotransformation of isolated podolactone with 1D-NMR

approach ...... 109

D. Experimental ...... 111

1. General experimental procedures ...... 111

2. Fractionation and isolation coupled with 1D-NMR spectroscopy ...... 111

2.1. Plant material ...... 111

2.2. Large-scale extraction of the roots of P. neriifolius D. Don ...... 111

2.3. 1H NMR-spectroscopy-directed fractionation of the hexane partition

(AA06795LG.D1) ...... 113

2.4. 1H NMR-spectroscopy-directed fractionation of the aqueous partition

(AA06795LG.D2) ...... 114

2.5. 1H NMR-spectroscopy-directed fractionation of the EtOAc partition

(AA06795LG.D3) ...... 116

2.5.1. Fractionation of AA06795LG.D3 on a reversed-phase C18 column ..... 116

2.5.2. Fractionation of AA06795LG.D3 on Diaion® HP-20 resin ...... 118

2.5.3. Fractionation of AA06795LG.D3 through liquid-liquid partitioning ... 119

2.6. Use of 1D-TOCSY NMR spectroscopy for the localization of the targeted

podolactone glucoside (83) in crude fractions ...... 120

3. Characterization of compounds detected and isolated from NMR spectroscopy-

based dereplication of P. neriifolius ...... 121

3.1. Characterization of nagilactone G-2β-O-β-D-glucoside (83) ...... 121

3.2. Detection of totarol, totaral, and sandaracopimaric acid (85-87) ...... 121

3.3. Characterization of macrophyllic acid (88) ...... 123

3.4. Characterization of inumakilactone A-15-O-β-D-glucoside (89) ...... 123

3.5. Characterization of makilactone F (90) ...... 123

3.6. Characterization of podolactone C (91) ...... 123

3.7. 1-epi-makilactone E (92) ...... 123

4. Biotransformation of inumakilactone A-15-O-β-D-glucoside (89) and its aglycone

(78) in fungal culture ...... 129

4.2. Source of podolactone for fungal-assisted biotransformation ...... 129

4.2.1. Pilot biotransformation of P. neriifolius aqueous fraction

AA06795LG.D2.2 in Penicillium concentricum potato dextrose broth ...... 130

4.2.2. Biotransformation of inumakilactone A (78) from the P. neriifolius

EtOAc-soluble fraction in a P. concentricum potato dextrose broth ...... 131

4.2.3. Biotransformation of the P. neriifolius aqueous extract AA06795LG.D2

in Penicillium concentricum potato dextrose broth ...... 131

4.2.4. Biotransformation of P. neriifolius aqueous fraction AA06795LG.D2 in

Penicillum expansum potato dextrose broth ...... 134

4.3. 1H NMR spectroscopy monitoring of the biotransformation procedure ...... 136

4.4. 1D-TOCSY-NMR-spectroscopy-based structural confirmation ...... 136

xvi

4.5. Identification and isolation of inumakilactone A (78) from fungal

biotransformation ...... 137

5. Biological evaluation ...... 138

5.1. Cytotoxicity assay in a panel of four cancer cell lines ...... 138

5.2. In vivo hollow fiber assay ...... 138

5.3. Insecticidal activity evaluation ...... 139

5.4. Cytotoxicity of fungal fermentation extracts ...... 139

E. Results and Discussion ...... 140

1. Structure elucidation of nagilactone G-2β-O-β-D-glucoside (83) ...... 140

2. Detection and identification of compounds obtained using dereplication methods

...... 145

2.1. 1H NMR-guided fractionation of the P. neriifolius aqueous extract

(AA06795LG.D2) for the identification of glycosidic podolactone derivatives 147

2.1.1. Identification of inumakilactone A-15-O-β-D-glucoside (89) ...... 150

2.2. 1D-NMR-dereplication-guided fractionation of the ethyl acetate-soluble

fraction (AA06795LG.D3) ...... 155

2.2.1. 1H NMR-complemented fractionation of AA06795LG.D3 by reversed-

phase C18 chromatography ...... 155

2.2.2. 1H NMR-spectroscopy and HPLC-guided purification of

AA06795LG.D3Di2 ...... 158

2.2.3. 1D-NMR-dereplication-guided purification of the liquid-partitioned

xvii

fractions of the EtOAc extract ...... 167

2.2.4. Screening of the hexane extract by 1H NMR dereplication ...... 184

3. Monitoring podolactone fungal biotransformation with 1D-NMR spectroscopy194

3.1. Biotransformation in Penicillium concentricum culture ...... 195

3.1.1. Monitoring P. concentricum-assisted biotransformation of

AA06795LG.D2.2 through 1H NMR spectroscopy ...... 195

3.1.2. Monitoring P. concentricum-assisted biotransformation of

AA06795LG.D2 through 1H NMR spectroscopy ...... 201

3.1.3. Monitoring P. concentricum-assisted biotransformation of inumakilactone

A (78) through 1H NMR spectroscopy ...... 212

3.2. Biotransformation in Penicillium expansum culture ...... 214

3.2.1. Monitoring P. expansum-assisted biotransformation of the aqueous extract

AA06795LG.D2 through 1H NMR spectroscopy ...... 215

3.2.2. 1D-NMR spectroscopy for the structural identification of inumakilactone

A (78) as a fungal-assisted biotransformation product ...... 223

4. Biological activity ...... 224

4.1. In vitro cytotoxicity evaluation of components obtained from the large-scale

extraction of P. neriifolius root sample ...... 224

4.1.1. Antiproliferative activity of the main partitions of the methanol crude

extract in a panel of four cancer cell lines ...... 224

4.1.2. Antiproliferative activity of the identified compounds (85-89, 91) from the

xviii

large-scale extraction of P. neriifolius in a panel of four human cancer cell lines

...... 225

4.2. Biological activity of inumakilactone A (78) in vivo...... 227

4.3. Insecticidal potential of inumakilactone A (78) and inumakilactone A-15-O-β-

glucoside (89)...... 227

4.4. Cytotoxic activity of extracts obtained from Penicillium concentricum

biotransformation procedures ...... 227

4.4. Cytotoxic activity of extracts obtained from Penicillium expansum

biotransformation procedures ...... 228

E. Conclusion ...... 233

Bibliography ...... 237

Appendix A: Permission to Reproduce Published Materials ...... 269

Appendix B: HT-29 Cytotoxicity Using an SRB Assay Protocol (OSU) ...... 270

Appendix C: Cytotoxicity Assay Using a Panel of Four Human Cell Lines (UIC) ...... 271

Appendix D: In vivo Evaluation of inumakilactone A (78) in the Murine Hollow Fiber

Assay ...... 272

Appendix E: Cytotoxicity Evaluation of Fungal Biotransformation Fractions against a

Human Ovarian and a Breast Cancer Cell Line Using the SRB Assay ...... 275

Appendix F: Penicillium concentricum Source, Identification, and Culture ...... 277

Appendix G: Penicillium expansum Source, Identification, and Culture ...... 278

xix

List of Tables

Table 1. 1H NMR data of makilactone E (77) and inumakilactone A (78) (400 MHz, in

C5D5N) compared with reported data ...... 56

Table 2. 13C NMR data of makilactone E (77) and inumakilactone A (78) (100 MHz, in

C5D5N) compared with reported data ...... 57

Table 3. 1H and 13C NMR data of 3-deoxy-2β-hydroxynagilactone E (79) (400 and 100

MHz) compared with reported data ...... 58

Table 4. 1H and 13C NMR data data of of inumakiols D (80) (400, 175 MHz) and E (81)

(400, 100 MHz) ...... 60

1 Table 5. H NMR data of inumakiol E (81) (400 MHz, in C5D5N) compared with reported data ...... 62

Table 6. 1H (400 MHz) and 13C (100 MHz) NMR data of 82 and 84 ...... 63

Table 7. Cytotoxic activity of isolated compounds ...... 91

Table 8. Comparison of the 1H and 13C NMR data of nagilactone G-2β-O-β-D-glucoside

(83) and its aglycone (79) ...... 122

Table 9. 1H (400 MHz) and 13C (100 MHz) NMR data of macrophyllic acid (88) ...... 124

Table 10. Comparison of the 1H and 13C NMR data of inumakilactone A-15-O-β-D- glucoside (89) with reported data and inumakilactone A (78) ...... 125 xx

Table 11. 1H NMR data of makilactone F (90) and 1-epi-makilactone E (92) ...... 126

Table 12. 13C NMR data of makilactone F (90) and 1-epi-makilactone E (92) ...... 127

Table 13. 1H and 13C NMR data of podolactone C (91) (400 MHz) ...... 128

Table 14. Biotransformation procedure of AA06795LG.D2 (250 mg) in Penicillum expansum fungal culture ...... 135

Table 15. Antiproliferative activity of fractions from the large-scale extraction of P. neriifolius in a panel of four human cancer cell lines ...... 226

Table 16. Antiproliferative activity fungal biotransformation fractions from two

Penicillium strains against the A2780 human ovarian cancer cell line ...... 230

Table 17. Antiproliferative activity fungal biotransformation fractions from two

Penicillium strains against the MCF-7 human breast cancer cell line ...... 231

xxi

List of Figures

Figure 1. Structures of vinca alkaloids: vinblastine, vincristine, vindesine, and vinorelbine

(1-4)...... 11

Figure 2. Structures of podophyllotoxin and analogs (DEPBD, etoposide, teniposide) (5-

8) ...... 13

Figure 3. Structures of taxanes: paclitaxel, docetaxel, and cabazitaxel (9-11) ...... 16

Figure 4. Structures of camptothecin derivatives (12-15) ...... 18

Figure 5. Additional examples of plant-derived antitumor and cancer-related agents (16-

19) ...... 20

Figure 6. Botanical sketch of Podocarpus neriifolius D. Don ...... 26

Figure 7. Structures of representative diterpenes (20-24) from P. neriifolius ...... 32

Figure 8. Structural classification of the podolactones ...... 33

Figure 9. Representative podolactones (25-29) from P. neriifolius ...... 34

Figure 10. Examples of flavonoids (30-33) from P. neriifolius ...... 35

Figure 11. Structures of additional compounds from P. neriifolius...... 36

Figure 12. Examples of podocarpane (39) and totarane-type (40-49) diterpenes and bisditerpenes from Podocarpus species ...... 39

Figure 13. Examples of sempervirol- (50, 51) and abietane-type (52-56) diterpenes from xxii

Podocarpus species ...... 40

Figure 14. Examples of type-A podolactones (57-64) from Podocarpus species ...... 41

Figure 15. Examples of type-B podolactones (65-71) from Podocarpus species ...... 43

Figure 16. Examples of type-C podolactones (72-76) from Podocarpus species ...... 44

Figure 17. P. neriifolius extraction and partition scheme ...... 52

Figure 18. HRESIMS [(full, (a) and zoomed with prediction (b)] spectra of makilactone E

(77) ...... 67

Figure 19. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of makilactone E (77)

(in C5D5N) ...... 68

Figure 20. HSQC (a) and HMBC (b) spectra of makilactone E (77) (400 MHz, in C5D5N)

...... 69

Figure 21. HRESIMS spectrum of inumakilactone A (78) ...... 70

Figure 22. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of inumakilactone A

(78) ...... 71

Figure 23. HSQC (a) and HMBC (b) spectra of inumakilactone A (78) (in C5D5N) ...... 72

Figure 24. HRESIMS spectrum of 3-deoxy-2β-hydroxynagilactone E (79) ...... 73

Figure 25. 1H (400 MHz) (a) and 13C (175 MHz) (b) NMR spectra of 3-deoxy-2β- hydroxynagilactone E (79) (in C5D5N) ...... 74

Figure 26. HSQC (a) and HMBC (b) spectra of 3-deoxy-2β-hydroxynagilactone E (79) (in

C5D5N) ...... 76

Figure 27. HRESIMS spectrum of inumakiol D (80) ...... 77

xxiii

Figure 28. 1H (400 MHz) (a) and 13C (175 MHz) (b) NMR spectra of inumakiol D (80) in

CD3OD ...... 78

Figure 29. HSQC (400 MHz) (a) and HMBC (700 MHz) (b) spectra of inumakiol D (80) in CD3OD ...... 79

Figure 30. HRESIMS spectrum of inumakiol E (81) ...... 81

13 Figure 31. C NMR spectrum of inumakiol E (81) (100 MHz, in CDCl3) ...... 81

1 Figure 32. H NMR spectra of inumakiol E (81) in CDCl3 (a) and in C5D5N (b) (400 MHz)

...... 82

Figure 33. HSQC (a) and HMBC (b) spectra of inumakiol E (81) (400 MHz, in CDCl3)

...... 83

Figure 34. HRESIMS spectrum of 4β-carboxy-19-nortotarol (82) ...... 84

Figure 35. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of 4β-carboxy-19-nortotarol (82) (in CDCl3) ...... 85

Figure 36. Selective 1D-TOCSY (a) and 1H (b) NMR spectra of 4β-carboxy-19-nortotarol

(82) (400 MHz, in CDCl3) ...... 86

Figure 37. HSQC (a) and HMBC (b) spectra of 4β-carboxy-19-nor-totarol (82) (400 MHz, in CDCl3) ...... 87

Figure 38. HRESIMS spectrum of makilactone G (84) ...... 88

Figure 39. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of makilactone G (84)

(in CD3OD) ...... 89

Figure 40. HSQC (a) and HMBC (b) spectra of inumakiol G (84) (400 MHz, in CDCl3)

xxiv

...... 90

Figure 41. Overview of structural diversity on the B-type podolactone core...... 108

Figure 42. Large-scale solvent extraction and partition scheme of P. neriifolius root sample...... 113

Figure 43. 1H NMR spectroscopy-guided fractionation scheme of the hexanes partition

(AA06795LG.D1) ...... 115

Figure 44. 1H NMR-spectroscopy-guided fractionation of the aqueous partition

(AA06795LG.D2) ...... 117

1 Figure 45. H NMR spectroscopy-guided reversed-phase C18 fractionation scheme of

AA06795LG.D3...... 118

Figure 46. Biotransformation procedure of 250 mg P. neriifolius aqueous extract in

Penicillium concentricum culture ...... 133

Figure 47. HRESIMS spectrum of nagilactone G-2β-O-β-D-glucoside (83) ...... 141

Figure 48. MS/MS spectrum of nagilactone G-2β-O-β-D-glucoside (83) ...... 141

Figure 49. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of nagilactone G-2β-

O-β-D-glucoside (83) ...... 142

Figure 50. 1H (400 MHz) NMR spectrum of nagilactone G-2β-O-β-D-glucoside (83) in

CD3OD ...... 143

Figure 51. HSQC (a) and HMBC (b) spectra of 83 ...... 144

Figure 52. Key HMBC (a) and NOESY correlation for 83 ...... 145

Figure 53. Additional compounds isolated during the large-scale isolation of P. neriifolius

xxv

...... 147

Figure 54. 1H NMR spectroscopic profile of the P. neriifolius aqueous extract

(AA06795LG.D2) ...... 149

Figure 55. 1H NMR spectra of AA06795LG.D2 and its fractions from Diaion® HP-20 resin separation...... 149

Figure 56. 1H NMR profiling of AA06795LGD2 with its first (D2.1) and last fractions

(D2.3) from Diaion® HP-20 resin separation ...... 150

Figure 57. Overlay of the 1H NMR spectra of AA06795LGD2 and its fraction

AA06795LGD2.2...... 151

Figure 58. HRESIMS spectrum of inumakilactone A-15-O-β-D-glucoside (89) ...... 152

Figure 59. HSQC data for inumakilactone A-15-O-β-D-glucoside (89) ...... 153

Figure 60. HMBC data for inumakilactone A-15-O-β-D-glucoside (89) ...... 154

Figure 61. 1H NMR spectrum of fraction AA06795LG.D3.1 including key podolactone core signals...... 157

Figure 62. Overlay of 1H NMR spectra of AA06795LG.D2.2 and AA06795LG.D3.1.5

(89) ...... 157

Figure 63. Overlay of 1H NMR spectra of AA06795LG.D3.1.2 and 78 ...... 158

Figure 64. Overlay of 1H NMR spectra of AA06795LG.D3 and its Diaion® HP-20 fractions...... 159

Figure 65. HPLC trace of fraction AA06795LG.D3.Di2 and UV pattern of selected peaks.

A: (89); B: (90); C: (84); D: (78)...... 160

xxvi

Figure 66. Determination of AA06795LG.D3.Di2.H7 (89) by NMR spectroscopic dereplication ...... 161

Figure 67. Determination of AA06795LG.D3.Di2.H10 (84) by NMR spectroscopic dereplication ...... 161

Figure 68. Determination of AA06795LG.D3.Di2.H14 (78) by NMR spectroscopic dereplication ...... 162

Figure 69. Overlay of 1H NMR spectra of AA06795LG.D3.Di2.H8 (90) and

AA06795LG.D3.Di2.H10 (84) ...... 164

Figure 70. HRESIMS spectrum of makilactone F (90) ...... 166

1 Figure 71. H NMR spectrum of makilactone F (90) (700 MHz in C5D5N) ...... 166

13 Figure 72. C NMR spectrum of makilactone F (90) (175 MHz in C5D5N) ...... 166

Figure 73. HSQC spectrum of makilactone F (90) (in C5D5N) ...... 167

Figure 74. Comparison of the HPLC traces of the CHCl3 (a) and aqueous (b) partitions of the EtOAc extract with that of the original glucoside-containing fraction (c) ...... 168

Figure 75. HRESIMS spectrum of podolactone C (91) ...... 169

Figure 76. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of podolactone (91)

...... 170

Figure 77. HSQC (a) and HMBC (b) spectra of podolactone (91) ...... 171

Figure 78. HRESIMS of 1-epi-makilactone E (92) ...... 173

Figure 79. 1H NMR spectrum (a) of and key NOESY correlations (b) for 1-epi- makilactone E (92) ...... 174

xxvii

Figure 80. HSQC (a) and HMBC (b) spectra of 1-epi-makilactone E (92) ...... 175

Figure 81. Overlay of 1H (a) and 1D-TOCSY NMR (b) spectra of 83 ...... 177

Figure 82. 1D-TOCSY NMR experiment for P. neriifolius EtOAc extract in CD3OD 178

Figure 83. 1D-TOCSY NMR spectra for P. neriifolius EtOAc extract ...... 179

Figure 84.1D-TOCSY NMR spectra for the aqueous fraction (AA06795LG.D3.W) ... 180

Figure 85. 1D-TOCSY NMR spectra for the CHCl3 fraction (AA06795LG.D3.C) ..... 181

Figure 86. Overlay of the 1H NMR spectra of (a) 83 and (b) that of peak 2 from

AA06795LG.D3.W ...... 182

Figure 87. Overlay of 1H NMR (a) and 1D-TOCSY (b) NMR spectra of AA06795LG-

D3.F1 with that of 83 (c) (zoomed 1.30-3.00 ppm region)...... 183

Figure 88. 1H NMR spectroscopic profile of the hexane extract (AA06795LG.D1) ... 186

Figure 89. Overlay of 1H NMR spectrum of the hexane extract (AA06795LG.D1) with that of compounds 82, 87, and 88...... 186

Figure 90. HRESIMS spectrum of macrophyllic acid (88) ...... 187

Figure 91. 1H (a) (400 MHz) and 13C (b) (100 MHz) NMR spectra of macrophyllic acid

(88) (Measured in CDCl3 at 300 K) ...... 188

Figure 92. HSQC (a) and HMBC (b) spectra of macrophyllic acid (88) ...... 189

Figure 93. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of totarol (85)- containing sub-fraction ...... 190

Figure 94. HSQC (a) and HMBC (b) NMR spectra of totarol (85)-containing sub-fraction

...... 191

xxviii

Figure 95. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra totaral (86)-containing sub-fraction ...... 192

Figure 96. HSQC (a) and HMBC (b) NMR spectra of totaral (86)-containing sub-fraction

...... 192

Figure 97. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra sandaracopimaric acid

(87)-containing sub-fraction ...... 193

Figure 98. HSQC and HMBC NMR spectra of sandaracopimaric acid (87)-containing sub- fraction ...... 194

Figure 99. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-4d-W, (b) 89 and, (c) 78

...... 197

Figure 100. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-6d-W, (b) 89 and, (c) 78

...... 198

Figure 101. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-6d-E, (b) 89 and, (c) 78

...... 199

Figure 102. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-2d-W, (b) PN-D2.2-PC-

II-2d-W, (c) 89 and, (d) 78. Measured in CD3OD, 400 MHz...... 202

Figure 103. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-2d-W, (b) PN-D2.2-PC-

I-3d-W Measured in CD3OD, 400 MHz ...... 203

Figure 104. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-2d-E, (b) PN-D2.2-PC-

I-3d-E ...... 204

Figure 105. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-3d-E, (b) PN-D2.2-PC-

xxix

I-6d-E, and (c) PN-D2.2-PC-II-6d-E ...... 204

Figure 106. Overlay of the 1H NMR spectra of (a) PN-D2-PC-2d-E, (b) 89, and (c) 78

...... 206

Figure 107. Overlay of the 1H NMR spectra of (a) PN-D2-PC-2d-W and (b) PN-D2-PC-

6d-W ...... 207

Figure 108. Overlay of the 1H NMR spectra of (a) 89 and (b) 78 ...... 209

Figure 109. Detection of 89 and 78 in PN-D2-PC-6d-E fraction by 1D-TOCSY NMR spectroscopy ...... 210

Figure 110. Overlay of the 1H NMR spectra from the biotransformation of 78 in P. concentricum ...... 214

Figure 111. Overlay of 1H NMR spectra of the EtOAc partitions from flasks I and IV at days 2 and 3...... 216

Figure 112. Overlay of 1H NMR spectra of the ETOAc partitions from flasks I and IV at days 2 and 3...... 217

Figure 113. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-I-2d-W and (b) 89 . 218

Figure 114. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-I-7d-W, (b) 89, and (c)

78...... 219

Figure 115. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-14d-W and (b) 89 .. 221

Figure 116. Detection of 79 as biotransformation product in PN-D2-PE-2g-14d-E by 1H

NMR spectroscopy...... 221

Figure 117. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-14d-EP, (b) PN-D2-PE-

xxx

2g-15d-EP, and (c) 78 ...... 222

xxxi

List of Abbreviations

1D, 2D: one-dimensional, two-dimensional br: broad (NMR splitting)

C5D5N: deuterated pyridine

CDCl3: deuterated chloroform

CD3OD: deuterated methanol

CH2Cl2: dichloromethane

CH3Cl: chloroform

CH3CN: acetonitrile d: doublet

D1, D2, D3, D4: hexanes, aqueous, ethyl acetate, and acetone extracts, respectively

δC: carbon-13 chemical shift (in parts per million, ppm)

δH: proton chemical shift (in parts per million, ppm)

DMSO: dimethyl sulfoxide

EtOAc: ethyl acetate

HMBC: heteronuclear multiple bond correlation spectroscopy

HPLC: high-performance liquid chromatography

HRESIMS: high resolution electrospray ionization mass spectrometry xxxii

HSQC: heteronuclear single quantum coherence spectroscopy

Hz: Hertz

IC50: concentration required to inhibit 50% cell growth compared to untreated control

J: coupling constant kg: kilogram

λ: wavelength in nanometer

LC: liquid chromatography

LG: large scale

MeOH: methanol mg: milligram min: minute m/z: mass to charge ratio nm: nanometer

NMR: nuclear magnetic resonance

NOESY: nuclear Overhauser enhancement spectroscopy s: singlet

SPE: solid phase extraction

TLC: thin-layer chromatography

μg: microgram

μM: micromolar

UV: ultraviolet

xxxiii

Chapter 1. Anticancer Agents of Plant Origin

A. Cancer statistics

Cancer continues to affect millions worldwide, constituting a major public health burden and being one of the primary causes of death globally. In the year 2012, over 14 million members of the global population were estimated to have contracted cancer, and

8.2 million deaths were associated with this group of diseases (Torre et al., 2015). In the

United States, cancer is the second-leading cause of death, with more than 1.7 million new cases expected this year (2018), and a mortality rate estimated to reach nearly 1,700 deaths per day, surpassed by accidents in children and heart diseases in adults (Ward et al., 2014;

Siegel et al., 2018). However, the last few decades have witnessed a decline in the overall cancer death rate in the U.S., with a 25% drop from 1991 to 2014 (Siegel et al., 2017). This positive outcome has been attributed mainly to reduced tobacco use, and the access to early detection and to advanced treatment protocols (Siegel et al., 2017). While similar trends have been observed in other developed countries, a heavier cancer burden is projected to shift towards less economically developed countries due to the adoption of behaviors favoring the risk factors of cancer, the prevalence of cancer-causing infections, and the limited adherence to and availability of preventive and therapeutic modalities, such as vaccination and chemotherapy (Torre et al., 2015). In addition, the growth and aging of the 1

population in these countries are contributing to this alarming drift (Torre et al., 2015).

Cancers of the liver, stomach and cervix are the most common in less developed countries, in contrast to lung, colorectal, prostate, and female breast cancers in more developed countries (Torre et al., 2015). While the number of deaths attributed to these four major cancers (lung, prostate, breast, and colorectum) has declined in the U.S., liver, uterine and pancreatic cancer death rates have continued to rise for the past few years (Siegel et al.,

2017). Continuous efforts in cancer prevention, in addition to advances in early detection and treatment approaches have improved the life expectancy for patients affected by certain cancer subtypes. However, independently of the therapeutic procedure employed, this remission is often short-lived due to tumor recurrence, and the relapse is commonly accompanied with resistance to conventional therapies and metastasis (Viale & Draetta,

2016). Additionally, specific therapies for certain cancers, such as solid tumors, remain limited, thus leading to poor prognosis, especially when diagnosed at advanced stages

(Benson, 2007; Bailly, 2009). Another challenge facing cancer therapy as is the case with many other disease conditions is the prevalence of therapy-induced side effects, which in some cases can lead to severe long-term life-altering conditions including chronic pain, impaired fertility, and even an increased risk to develop other cancers (Miller et al., 2016).

Therefore, a tremendous effort remains to be undertaken in the fight against cancer and its related collateral conditions. Among these efforts are the development of enhanced detection methods and accessibility, as well as continuous research for improved treatment, including chemotherapy approaches with increased safety and efficacy against resistant

chemotypes (Ismael et al., 2008; Siegel et al., 2017).

B. Relevance of natural products as source of medicine since ancient times

For millennia, Nature has been a source of inspiration for humankind to meet basic needs, from clothing and shelter to food and medicine (Newman et al., 2000). The long history of use of natural resources for healing purposes can be traced from the early cuneiform records of Mesopotamia around 2600 BC, the Egyptian “Eber papyrus” of 1500

BC, to the Chinese Materia Medica and the Indian Ayurveda of 1100 and 1000 BC, respectively (Newman et al., 2000). These were then followed by the contribution of the

Greeks, including Theophrastus, Dioscorides, and Galen, then the expansion by the Arabs, which eventually led to the more contemporary Pharmacopeia, such as that published in the United Kingdom in the early 1600s (Newman et al., 2000). Although plants constitute the most common folk medicine ingredients in many civilizations from ancient eras to current times, other organisms have been employed, and examples include the fungus

Peptoporus betulinus used as antiseptic and disinfectant and lichens for treatment of inflammations and scalp diseases (Dias et al., 2012).

In its first Traditional Medicine Strategy (2002-2005), the World Health

Organization (WHO) predicted that traditional medicine constitutes the primary health care source for an estimated 80% of the world’s population (Newman et al., 2000; World Health

Organization, 2002), and, more recently, the updated version of this Strategy underlined the continuous growth in popularity of traditional and contemporary medicinal practices

both in developing and more economically developed countries (World Health

Organization, 2013). As one of the main components of such practices, natural products, therefore, represent an important source of traditional means of healing.

Additionally, Nature, by affording Western conventional medicine with its wealthy collection of secondary metabolites, constitutes an essential source of compound leads for prescription drugs against an array of illnesses (David et al., 2015). Among the earliest- used bioactive constituents of natural origin were morphine, atropine, and colchicine, isolated from the plant species Papaver somniferum (Papaveraceae), Atropa belladonna

(Solanaceae), and Colchicum autumnale (Colchicaceae), respectively (Newman et al.,

2000). Several reviews have been published detailing the significant contribution of natural products as approved therapeutic agents, clinical candidates, or leads thereto, for the treatment of a plethora of human disease conditions. Among these are the notable series of reports by Newman and Cragg over the years (Cragg et al., 1997; Newman et al., 2003;

Newman & Cragg, 2007; Newman & Cragg, 2012, 2016), with the latest including a survey of all approved drugs between the 1981-2014 period. Reports by Butler and co-workers citing the proportion of natural products and their derivatives in clinical trials further demonstrate the continuous relevance of secondary metabolites in the pharmaceutical industry (Butler, 2004; Butler et al., 2014). Some of these drug candidates have now been launched, including the microorganism-derived antineoplastic agent and a staurosporine derivative, midostaurin (Rydapt®), approved in April 2017 for the treatment of FLT3- positive acute myeloid leukemia (AML) and systemic mastocytosis (Butler et al., 2014;

Food and Drug Administration, 2017; Levis, 2017).

According to the above reports, while natural products have provided chemotherapeutic agents and leads against a variety of human diseases, their impact has been most predominantly in the areas of infections and cancers. As such, of the 112 small- molecule antibacterial agents approved in the years 1981-2014, 73% were either unmodified natural products or their derivatives (Newman & Cragg, 2016). Similarly, as described in the same account, when one considers the 136 antineoplastic small molecules approved in that time period, over 80% are either natural products, derivatives therefrom, or their mimics. In contrast, secondary metabolites have not been well-represented in the antifungal arena, as most of such agents are of synthetic origin, with the rather “ancient” agents, such as griseofulvin and amphotericin, both launched in the late 1950s, still involved in current treatment regimen (Newman & Cragg, 2016). Nonetheless, newer examples of naturally based antifungal agents include the pneumocandin-derived capsofungin, and the related micafungin (FK463) (Baker et al., 2007; Fujie, 2007).

When the natural origins of the above-mentioned approved therapeutic agents are examined, microbes and plants constitute the principal source organisms, as these have been the most predominantly accessible study material in the past. Indeed, spurred by the discovery of penicillin, a “Golden Era” of natural products, mainly of microbial origin, was born. While these chemotherapeutics are now considered “old drugs”, they continue to serve as important scaffolds to the synthesis of thousands of derivatives in recent years

(Newman & Cragg, 2014), thus further enforcing the sustained importance of natural

sources for drug discovery. Examples include tigecycline (a derivative of the tetracycline doxycycline), televancin (a glycopeptide derived from vancomycin), and the cephalosporin-derived ceftaroline fosamil, approved in 2005, 2009, and 2011, respectively

(Newman & Cragg, 2014). Higher plants, previously described as “the sleeping giant of drug development” (Farnsworth & Morris, 1976), have also constituted a valuable source of approved drugs as well, particularly considering the four main classes of antitumor agents described below. The significant roles of plants and microorganisms were again demonstrated in perhaps one of the most important examples of appreciation of the field of pharmacognosy, when the 2015 Nobel Prize in Physiology and Medicine jointly awarded to William C. Campbell and Satoshi Omura, and to Youyou Tu, in recognition of their discoveries of the antiparasitic (river blindness and elephantiasis) microbial drug ivermectin (an avermectin analog) and the antimalarial plant metabolite artemisinin, respectively (Shen, 2015).

Since the isolation of sponge metabolites, spongouridine and spongothymidine, the approval of several marine-based therapeutics has occurred, including the dolastatin 10- derived monomethyl auristatin E (MMAE) as component of the drug antibody conjugate

(ADC) brentuximab vedotin (Adcetris®), the halichondrin B analog eribulin mesylate

(Halaven®), the marine snail isolate ziconotide (Prialt®), and the tunicate product trabectedin (Cragg & Newman, 2013; Harvey et al., 2015) . While medicines from the sea have not yet been established significantly in terms of quantity among currently approved drugs, considerable advances have been and continue to be undertaken in the field of

marine natural product research, as shown by the vast number, extensive chemical diversity, and unprecedented classes of bioactive marine secondary metabolites reported to date. For instance, 840 and 1003 new structures were reported from marine sources in the years 1998 and 2010, respectively (Faulkner, 2000; Blunt et al., 2012; Cragg & Newman,

2013), and the voluminous publication compilations on the bioactive compounds from the ocean continue to attest to the great potential offered by organisms from this source.

Sources belonging to the Animal Kingdom have also supplied small molecules that have acted as leads towards approved therapeutics or the development of novel drug candidates against human diseases. This was the case of captopril and enalapril, designed from teprotide, obtained from the venom of Bothrops jaracaca (pit viper) for the treatment of cardiovascular conditions, and that of exendin-4, isolated from the Gila monster

(Heloderma suspectum) venom, from which the antidiabetic extenatide polypeptide

(Byetta®) was derived (Cragg & Newman, 2013). Moreover, epibatidine, produced by the poisonous frog Epipedobates tricolor led to a novel class of potential analgesic compounds

(Cragg & Newman, 2005a).

The above examples, while representing only a glimpse of the considerable previous contributions of Nature towards the discovery and development of chemotherapeutics, also give a foretaste of the tremendous opportunities that natural products will continue to offer in the future. First, as introduced above, “old drugs” have constituted the basis of more recent synthetic drugs and drug candidates, including many antibiotics being chemically modified for further development (Newman & Cragg, 2014).

Additional repurposing aspects of secondary metabolites can be seen through the evaluation of known compounds against new biological targets, including their use as toxic warhead components of ADC drugs, as was the case of Adcetris® (dolastatin 10 analog),

Mylotarg® (gemtuzumab ozogamicin, calicheamicin based), and ado-trastuzumab emtansine (Kadcyla®, a maytansine derivative), and these demonstrate a promising new horizon for those natural product leads previously limited by their narrow therapeutic indices (Cragg et al., 2014).

Moreover, it has been stated that only a small portion of the estimated known living organisms have been explored to date, including both terrestrial and marine sources, leaving “a virtually untapped reservoir of novel drugs awaiting imaginative and progressive organizations” (Cragg et al., 1997). In particular, microorganisms, identified and currently unknown (not yet discovered), hold great promise in this regard as an unbounded source of novel chemical entities. These include extremophiles, endophytes, and microbes for which artificial culture is not yet facile, all of which remaining to be investigated (Cragg & Newman, 2005b; Wilson & Brimble, 2009). Perhaps two of the most important innovations in the exploration of the potential of these organisms are heterologous gene expression circumventing the production limitation in the original producing source, and the genetic manipulation of “dormant” gene clusters awaiting activation toward the production of new or targeted molecules, and offering deeper understanding in the biosynthetic pathways involved (Cragg & Newman, 2005b; Newman

& Cragg, 2014). Consequently, it has been predicted that microbes, their commensals, and

other symbiotic interactions with their hosts will constitute a great source of novel agents of natural origin in the future (Newman & Cragg, 2014).

Finally, another advantage of natural products compared to other sources such as combinatorial and synthetic chemistry is demonstrated by the degree of similarity they share with approved chemotherapeutic agents, making them a privileged class of compounds for drug discovery (Lam, 2007; El-Elimat et al., 2012). Natural products occupy a much larger region of the chemical space, also termed “biologically relevant chemical space”, further asserting of their superiority over synthetic sources both in regarding their chemical diversity and drug-like properties (Feher & Schmidt, 2003;

Atanasov et al., 2015).

C. Overview of the relevance of higher plants in anticancer drug discovery

1. The four main classes of anticancer agents As the work discussed in this dissertation is centered primarily on the investigation of potential antiproliferative agents from a plant, the following section will provide a brief overview of clinically used plant-derived anticancer agents, and the continued value of this natural source for future antitumor drugs. The classic antineoplastic agents of plant origin are categorized into four main structural groups, namely, the bisindole (vinca) alkaloids, the epipodophyllotoxins, the taxanes, and the camptothecins (Newman et al., 2000;

Balunas & Kinghorn, 2005; Bueno Pérez et al., 2014).

1.1. The vinca alkaloids The pioneering plant-derived anticancer compound class was the vinca alkaloids, so-called by being based on the previous taxonomy of their plant source, Catharanthus roseus (L.) G. Don (formerly Vinca rosea) (Apocynaceae), also commonly known as the

Madagascar periwinkle (Newman et al., 2000; Cragg & Newman, 2005a; Roussi et al.,

2012). Although this plant is endemic to this named country, the first isolations of the parent antitumor bisindole alkaloids, vinblastine (1) and vincristine (2) (Figure 1), were performed on plant specimens collected in Jamaica and the Philippines (Cragg & Newman,

2005a). Owing to its folkloric medicinal usage as a treatment of diabetic conditions, the leaf extracts of C. roseus were investigated initially for their glucose level-lowering constituents, when observations of the effects on white blood cells steered subsequent research towards the antitumor agents later introduced into the clinic (Newman et al., 2000;

Roussi et al., 2012). The principal mechanism of action of these agents is concentrated on the disruption of microtubule assembly through interaction with tubulin on a specific binding domain (the vinca domain), which then results in cell cycle arrest followed by apoptosis (Grothaus et al., 2010; Roussi et al., 2012). These secondary metabolites, still among the widely used cancer chemotherapeutic agents over five decades later, have been studied extensively and continue to be the further developed to ameliorate their pharmacokinetic properties. Hence, additional semi-synthetic vinca-derived analogs have joined this class of the antineoplastic armamentarium, including vindesine (3), vinorelbine

(4), and they are used mostly in combination with other therapies for the treatment of

several cancers, such as lymphomas, leukemias, and breast, lung, and testicular cancers

(Grothaus et al., 2010) (Figure 1). More recently, a vincristine sulfate liposome injection

(Marqibo®) was introduced for the treatment of adult advanced Philadelphia chromosome- negative acute lymphocytic leukemia (Silverman & Deitcher, 2013; Bueno Pérez et al.,

2014). Prompted by the initial discovery and approval of the vinca alkaloids, efforts to search for clinically relevant phytochemicals were undertaken by several organizations, including the U.S. National Cancer Institute (NCI), which later resulted in the introduction of two of the remaining anticancer chemotherapeutic classes discussed below.

Figure 1. Structures of vinca alkaloids: vinblastine, vincristine, vindesine, and vinorelbine

(1-4)

1.2. The podophyllotoxins Plants of the genus Podophyllum, such as P. peltatum L. (American mandrake or

Mayapple) and Podophyllum emodi Wallich (Berberidaceae), have been known for their usage as cathartics and cholagogues in Native North American and Himalayan traditional medicine, as well as for the treatment of skin cancer and warts (Newman et al., 2000; Lee

& Xiao, 2012). The isolation of the podophyllotoxin lignans was instigated by the discovery of the biological properties of podophyllin, the alcoholic extract of Podophyllum rhizomes, as it restored benign genital warts (condylomata acuminata) and later proved toxic to mitotic cells (Lee & Xiao, 2012). Following phytochemical investigation of the plant, podophyllotoxin (5, Figure 2) was isolated, and it was shown to interfere with the mitotic spindle assembly in a similar manner to the vinca alkaloids (Grothaus et al., 2010;

Lee & Xiao, 2012). However, podophyllotoxin clinical development was aborted due to insufficient efficacy and dose-limiting toxicity, notably gastrointestinal adverse effects

(Lee & Xiao, 2012). Then, extensive efforts were conducted leading to the discovery of the epipodophyllotoxin glycoside analogs (e.g., 4′-O-demethyl-epipodophyllotoxin- benzylidene β-D-glucoside, DEPBD, 6, Figure 2), in addition to semi-synthetic modifications, resulting in the now clinically used etoposide (7) and teniposide (8) (Figure

2) (Lee & Xiao, 2012). Surprisingly, with a slight structural change, these analogs exhibited increased therapeutic effectiveness as well as a mechanism of action different than that of the closely-related podophyllotoxin, as they prohibited cell division by inhibiting topoisomerase II, leading to DNA breakage and cell death (Stähelin & von Wartburg, 1991;

Grothaus et al., 2010; Lee & Xiao, 2012).

Figure 2. Structures of podophyllotoxin and analogs (DEPBD, etoposide, teniposide) (5-

Thus, the development of the clinically used podophyllotoxin analogs exemplifies the lengthy process of drug discovery, which also may involve serendipitous events.

Indeed, over a century passed from the first isolation of podophyllotoxin lignans in 1880

to the FDA approval of the semi-synthetic derivatives etoposide and teniposide in 1983 and 1992, respectively (Lee & Xiao, 2012). In addition to these two drugs, etoposide phosphate is clinically available, and together they are used against a variety of cancers, including bronchial and testicular cancers, leukemia, and Kaposi’s sarcoma (Cragg &

Newman, 2005a; Lee & Xiao, 2012).

1.3. The taxanes Paclitaxel (Taxol®, 9, Figure 3), an amino acid-containing diterpenoid, was first isolated from the alcoholic extract of the stem bark of Taxus brevifolia Nutt. (Pacific yew)

(Taxaceae) in 1966, but its complex structure was only reported in 1971 owing to the remarkable contribution of Drs. Wall and Wani and their colleagues (American Chemical

Society; Wani et al., 1971). Although a highly promising compound during those early years, the initiation of paclitaxel into preclinical studies was hampered by several limitations, notably its low yield in the plant source and its lack of water solubility

(Kingston, 2012). However, the significant potency of paclitaxel in in vivo murine models such as CX-1 (colon) and MX-1 (breast) human tumor xenografts allowed for its continued investigation (Oberlies & Kroll, 2004; Kingston, 2012). Moreover, a crucial discovery by

Dr. Susan Band Horwitz and co-workers on the mechanism of action of paclitaxel through an unprecedented interaction with tubulin further rekindled interest in more advanced clinical studies (Schiff et al., 1979; Kingston, 2012). Through a unique tubulin-binding property, paclitaxel promotes and stabilizes microtubule formation (tubulin

polymerization), forcing the cancer cell to undergo DNA replication without cytokinesis, thus resulting in endoreduplication and eventually to cell death by apoptosis (Oberlies &

Kroll, 2004). Moreover, with the development of an improved formulation in castor oil

(Cremophor EL®), paclitaxel, although it elicited allergic responses, was successfully introduced into the market for the treatment of refractory ovarian and breast cancers, in

1992 and 1994, respectively (Kingston, 2012). With a continuing supply problem that was related to paclitaxel, then obtainable only from T. brevifolia bark, the discovery of an alternative source was most welcomed. Potier and his co-workers not only uncovered the taxane 10-deacetylbaccatin III (10-DAB) being produced in high abundance in the needles of the European yew Taxus baccata, but, by using 10-DAB, they semi-synthesized the second-generation taxane anticancer drug, docetaxel (10, Figure 3), approved for advanced breast cancer in 1996 and non-small cell lung cancer (NSCLC) three years later (Oberlies

& Kroll, 2004; Cragg & Newman, 2005a; Kingston, 2012). Cabazitaxel (11, Figure 3), another semi-synthetic paclitaxel derivative, exhibited lower affinity for the multidrug resistance protein P-glycoprotein (P-gp, efflux pump) and potency against both docetaxel- sensitive and resistant cancers, and was approved in 2010 to treat hormone-refractory metastatic prostate cancer (Galsky et al., 2010). Additional paclitaxel-based chemotherapeutics, inclusive of paclitaxel polyglomex, and the nanoparticle preparations,

AbraxaneTM (albumin-bound paclitaxel) and Genoxol-PMTM (polymeric micelle formulation), have now entered the clinic for the treatment of various malignancies, such as NSCLC, breast and pancreatic cancers (Bueno Pérez et al., 2014; Hare et al., 2017).

1.4. The camptothecins In another noteworthy contribution of the Wall-Wani team, a second class of anticancer agents was named after the parent alkaloid, camptothecin (12, Figure 4), the isolation of which was first published in 1966 from the bioactivity-guided isolation of a stem wood sample of the Chinese native “happy tree”, Camptotheca acuminata Decne.

(Nyssaceae), grown in the United States (Wall et al., 1966; Wall & Wani, 1996).

Figure 3. Structures of taxanes: paclitaxel, docetaxel, and cabazitaxel (9-11)

Camptothecin and its derivatives are characterized structurally by a pentacyclic core composed of quinoline, pyrrolinic-δ-lactam, and α-hydroxy group-bearing-δ-lactone

moieties defined as rings A-B, C-D, and E, respectively (Martino et al., 2017). While preclinical development of 12 showed great promise, early clinical studies were hampered by insufficient efficacy accompanied by substantial side effects related to the poor solubility and stability of the drug candidate in its salt form (Wall & Wani, 1996; Oberlies

& Kroll, 2004). Thus, clinical development of camptothecin was temporarily discontinued until the discovery of its mechanism of action was reported by Hsiang and colleagues in

1985. This compound was found to bind the DNA-replicating enzyme topoisomerase I, blocking religation of the cleaved DNA single strand leading to irreversible DNA cleavage resulting in downstream cell death (Hsiang et al., 1985). This discovery revived the development of more clinically amenable camptothecin analogues as topoisomerase I inhibitors with improved water solubility and manageable toxicity. A decade later, topotecan (HycamtinTM, 13, Figure 4), a semi-synthetic (9-[dimethylamino)methyl]-10- hydroxy-camptothecin, became the first U.S. FDA-approved camptothecin derivative, prescribed for the treatment of relapsed ovarian cancer (Pizzolato & Saltz, 2003; Rahier et al., 2012). Topotecan has since been approved for the treatment of additional cancers such as cervical and small-cell lung cancers, as second-line chemotherapy and in combination regimens, and can be administered intravenously or orally (Rahier et al., 2012). Shortly following the approval of topotecan, irinotecan (CamptosarTM, 14, Figure 4) was approved in the US for the treatment of metastatic colorectal cancer. Structurally, irinotecan has been characterized as a prodrug, as its dipiperidine moiety is cleaved by hepatic and gastrointestinal carboxylesterases releasing the more potent metabolite SN38 (Pizzolato &

Saltz, 2003). The indications for irinotecan have been extended to other cancers including breast, gastrointestinal, cervical, ovarian, as well as lymphomas, as a component of combination therapies (Rahier et al., 2012). A third semi-synthetic analog with a 7- isopropylaminoethyl attachment, belotecan (CamptobellTM, 15, Figure 4), is currently only approved in South Korea for the treatment of ovarian and small-cell lung cancers (Rahier et al., 2012; Shang et al., 2017).

Figure 4. Structures of camptothecin derivatives (12-15)

Comparable to the previous antineoplastic agents described above, continuous

clinical development studies on the camptothecin derivatives are undergoing, focusing on improved formulation and the delivery of new and approved analogs to expand the therapeutic indices, circumvent resistance, and broaden the application of compounds of this class to additional malignancies.

2. Additional plant-derived anticancer agents In addition to the four chemotherapeutic classes mentioned above, higher plants have been the source of new agents, of which three used as antineoplastic or related drugs will be mentioned briefly. In 2012, two plant-derived drugs, namely, omacetaxine mepesuccinate (17, Synribo®) and ingenol mebutate (18, Picato®) (Figure 5) were approved, respectively, for the treatment of chronic myeloid leukemia (CML) and actinic keratosis, a common precursor of squamous cell carcinoma (Lebwohl et al., 2012; Alvandi et al., 2014; Bueno Pérez et al., 2014). The former is a derivative of the naturally occurring compound, homoharringtonine (16, Figure 5), first isolated from the Chinese plant

Cephalotaxus harringtonia (Knight ex J. Forbes) K. Koch (Cephalotaxaceae) (Powell et al., 1970). Omacetaxine mepesuccinate exerts its antineoplastic activity by preventing oncoprotein expression through ribosomal interference (Gandhi et al., 2014). Ingenol mebutate (ingenol-3-angelate), which is applied topically for the indication mentioned above, was isolated as an active constituent of the sap of Euphorbia peplus L.

(Euphorbiaceae), a plant used in Australian folkloric treatment of skin cancers (Hampson et al., 2005; Cragg et al., 2014). Furthermore, this compound promotes the activation of

the protein kinase C and leads to apoptosis (Ogbourne et al., 2004; Hampson et al., 2005).

Moreover, ado-trastuzumab emtansine (T-DM1, 19, Figure 5), an antibody-drug conjugate (ADC) composed of the antibody trastuzumab (Herceptin) and a maytansine- derivative as the warhead was introduced in 2013 for the treatment of Her-2 positive metastatic breast cancer (Ornes, 2013; Amiri-Kordestani et al., 2014).

Figure 5. Additional examples of plant-derived antitumor and cancer-related agents (16-

19)

Maytansine, initially reported from Maytenus species and Putterlickia verruosa

(Celastraceae) (Kupchan et al., 1972), was later shown to be biosynthesized by endophytes associated with the plants (Wings et al., 2013; Kusari et al., 2014). Although the maytansinoids proved too toxic for continued clinical trials, and thus were abandoned, this class of compounds was later resurrected through the development of the more selective

ADCs, which extended their efficacy and safety (Cassady et al., 2004; Cragg et al., 2014).

3. The continued role of higher plants as sources of potential anticancer agents The examples presented in the above sections underline the major contribution of higher plants as initial sources of potent chemotherapeutic agents used to treat various cancer types. For the reasons stated previously, plants will also continue to be a significant source of future potential agents, including those aimed at the treatment of cancer, as numerous drug candidates derived from higher plants and their secondary metabolites are currently in preclinical and clinical studies (Cragg & Newman, 2005a; Grothaus et al.,

2010; Kinghorn et al., 2011). Moreover, while several of the nominally plant-derived leads were later found to be produced by endophytes or the microbial interaction with the plant host, the discovery of these leads would have been delayed, if not missed, without the investigation of their “plant hosts” (Grothaus et al., 2010). Likewise, unprecedented biological findings originated from the discovery of these compounds through the study of their mode of action. The cases of taxol and camptothecin, among others, perfectly portray this observation (Oberlies & Kroll, 2004). In addition, the threat to biodiversity, especially that resulting to the loss of plant species in biodiversity “hotspots”, accentuates the need

for chemical and biological investigations, which will require expanded interdisciplinary partnerships to be successful (Cragg & Newman, 2005b).

Therefore, natural products, including plants, are still a valuable source for the discovery of new therapeutics, and as part of an effort to use this resource for the search for potential anticancer agents, tropical plants, including the subject of the present dissertation, were investigated for the isolation of their secondary metabolites, focusing both on novelty and antiproliferative activity against human cancer cell lines (Bueno Pérez et al., 2014; Kinghorn et al., 2016). The phytochemical and biological investigations, as well as the related studies arisen from these will be described in subsequent chapters of this dissertation.

Chapter 2. Bioactivity-guided isolation of Podocarpus neriifolius secondary

metabolites

A. Background on Podocarpus neriifolius D. Don

The study performed in this dissertation research is centered on Podocarpus neriifolius D. Don (Podocarpaceae). Accordingly, this section will include a background on the taxonomy of this species, comprising a brief overview on the plant family and the genus, as well as a summary of the folkloric uses of the Podocarpus species, along with their phytochemical and pharmacological characteristics. It seems noteworthy to mention that the genus Podocarpus and several of its species have undergone recent taxonomic reclassification, and, according to certain authors, some of these may require further investigation (de Laubenfels, 1985; Liguo et al., 1999; Mill, 2003; Abdillahi et al., 2010; de Laubenfels, 2015; Mill, 2015). Hence, the discussion provided below may be subject to change from this point of view.

1. Phylogenetic background and ethnobotanical uses of Podocarpus neriifolius 1.1. The family Podocarpaceae and the genus Podocarpus Considered to be one of the largest families of cone-bearing plants, the

Podocarpaceae is believed to have originated in the southern hemisphere before spreading to the northern continents, and it consists of an estimated 190 species grouped into twenty genera (Mill, 2003; The Plant List, 2013). While the presence of podocarp fossils suspected to have a Laurasian source has challenged this well-accepted Gondwanan theory on the 23

ancestry of the Podocarpaceae, the latter view remains correct until fully proven otherwise

(Mill, 2003). Plants belonging to this family are comprised of trees and evergreen shrubs dispersed across tropical, subtropical, and temperate regions of the southern hemisphere, as well as in Central America, Japan, and mainland China (Liguo et al., 1999).

Among the Podocarpaceae genera, the genus Podocarpus L’Hér. ex. Pers. consists of dioecious trees and shrubs possessing the following characteristics provided by Liguo and colleagues (Liguo et al., 1999):

“Leaves spirally arranged to subopposite, ± monomorphic, juvenile leaves similar

to adult leaves in shape but often larger and/or wider, linear, lanceolate, or ovate-

elliptic, more than 5 mm, with single, obvious, often raised midvein on 1 or both

surfaces, stomatal lines present on abaxial surface. Pollen cone complexes axillary,

solitary or clustered, pedunculate or sessile; microsporophylls numerous, spirally

arranged; microsporangia 2; pollen 2-saccate. Seed-bearing structures usually

borne in leaf axils (rarely terminal), solitary (rarely more than 1); apical bracts

fertile; basal bracts often fused to form a receptacle (obsolete in some species);

ovule 1 (rarely few), inverted. Epimatium wholly enveloping seed, sometimes

colored and succulent. Seed ripening in 1st year, drupelike, dry, or leathery.”

(Liguo et al., 1999).

According to these authors, plants in this genus grow in tropical and subtropical zones worldwide (Liguo et al., 1999). However, de Laubenfels divided the genus into two subgenera, namely, Podocarpus and Foliolatus, distributed in different regions, thus

providing additional distinction between the Podocarpus L’Hér. ex. Pers. species (de

Laubenfels, 1985). Accordingly, species belonging to the subgenus Podocarpus are found in New Zealand, the Australian state of Tasmania, and Chile, as well as in the tropical

African and American highlands, while the members of the Foliolatus subgenus are mainly concentrated in Asia, Australia, and the Pacific regions (de Laubenfels, 1985).

1.2. Phylogeny of Podocarpus neriifolius Podocarpus neriifolius D. Don (Figure 6) was assigned taxonomically for the first time in 1824, and constituted the “the first tropical Asian Podocarpus L’Hér. ex Pers. species to be described” (de Laubenfels, 2015). P. neriifolius, also known as Margbensonia neriifolia D. Don, possesses a number of other synonyms, including those previously assigned to different species of the genus Nageia [e.g., N. discolor (Blume) Kuntze and N. neglecta (Blume) Kuntze] (The Plant List, 2013). In the course of nearly two centuries, various changes have been adopted regarding the taxonomy of this genus and its related species. For instance, several species previously considered as synonyms or classified under P. neriifolius have been assigned as distinct species, while others, including P. macrophyllus var. acuminatissimus E. Pritz, remain to be further validated (de Laubenfels,

2015).

Figure 6. Botanical sketch of Podocarpus neriifolius D. Don

[Reproduced without adaptation, extracted from Tropicos.org. Missouri Botanical Garden,

2018, http://tropicos.org/Image/68656, with permission under the Creative Commons

Attribution-NonCommercial-ShareAlike 3.0 Unported (CC BY-NC-SA 3.0) License https://creativecommons.org/licenses/by-nc-sa/3.0/, copyright MBG-FOC, (Tropicos.org,

2018), see Appendix A]

P. neriifolius plants are described as follows, according to the assignment of Liguo and co-workers:

“Trees to 25 m tall; trunk usually to 5 cm d.b.h.; bark grayish brown, thin, fibrous,

peeling off in longitudinal flakes; branches spreading or ascending. Foliage bud

scales erect, triangular, 1–1.5 mm wide, apex acute. Leaf blade lanceolate, usually

slightly curved, (4–)7–15(–20) × (0.5–)0.9–1.3(–2) cm, leathery, midvein raised

adaxially, flat or slightly raised abaxially, base cuneate into short petiole, apex

long acuminate; juvenile leaves wider, with obtuse, mucronate apex. Pollen cones

solitary or in clusters of 2 or 3, normally sessile, 2.5–5 cm, with several spirally

arranged, basal bracts. Seed-bearing structures axillary, solitary; peduncle 0.9–

2.2 cm. Receptacle orange-red when ripe, obconical-ellipsoid, 8–10 × 5–8 mm,

base with 2 subulate bracts 2–6 mm. Epimatium purplish red when ripe. Seed ovoid

or ovoid-subglobose, 0.8–1.6 cm, apex rounded or obtuse. Pollination May, seed

maturity Aug–Nov. 2n = 34.” (Liguo et al., 1999).

As a member of the subgenus Foliolatus (de Laubenfels, 1985) and consistent with the geographical distribution described above, P. nerifolius is spread across southeast Asia,

growing in Northeast India, Nepal, Malaysia, Vietnam, and mainland China, as well as in

Papua New Guinea and the Pacific Islands (Liguo et al., 1999).

1.3. Uses of Podocarpus species The ethnobotanical uses of several Podocarpus species, including P. neriifolius, has been compiled previously (Hocking, 1997; Abdillahi et al., 2010). Trees of the genus

Podocarpus, referred to as “yellow wood”, are well-known for their value as timber across various geographical areas and cultures. Hence, P. amara (Java), P. spicata (New

Zealand), P. imbricatus (Malaysia), P. madagascariensis (Madagascar), P. falcatus (South

Africa), and P. coriaceus (Venezuela) are all used as construction materials, from house beams and flooring, to boats and furniture (Abdillahi et al., 2010; Feleke et al., 2012).

Several species are considered as ornamental plants, and these include the African P. henkelii and the “Japanese yew”, P. macrophyllus. Moreover, the edible fruits of certain

Podocarpus plants, such as P. totara, P. nivalis, and P. spinulosa, are consumed, raw or cooked, or prepared as jams by the local populations in New Zealand, Japan, and

Queensland, respectively, while in African countries, the fruits of P. falcatus are used to make an edible oil (Abdillahi et al., 2010; Feleke et al., 2012).

In addition to the above-mentioned usage, the different parts of Podocarpus plants are also valued in traditional medicine to relieve a number of ailments and conditions. For instance, a decoction of the fruits of P. macrophyllus is consumed as a tonic to restore the health of the heart and lungs, while the bark of this species is used to treat worm-related

infections (Hocking, 1997). Furthermore, preparations of the bark, stem bark, fruits, and seeds of P. nagi are used to alleviate fever, skin diseases, stomachache, and cholera, respectively (Abdillahi et al., 2010). Finally, certain species growing on the African continent, such as P. falcatus, P. henkelii, and P. latifolius, are used to relieve chest pain and headaches in humans, as well as to treat animal diseases in dogs and cattle (Abdillahi et al., 2010).

1.4. Uses of Podocarpus neriifolius Similar to other Podocarpus species, P. neriifolius has been utilized for both medicinal and non-medicinal purposes. While the tree itself is considered as an ornamental, the yellow wood is used in carpentry for the making of furniture and musical instruments, as well as paper (Liguo et al., 1999; Abdillahi et al., 2010). Additionally, the fruits are cooked in Nepal, and the juice thereof is consumed as a beverage (Hocking, 1997). The medicinal significance of P. neriifolius is concentrated in its leaves, of which an infusion is used to alleviate bronchitis, and to treat rheumatism and painful joints in Ayurvedic medicine (Hocking, 1997; Abdillahi et al., 2010).

2. Phytochemical background of P. neriifolius and the genus Podocarpus The phytochemistry of plants of the genus Podocarpus has been investigated widely, with studies dating as early as 1930 (Briggs & Cawley, 1948). While many of these initial studies were conducted solely for phytochemical or chemotaxonomic purposes

(Cambie et al., 1963; Cambie, 1968), several subsequent reports were concentrated on the investigation of the bioactive principles from these plants responsible for wood durability, resistance to insect attack, plant-growth inhibition, and cytotoxicity against cancer cells, among other their biological properties (Galbraith et al., 1972a; Hayashi et al., 1975;

Cambie et al., 1983; Kubo et al., 1984).

Secondary metabolites isolated from these plants are composed primarily of terpenoids such as the phytoecdysones and the more abundant diterpenoids, and flavonoids including mono- and biflavones as well as anthocyanins (Imai et al., 1968b; Lowry, 1972;

Ito & Kodama, 1976; Abdillahi et al., 2010). Several review articles have covered the phytochemistry and bioactivity of Podocarpus species, and these predominantly have been focused on the nor- and bisnorditerpene dilactones, referred to as the “podolactones” (also called “nagilactones”) and classified as chemotaxonomic markers of this genus (Brown Jr.

& Sanchez L., 1974; Ito & Kodama, 1976; Hayashi et al., 1980; Barrero et al., 2003). An overview of these natural products will be presented in the sections below, with an emphasis on the phytochemical investigation of P. neriifolius, followed by examples of additional isolates obtained from other plants of the genus, and finally the bioactivity of

Podocarpus secondary metabolites will be briefly discussed.

2.1. Secondary metabolites isolated from Podocarpus neriifolius Using the SciFinder® database (Chemical Abstracts Service, CAS®, Columbus,

OH), the existing phytochemical literature on P. neriifolius and its twelve accepted

synonyms reported by The Plant List (The Plant List, 2013) was examined by using the species names as search entries. Surprisingly, among these, P. neriifolius appear to be the only species name for which earlier phytochemical investigations have been conducted.

Several compounds have been isolated from this plant, including diterpenoids, podolactones, flavones, and anthocyanins, as summarized below.

2.1.1. Diterpenes isolated from Podocarpus neriifolius

While investigating the chemical principles associated with the wood durability of ten plants within the Podocarpaceae family, including five Podocarpus species, P. neriifolius was shown to contain the phenolic diterpenoids, totarol, sempervirol, pododacric acid (20, 21, Figure 7), and the carboxytorarol dimer, macrophyllic acid

(Cambie et al., 1983), first isolated from P. macrophyllus (Bocks et al., 1963). However, the authors speculated that while these diterpenoids probably conferred this durability to the plant, additional components could also be involved. In addition, abietic and dehydroabietic acids (22, Figure 7) were obtained from the berries of P. neriifolius

(Cambie & Sidwell, 1983), while Δ8,9-isopimaric and isopimaric acids (23, 24, Figure 7) was isolated from the leaves of this plant (Wu et al., 2017).

Figure 7. Structures of representative diterpenes (20-24) from P. neriifolius

2.1.2. Podolactones isolated from Podocarpus neriifolius

As mentioned previously, the podolactones constitute the chemical markers of the genus Podocarpus (Barrero et al., 2003). These totarane-derived nor and bisnor-diterpenes are characterized by a δ-lactone between C12 and C14 and a γ-lactone linking C6 to C19

(Barrero et al., 2003) (Figure 8). Furthermore, these compounds are categorized into three main groups, namely, A-C, depending on the conjugated system between the B and C rings

(Ito & Kodama, 1976; Barrero et al., 2003). Thus, type A is distinguished by a

[8(14),19(11)-dienolide] (α-pirone) moiety, while types B and C contain a 7α,8α-epoxy-

9(11)-enolide and a 7,9(11)-dienolide, respectively (Ito & Kodama, 1976; Barrero et al.,

2003) (Figure 8).

Figure 8. Structural classification of the podolactones

Different plant parts of P. neriifolius have been investigated for their content in podolactones. These include, the plant growth inhibitors and type-B nagilactones, podolactones A and B (25, 26 Figure 9), isolated from the bark of a species collected in

Northern Queensland (Galbraith et al., 1970). Further study on the same plant led to the isolation of the first sulfoxide-containing terpenes, podolactones C and D (27) and inumakilactone B (Galbraith et al., 1971; Cassady et al., 1984; Park et al., 2003; Sato et al., 2009a), along with the type-C derivative, podolactone E (28, Figure 9) (Galbraith et al., 1972b). On the other hand, a study of the twigs of this plant species resulted in the purification of nagilactone C (29, Figure 9) as the principal antiproliferative compound

(Hayashi et al., 1968; Shrestha et al., 2001).

Figure 9. Representative podolactones (25-29) from P. neriifolius

However, the structures of several podolactones, including the above-mentioned compounds, have been revised based on X-ray crystallographic data, where the position and configuration of substituents on the A ring constitute the main structural revisions reported (Godfrey & Waters, 1975; Arora et al., 1976; Sato et al., 2009a; Addo et al.,

2015). Hence, podolactone A (25, Figure 9) was shown to possess a 2,3-β-epoxy ring rather than the initially reported 1,2-α-epoxy functionality (Galbraith et al., 1970;

Poppleton, 1975; Arora et al., 1976). A similar revision of podolactone C was provided by

Cassady and colleagues (Cassady et al., 1984). Furthermore, the unambiguous structure of podolactone D was reported several years after its first isolation, through X-ray

crystallography analysis, confirming the olefin group on the A ring, as well as the configuration of the sulfur-containing side chain (Park et al., 2003).

2.1.3. Flavonoids isolated from Podocarpus neriifolius

The leaves of P. neriifolius have been reported to contain the flavonoids, 7,4′- dimethylaromadendrin, robustaflavone (30), robustaflavone-7"-methyl ether, 2"-O- rhamnosylscoparin, and 2"-O-rhamnosylvitexin, as well as amentoflavone (31), and its derivatives, podocarpusflavones A and B (32), and isoginkgetin (33) (Figure 10) (Rizvi &

Rahman, 1974; Rizvi et al., 1974; Xu et al., 1993). In addition, amentoflavanone (31,

Figure 10) was also obtained from a fruit sample of this plant (Cambie & Sidwell, 1983).

Figure 10. Examples of flavonoids (30-33) from P. neriifolius

2.1.4. Additional secondary metabolites from Podocarpus neriifolius

Additional natural products isolated from the leaves of P. neriifolius are displayed in Figure 11, and these include the phytoecdysone, ponasterone A (34), and the phenolic compounds isovanillin, 4-hydroxybenzoic acid, and protocatechuic acid (35) (Figure 11)

(Wu et al., 2014). On the other hand, study of the twigs of this plant afforded 5α,6β- dihydroxysitosterol, and three sesquiterpenoids, spathulenol, 4α,10α-epoxy- aromadendrone, blumenol (36), as well as an unprecedented cyclopeptide and a lignan, neriitide A and neriilignan (37, 38) (Figure 11), respectively (Wu et al., 2017).

Figure 11. Structures of additional compounds from P. neriifolius

2.2. Secondary metabolites isolated from the genus Podocarpus As mentioned previously, the genus Podocarpus has been investigated extensively, both phytochemically and biologically. While flavonoids (Miura et al., 1968, 1969;

Hameed et al., 1973; Roy et al., 1987; Cheng et al., 2007), anthocyanins (Lowry, 1968;

Crowden & Grubb, 1971; Lowry, 1972), and phytoecdysones (Imai et al., 1968a; Imai et al., 1968b; Hikino et al., 1970), have been reported from these plants in addition to podolactones and other diterpenes, the latter two classes will be discussed briefly below, as they are considered more significant taxonomically.

2.2.1. Diterpenes from the genus Podocarpus

A number of diterpenes including those possessing the abietane, isopimarane, kaurane, podocarpane, sempervirol, and totarane skeletons have been reported from

Podocarpus species (Ito & Kodama, 1976; Sato et al., 2008). These species were P. halii,

P. totara, P. lambertius, P. nagi, and P. macrophyllus, with the latter two being among the most intensively investigated plants of this genus. Podocarpic acid (39, Figure 12), isolated from the heartwood of P. hallii, constitutes a podocarpane diterpene, the nomenclature of which is derived from the plant genus name (Cambie et al., 1963). Representatives of the totarane-type diterpenes isolated from Podocarpus plants include totarol, 19-oxototarol,

4β-carboxynortotarol, 4β-carboxy-17-hydroxy-19-nortotarol, and totaradiol (40-42,

Figure 12), and the inumakiols, including inumakiols C and H (43, 44, Figure 12), isolated from P. hallii and P. macrophyllus (Cambie et al., 1963; de Paiva Campello et al., 1975;

Park et al., 2004a; Sato et al., 2008). Moreover, the bark sample of P. latifolius afforded three totarane-derived diterpenoids (though reported as sempervirol-type), namely, cycloinumakiol, inumakal, and inumakoic acid (45-47, Figure 12) (Devkota et al., 2011).

In addition, the bisditerpenoids, podototarin and 7β-hydroxymacrophyllic acid (48, 49,

Figure 12), have been reported from P. totara and P. falcatus, respectively (Bocks et al.,

1963; Cambie et al., 1963; Addo et al., 2015). On the other hand, the phytochemical investigation of the bark samples of two Podocarpus species afforded sempervirol-type diterpenes, including inumakiols A and B (50, 51, Figure 13) obtained from P. macrophyllus (Sato et al., 2008).

Abietane diterpenes reported from the same genus include 16-hydroxylambertic acid, 7-oxo-18-hydroxyferruginol, and 5α,12-dihydroxy-6-oxa-abieta-8,11,13-trien-7-one

(52-54, Figure 13) from P. nagi (Feng et al., 2017a), sugiol (55, Figure 13) from P. hallii

(Cambie et al., 1963), and pododacric (21, Figure 7) and lambertic acids (56, Figure 13) isolated from P. dacrydioides and P. lambertii, respectively (Cambie et al., 1983).

Figure 12. Examples of podocarpane (39) and totarane-type (40-49) diterpenes and bisditerpenes from Podocarpus species

Figure 13. Examples of sempervirol- (50, 51) and abietane-type (52-56) diterpenes from

Podocarpus species

2.2.2. Podolactones from the genus Podocarpus

Of the reported podolactones, those belonging to the A and B types have been found to be the most abundant, while C-type derivatives constitute minor constituents (Hayashi et al., 1980; Hayashi & Matsumoto, 1982; Barrero et al., 2003). Examples from each structural type are briefly described.

Several type-A podolactones have been isolated from various parts of P. nagi. For instance, nagilactones A-D (57, Figure 14), (Hayashi et al., 1968; Ito et al., 1969; Hayashi et al., 1972b; Arora et al., 1976), as well as 3-epi-nagilactone C (58, Figure 14), (Kubo &

Ying, 1991), were obtained from the leaves of P. nagi.

Figure 14. Examples of type-A podolactones (57-64) from Podocarpus species

In addition, the root bark of this species contained 1-deoxy-2β,3β-epoxynagilactone

A (59, Figure 14), while nagilactones K and L (60, 61, Figure 14) and their derivatives were obtained from samples of its seeds (Feng et al., 2017b). Moreover, mono-, di-, and trisaccharide derivatives belonging to this same structural class, inclusive of nagilactone

B 1-O-β-D-glucoside (62) (Feng et al., 2018), nagilactoside C (63) (Xuan et al., 1995), and nagilactosides F and G (64) (Xu & Xuan, 1999) (Figure 14), have been reported from the seeds of P. nagi as well.

Representatives of type-B podolactones include sellowins A and B (65, Figure 15)

(Arora et al., 1976; Sato et al., 2009a), inumakilactones A and B, makilactones E-M (66,

Figure 15) (Sato et al., 2009a), and rakanmakilactones H-J (67, Figure 15) (Park et al.,

2004a), along with the sulfur-containing derivatives, makilactones N-R (68, Figure 15)

(Sato et al., 2009a) and rakanmakilactones A-F (69, Figure 15) (Park et al., 2004b), all of which have been isolated from P. macrophyllus. Additionally, type-B podolactone glycosides, including 3-O-β-D-glucopyranosylnagilactone G, and 16-O-β-D- glucopyranosylnagilactone G (70, 71, Figure 15), were recently obtained from the twigs of P. nagi (Zheng et al., 2018).

Figure 15. Examples of type-B podolactones (65-71) from Podocarpus species

The third podolactone class, type C, is represented on Figure 16 by nagilactone F

(72) (Sato et al., 2009b) and its derivatives 16-hydroxynagilactone F, 2β,16- hydroxynagilactone, and nagilactone I (73-75) (Addo et al., 2015), isolated from P. macrophyllus and P. falcatus, respectively. A fifth example, makilactone B (76, Figure

16), represents a chlorinated derivative from the makilactone A-D series obtained from the bark and root of P. macrophyllus (Sato et al., 2009b).

Figure 16. Examples of type-C podolactones (72-76) from Podocarpus species

3. Bioactivity of secondary metabolites from Podocarpus species Plants belonging to the Podocarpus genus, along with their secondary metabolites, notably the podolactones, have been the subject of several biological studies ranging from the investigation of their insect antifeedant activities to their cancer cell antiproliferative properties. Such biological activities along with some structure-activity relationship (SAR) observations have been described previously (Galbraith et al., 1972a; Barrero et al., 2003;

Park et al., 2004b; Abdillahi et al., 2010).

Podolactones and other natural products obtained from Podocarpus species have been shown to possess insecticidal, as well as insect antifeedant and growth inhibition effects, in addition to insect-molting activity (Kubo et al., 1984; Zhang et al., 1992). For

instance, the strong resistance of P. gracilior against insect attack has been speculated to be due to a combination of its constituents, namely, the feeding deterrent nagilactones C,

D, and F, the insectidal podolide, the non-lethal growth inhibitor podocarpusflavone, and the phytoecdysone ponasterone A (Kubo et al., 1984). Interestingly, the antifeedant activities of nagilactones A and C from the leaves of P. nagi also extended to mammals such as guinea pigs (Hayashi et al., 1992).

Numerous studies have concentrated on the inhibitory activities of the Podocarpus plants against diverse cancer cells, and several podolactones have shown activities correlating to the cytotoxicity of their parent plant extracts (Hayashi et al., 1975; Shrestha et al., 2001; Park et al., 2003; Sato et al., 2009a; Zheng et al., 2018). For example, nagilactone C was cytotoxic against the murine colon carcinoma (26-L5) and the human fibrosarcoma HT-1080 cells in vitro (Shrestha et al., 2001). Moreover, this compound and its congener, nagilactone E were both found to be effective as antitumor compounds in P-

388 lymphocytic leukemic mice at a dose of 20 mg/kg, although the latter displayed some toxicity at a higher dose (Hayashi et al., 1975). Potential mechanistic pathways responsible for these cytotoxic effects of the podolactones may be related to the inhibition of oncogenic transcription factors, such as the activator protein-1 (AP-1) (Devkota et al., 2011), the initiation of autophagy (Feng et al., 2017b), or the induction of cell cycle arrest leading to apoptosis (Zheng et al., 2018).

In addition, the secondary metabolites from plants of this genus have shown potential plant-growth inhibitory effects (Galbraith et al., 1970; Galbraith et al., 1972a;

Hayashi et al., 1972a), as well as anti-inflammatory (Feng et al., 2017a), antibacterial (Sato et al., 2008), and antifungal (Cambie et al., 1983) activities.

B. Statement of problem

The discussion in the previous chapter of this dissertation underscored the significant social burden that cancer represents, and the continuous need to address this serious health concern. Managing and overcoming this highly complex group of diseases and its collateral impacts are multifaceted and challenging tasks involving several disciplines and areas of research, ranging from prevention to diagnosis and therapy. As mentioned in the same chapter, Nature constitutes a valuable source for drug leads against different diseases, including cancer, and thus, natural products-based drug discovery and development continue to be an important area in the quest for new and improved anticancer chemotherapeutic leads.

Thus, a program project grant (P01-CA125066) was established for the purpose of investigating potential new anticancer drug leads of natural origin (Kinghorn et al., 2009;

Kinghorn et al., 2016). This multi-institutional collaborative effort, funded by the U.S.

National Cancer Institute, investigates diverse classes of natural sources consisting of higher plants, aquatic and terrestrial cyanobacteria, filamentous fungi, and lichens, and involves scientific input from several disciplines, including botany, microbiology, analytical, isolation and synthetic chemistry, in vitro and in vivo biological testing, and biostatistics (Kinghorn et al., 2016). The bioprospecting of higher plants for this program

project and its former sister grant through the “National Cooperative Drug Discovery

Groups” (NCDDG; 2000-2006) has taken place in tropical and semi-tropical regions across continents, including Africa, Asia, and Latin America (Pan et al., 2010a; Kinghorn et al.,

2011; Henkin et al., 2018). Among these are the so-called “biodiversity hotspots”, known for their vast concentration in endemic species (Myers et al., 2000). Tropical plants growing in these regions are considered a rich source of unique and bioactive secondary metabolites, owing to their highly dense and competition-prone ecological habitats, which favor the production of a variety of chemical defense mechanisms among the existing species. Therefore, organisms within these “biodiversity hotspots” are perceived as being of a particular relevance to natural product drug discovery (Balunas & Kinghorn, 2005;

Pan et al., 2010a; Kinghorn et al., 2011). Accordingly, tropical rainforests in Vietnam and

Laos have been selected as collection sites for this program project in the last few years

(Bueno Pérez et al., 2014; Henkin et al., 2018). In the case of Vietnam, to extend the diversity of collected species, expeditions were carried out in regions of different climatic zones within this country, including the Hon Ba forest, the Kego Nature Reserve, the

Hoang Lien mountains, along with the Nui Chua and Xuan Son National Parks, and these have resulted altogether in the procurement of over 1300 plant specimens between the years

2004-2015 (Henkin et al., 2018). Initial plant collections are performed either randomly or using “informed” approaches, where the former focuses mainly on biodiversity and serendipity, while the latter may involve ethnomedicinal, chemotaxonomic, and dereplication considerations (Cordell, 1995; Henkin et al., 2018). These primary plant

samples (300-500 g dry weight) are first extracted and screened for cytotoxicity against a human colon cancer cell line, HT-29, and those displaying suitable bioactivity (IC50 <20

μg/mL) are then re-collected in kilogram amounts for further investigation of their bioactive constituents (Bueno Pérez et al., 2014; Kinghorn et al., 2016; Henkin et al., 2017;

Henkin et al., 2018).

Following the above-mentioned strategy, a root sample of Podocarpus neriifolius

D. Don, having exhibited a primary cytotoxic activity against HT-29 cells (IC50 = 8.2

μg/mL), was selected as a candidate for further phytochemical and biological investigation, and thus constituted the object of the present dissertation research. As mentioned in previous sections, plants of the genus Podocarpus are well-known for their content of cytotoxic podolactones, of which several have been identified thus far (Barrero et al., 2003;

Sato et al., 2009a; Zheng et al., 2018). However, on looking into the prior studies conducted with P. neriifolius, it was seen that only a handful of such podolactones have been characterized, with equally few investigated for their biological activities against human cancer cell lines (see section on P. neriifolius above). Furthermore, as the phytochemical isolation work on the more extensively studied species, such as P. nagi and

P. macrophyllus, have most recently afforded numerous bioactive and novel compounds, it could be hypothesized that findings of a similar nature would be made through the study of P. neriifolius. Additionally, as discussed previously, the discovery of unique chemical scaffolds has led to important new biological findings, such as the mechanisms of action of clinically useful drugs or the identification of previously unknown biochemical

pathways through pharmacological probes. Therefore, the rationale for this dissertation study involved the primary objective of the program project, which is the discovery of potential new anticancer lead compounds, and additional phytochemical and biological findings that would further the scientific knowledge on this plant species. Accordingly, the remainder of this chapter will describe the work performed on the roots of P. neriifolius collected in Vietnam, with a focus on the bioactivity-guided isolation and the structural determination of its secondary metabolites.

C. Experimental

1. General experimental procedures High-resolution electrospray ionization mass spectra (HRESIMS) were collected on a LTQ OrbitrapTM mass spectrometer (Thermo Fisher Scientific Inc., Bremen,

Germany), operated in the positive-ion mode using sodium iodide as the external calibrant, and on a 15T Bruker FT-ICR mass spectrometer (ESI positive mode). NMR spectroscopic data were measured on 400 MHz Bruker AVIII400HD and 700 MHz Bruker Ascend

(Bruker Biospin, Fällanden, Switzerland) spectrometers. High-performance liquid chromatography was performed on a Hitachi Primaide HPLC (Hitachi High-Technologies

Corporation, Tokyo, Japan) equipped with a Primaide 1110 pump with a degasser, a

Primaide 1210 autosampler, and Primaide 1430 diode array detector, and using semi- preparative C18 reversed-phased HPLC columns (Dynamax 250 mm × 10 mm i.d. and

Cogent bidendate 250 mm × 10 mm i.d.). Column chromatography was conducted using

Sephadex® LH-20 resin (Supelco, Bellefonte, PA), normal-phase silica gel [(40-63 µm particle size; 230 × 400 mesh, and 63-200 µm particle size, 65 × 250 mesh)] (Sorbent

Technologies, Atlanta, GA, USA), reversed-phase C18 silica gel (Sorbent Technologies,

Atlanta, GA, USA), and Diaion® HP-20 (Alfa Aesar, Ward Hill, MA, USA). Analytical thin-layer chromatography (TLC) was performed on precoated (200 µm normal phase, and

150 µm, reversed-phase C18) aluminum-backed silica gel plates supplemented with a fluorescence indicator (UV light at 254 nm) (Sorbent Technologies, Atlanta, GA, USA).

Chromatography and spectrometric analysis were achieved using ACS, HPLC, and MS- grade solvents (Fisher Scientific, Fair Lawn, NJ, USA), and NMR experiments were performed with deuterated solvents (Cambridge Isotope Laboratories, Inc., Andover, MA,

USA).

2. Plant material A root sample of Podocarpus neriifolius D. Don (Podocarpacea) (A06795) was first collected for primary testing on January 11, 2010 in the Cotuy forest of the Nui Chua

National Park in the province of Ninh Thuan, Vietnam (GPS reading: 11o 43.159’ N; 109o

08.208’ E.) by a team led by Dr. D. D. Soejarto [collection number: Soejarto et al. 14601].

The plant was taxonomically identified by Dr. Soejarto (College of Pharmacy, University of Illinois at Chicago (UIC); Field Museum of Natural History, Chicago, IL, USA), and a voucher specimen was deposited at the John G. Searle Herbarium of the Field Museum of

Natural History, Chicago, IL, USA. Following a positive report of the cytotoxic activity

(IC50 = 8.2 µg/mL), 2 kg (dry weight) of this root sample was recollected (AA06795) from the same tree in the months of July-August 2011.

The collection of this plant material was performed under a “Memorandum of

Agreement” (MOA) for collaborative research between UIC and the Institute of Ecology and Bioloigical Resources of the Vietnam Academy of Science and Technology, in accordance with the United Nations Convention on Biological Diversity (CBD) and the

Nagoya Protocol provisions on intellectual property and benefit-sharing requirements

(Cordell, 2010; Bueno Pérez et al., 2014; Kinghorn et al., 2016).

3. Cytotoxicity-guided isolation First, a small amount of the dried powder of the root sample of P. neriifolius

(AA06795, 100 g) was extracted exhaustively by percolation in methanol (MeOH), and the resulting percolate was then evaporated in vacuo. The MeOH extract (6.1 g) was resuspended in a mixture of MeOH and H2O (ca. 9:1 v/v) and partitioned successively with hexanes and ethyl acetate (EtOAc), as illustrated on Figure 17. Portions of the dried partitioned fractions, namely, the hexanes (D1, 296 mg), aqueous (D2, 3.0 g), and EtOAc

(D3, 2.6 g), were evaluated for their antiproliferative activity against the HT-29 human colon cancer cell line. Only the EtOAc partition (IC50 = 4.3 μg/mL) was deemed active

(IC50 <20 μg/mL), and thus it was subjected to further purification.

A portion of the EtOAc partition (608 mg) was initially loaded on a Sephadex LH-

20 gel column (41 cm × 1.3 cm) and eluted with CHCl2-MeOH (1:1 v/v), affording six

fractions, D3F1-D3F6. Fraction D3F3 (ca. 192 mg) was then chromatographed on a normal-phase silica gel column (30 cm × 1.2 cm) eluted with CHCl3-EtOAc-MeOH (7:2:1 v/v) solution, resulting in five sub-fractions, D3F3.1-D3F3.5. Two fractions, D3F3.2 and

D3.F3.3, formed white precipitated solids, later identified as compounds 77 (makilactone

E) (Sato et al., 2009a) and 78 (inumakilactone A) (Park et al., 2004a), respectively. A second aliquot of the EtOAc extract was also applied to separation by Sephadex LH-20 (41 cm × 1.3 cm) using a mixture of CHCl2-MeOH (1:1 v/v) as solvent, and similarly yielded six fractions labeled D3F1′- D3F6′. All fractions were submitted to the HT-29 cytotoxicity bioassay, and fractions D3F2′ and D3F3′ were found to be active, with IC50 values of 7.0 and 3.8 μg/mL, respectively.

Figure 17. P. neriifolius extraction and partition scheme

Podocarpus neriifolius (roots)

Extract with MeOH

Methanol extract (IC50 = 6.6 μg/mL)

Dissolve in MeOH-H2O, partition with hexanes

Hexanes extract Residue (IC50 >20 μg/mL) Dissolve in H2O, partition with ethyl acetate

Aqueous extract Ethyl acetate extract

(IC50 >20 μg/mL) (IC50 = 4.3 μg/mL)

Further purification by chromatography 52

Thus, fraction D3F3′ (327.1 mg) was further separated by normal-phase silica gel chromatography eluted in CHCl3-EtOAc-MeOH (7:2:1 v/v), affording eight sub-fractions,

D3F3′.1- D3F3′.8. Of these, consecutive fractions, D3F3′.3 (46 mg; IC50 = 0.4 μg/mL) and

D3F3′.4 (20.7 mg; IC50 = 0.5 μg/mL) both formed compound 78 as a precipitated product.

Moreover, fraction D3F3′.2 (131.5 mg), with an IC50 value of 15.1 μg/mL also formed a white crystalline precipitate (77, 9.7 mg), and its supernatant was chromatographed on a reversed-phase C18 column with 40, 70, and 100% MeOH/H2O into three respective sub- fractions, D3F3′.2.1- D3F3′.2.3, all evaluated for their cytotoxic activity against HT-29 cells. The cytotoxic D3F3′.2.1 (IC50 = 2.1 μg/mL) and D3F3′.2.2 (IC50 = 7.8 μg/mL), were purified as follows. The most polar sub-fraction, D3F3′.2.1, was subjected to semi- preparative HPLC with an increasing H2O/CH3CN gradient (70:30 to 60:40 in 10 min,

60:40 to 0:100 in 12 min, 100:0 for 5 min) and at a detection wavelength of 234 nm, affording compounds 79 (3-deoxy-2β-hydroxynagilactone E, 2.1 mg) (Kubo & Ying,

1991; Zheng et al., 2018) and 80 (inumakiol D, 1.4 mg) (Sato et al., 2008), eluted at 19.1 and 20.1 min, respectively. Likewise, fraction D3F3′.2.2 was purified by semi-preparative

C18 HPLC, eluted with a gradient of H2O/CH3CN solvent system (50:50 to 30:70 in 10 min, 30:70 to 15:85 in 6 min, 15:85 to 0:100 in 1 min, 0:100 for 3 min, 0:100 to 50:50 in 2 min, 50:50 for 5 min, at 210 nm), and afforded 79 (tR = 8.7 min, 1.2 mg), as well as 81

(inumakiol E, tR = 15.4 min, 5.0 mg) (Sato et al., 2008), and 82 (4β-carboxy-19-nortotarol, tR = 8.7 min, 1.2 mg) (Park et al., 2004a).

Fraction D3F2′ (329 mg, IC50 = 7.0 μg/mL) was fractionated on a C18 column with two solvent systems, 30:70 and 100:0 H2O/MeOH (v/v), into two sub-fractions, D3F2′.1 and D3F2′.2. The first fraction was then purified by semi-preparative HPLC, using a C18 column, with a gradient of H2O/CH3CN (80:20 to 50:50 in 20 min, 50:50 to 100:0 in 1 min,

100:0 for 5 min, at 234 nm), resulting in the isolation of compounds 83 (3-deoxy- nagilactone E-2β-O-β-D-glucoside, tR = 16.2 min, 2.5 mg) and 79 (tR = 22.2 min, 1.5 mg).

Moreover, fraction D3F3′.7, also formed a precipitate, which was isolated and identified as the known compound makilactone G (84) (Sato et al., 2009a). The structures of these compounds (77-84) were determined by comprehensive one- and two-dimensional NMR spectroscopic, as well as high-resolution mass spectrometric data analysis, and complemented with comparison with values in the published phytochemical literature.

Moreover, the in vitro bioactivity of these isolates was further evaluated, as described in section 5 below.

4. Characterization of the isolated secondary metabolites 4.1. Makilactone E (77) Colorless crystalline solid; HRESIMS, positive mode, observed m/z: 423.0806 (calcd. for

+ + 1 13 C18H21ClO8Na , 423.0817 [M + Na] ) (Figure 18); H NMR (Table 1) C NMR (Table

2).

4.2. Inumakilactone A (78) White amorphous solid; HRESIMS, positive mode, observed m/z: 387.1041 (calcd. for 54

+ + 1 13 C18H20O8Na , 387.1050 [M + Na] ) (Figure 21); H NMR (Table 1) C NMR (Table 2).

4.3. 3-Deoxy-2β-hydroxynagilactone E (79)

+ White solid; HRESIMS, positive mode, observed m/z: 371.1469 (calcd. for C19H24O6Na ,

371.1465 [M + Na]+) (Figure 24); 1H NMR (Table 3) 13C NMR (Table 3).

4.4. Inumakiol D (80) White amorphous solid; HRESIMS, positive mode, observed m/z: 355.1897 (calcd. for

+ + 1 13 C20H28O4Na , 355.1880 [M + Na] ) (Figure 27); H and C NMR (Table 4).

4.5. Inumakiol E (81) Pale yellow amorphous solid; HRESIMS, positive mode, observed m/z: 369.2039 (calcd.

+ + 1 13 for C21H30O4Na , 369.2036 [M + Na] ) (Figure 30); H NMR (Tables 4 and 5) C NMR

(Table 4).

1 Table 1. H NMR data of makilactone E (77) and inumakilactone A (78) (400 MHz, in C5D5N) compared with reported data

Makilactone E (77) Makilactone E (77) Inumakilactone A (78) Inumakilactone A (78) Position (Sato et al., 2009a) (present data) (Park et al., 2004a) (present data) 1 4.97 (1H, d, 3.0) 4.97 (1H, d, 2.9) 3.68 (1H, d, 4.2) 3.66 (1H, d, 4.2) 2 4.69 (1H, br m) 4.69 (1H, m) 3.56 (1H, dd, 4.2, 5.0) 3.54 (1H, dd, 4.2, 6.2) 3 4.35 (1H, m) 4.33 (1H, m) overlap 4.69 (1H, t, 5.0) 4.67 (1H, dd, 6.1, 4.7) 5 2.75 (1H, d, 4.2) 2.74 (1H, d, 4.24) 2.18 (1H, d, 5.1) 2.16 (1H, d, 5.2) 6 5.34 (1H, dd, 4.2, 1.2) 5.34 (1H, dd broad, 0.8, 4.3) 5.15 (1H, d, 5.1) 5.13 (1H, dd, 5.2, 1.3) 7 5.18 (1H, d, 1.3) 5.18 (1H, d, 1.1) 5.18 (1H, s) 5.16 (1H, br d, 1.2) 11 6.62 (1H, s) 6.60 (1H, s) 6.84 (1H, s) 6.82 (1H, s) 14 4.75 (1H, d, 8.5) 4.74 (1H, d, 8.6) 4.79 (1H, d, 8.7) 4.76 (1H, 8.6) 15 4.39 (1H, m) 4.37 (1H, m) overlap 4.39 (1H, m) 4.37 (1H, m) 17 1.57 (3H, d, 6.2) 1.56 (3H, d, 6.26) 1.61 (3H, d, 5.9) 1.58 (3H, d, 6.2) 18 1.64 (3H, s) 1.68 (3H, s) 1.42 (3H, s) 1.39 (3H, s) 20 2.00 (3H, s) 2.00 (3H, s) 1.60 (3H, s) 1.58 (3H, s) OH-2 8.30 (1H, br d) 8.33 (1H, d, 4.4 Hz) OH-3 5.48 (1H, d, 8.7) OH-15 7.01 (1H, d, 5.8) 7.02 (1H, d, 5.4)

Table 2. 13C NMR data of makilactone E (77) and inumakilactone A (78) (100 MHz, in

C5D5N) compared with reported data

Makilactone E (77) Makilactone E (77) Inumakilactone A (78) Inumakilactone A Position (Sato et al., 2009a) (present data) (Park et al., 2004a) (78) (present data)

1 63.6 63.4 55.9 55.7

2 73.1 72.8 51.1 50.8

3 71.4 71.2 68.3 68.1

4 44.6 44.4 49.0 48.8

5 40.6 40.4 45.5 45.3

6 72.6 72.4 71.8 71.6

7 55.6 55.4 56.1 55.8

8 57.6 57.4 57.2 57.0

9 155.5 155 158.2 158.0

10 42.6 42.4 37.9 37.7

11 119.5 119.0 119.3 119.0

12 163.3 163.0 163.3 163.1

14 83.0 82.7 82.9 82.7

15 63.8 63.6 63.8 63.6

16 21.0 20.8 21.1 20.8

18 22.3 22.1 25.3 25.0

19 178.1 177.9 176.7 176.4

20 26.1 25.9 20.8 20.5

Table 3. 1H and 13C NMR data of 3-deoxy-2β-hydroxynagilactone E (79) (400 and 100 MHz) compared with reported data a b Appeared as dddd (13.5, 8.5, 7.4, 4.9) in CD3OD. Value obtained from HMBC spectrum.

3-Deoxy-2β-hydroxynagilactone E (79) 3-Deoxy-2α-hydroxynagilactone E 3-Deoxy-2β-hydroxynagilactone E Position (in C5D5N) (present data) (in C5D5N) (Kubo & Ying, 1991) (in C5D5N) (Zheng et al., 2018)

δC δH δC δH δC δH 1a 40.4 2.42 (overlap, dd, 13.5, 8.5) 40.65 2.42 (1H, dd, 13.2, 9.1) 41.1 2.47 (dd, 13.5, 8.7) 1b 1.80 (dd, 13.5, 7.4) 1.81 (1H, dd, 13.2, 7.3) 1.84 (dd, 13.5, 7.5) 2 63.5 4.20 (m)a 63.70 4.22 (1H, m) 64.2 4.24 (m) 3a 38.3 2.14 (dd, 13.5, 4.9) 38.52 2.15 (dd, 13.6, 4.4) 39.0 2.19 (dd, 13.7, 4.8) 3b 2.48 (overlap, t, 13.5) 2.48 (1H, dd, 13.6, 13.2) 2.52 (t, 13.7) 4 42.0 42.32 42.8 5 42.1 1.95 (d, 5.0) 42.32 1.96 (1H, d, 4.0) 42.7 1.99 (d, 5.0) 6 72.8 5.14 (dd, 5.0, 1.4) 73.03 5.13 (1H, d, 4.0) 73.5 5.18 (dd, 5.0, 1.5) 7 54.1 4.24 (d, 1.4) 54.33 4.23 (1H, s) 54.8 4.28 (d, 1.5) 8 57.8 58.00 58.5 9 159.1 159.34 159.8 10 36.7 37.01 37.5 11 118.2 6.30 (s) 118.44 6.30 (1H, s) 118.9 6.35 (s) 12 163.6b 163.80 164.4 14 82.6 b 4.58 (d, 3.3) 82.83 4.58 (1H, d, 2.0) 83.2 4.62 (d, 3.3) 15 26.7 1.96 (overlap, m) 26.98 1.97 (1H, m) 27.4 1.98 (m) 16 21.1 1.16 (d, 6.7) 16.51 1.18 (3H, d, 6.6) 21.8 1.20 (d, 6.7) 17 16.3 1.02 (d, 6.7) 21.32 1.03 (3H, d, 6.6) 17.0 1.05 (d, 6.7) Continued 58

Table 3 Continued

3-Deoxy-2β-hydroxynagilactone E 3-Deoxy-2α-hydroxynagilactone E 3-Deoxy-2β-hydroxynagilactone E Position (79) (in C5D5N) (present data) (in C5D5N) (Kubo & Ying, 1991) (in C5D5N) (Zheng et al., 2018) δC δH δC δH δC δH 18 22.7 1.31 (s) 28.80 1.32 (3H, s) 23.4 1.34 (s) 19 181.1b 181.30 181.8 20 28.6 1.28 (s) 22.95 1.30 (3H, s) 29.3 1.31 (s)

Table 4. 1H and 13C NMR data data of of inumakiols D (80) (400, 175 MHz) and E (81) (400, 100 MHz) a b Assigned by HMBC correlation with H3-18. No HMBC correlation observed.

Position Inumakiol D (80) (in CD3OD) Inumakiol E (81) (in CDCl3) δC δH δC δH 1 39.8 2.25 (1H, d or overlap) 39.3 1.42 (1H, m, overlap) 1.36 (1H, overlap) 2.19 (1H, broad dt, 13.1, 3.6) 2 20.12 2.07 (1H, overlap) 19.9 1.63 (1H, m) 1.60 (1H, broad d, 11.8) 1.99 (1H, m) 3 38.4 2.27 (1H, overlap) 36.9 1.16 (1H, td, 13.6, 4.2) 1.14 (1H overlap) 2.27 (1H, broad d, 13.4) 4 42.5a 43.3 5 44.9 2.0 (1H, m overlap) 45.2 1.98 (1H, m overlap) 6 31.1 2.3 (1H, overlap) 23.7 1.97 (1H, m, overlap) 2.2 (1H, overlap) 2.6 (1H, m) 7 64.8 4.95 (1H, broad s) 74.5 4.42 (1H, broad s) 8 133.6 133.1 9 140.0 141.2 10 48.1b 38.7 11 123.5 6.7 (1H, d, 8.5) 124.1 7.02 (1H, d, 8.5) 12 116.3 7.0 (1H, d, 8.5) 117.3 6.64 (1H, d, 8.5) 13 154 152.6 14 132.7 133.3 15 27.6 3.55 (1H, m) 27.8 3.17 (1H, m) 16 19.6 1.35 (3H, d, 7.1) 20.5 1.43 (3H, d, 7.0) 17 19.5 1.43 (3H, d,6.8) 20.8 1.44 (3H, d, 7.0)

Continued 60

Table 4 Continued

Position Inumakiol D (80) (in CD3OD) Inumakiol E (81) (in CDCl3) δC δH δC δH 18 27.6 1.29 (3H, s) 28.3 1.38 (3H, s) 19 182a 182.3 20 21.7 1.09 (3H, broad s) 22.2 1.09 (3H, s) 7-OCH3 55.3 3.5 (3H, s)

1 Table 5. H NMR data of inumakiol E (81) (400 MHz, in C5D5N) compared with reported data

a b Values obtained by selective 1D-NOESY, irradiating δH = 1.60 (H-2a), δH = 2.56 (H-3b), a d e δH = 1.48 (H3-18), δH = 2.90 (h-6b), δH = 4.66 (H-7).

Position Inumakiol E (81) (Sato et al., 2008) Inumakiol E (81) (present data) 1.39 (1H, m) 1.38 (overlap) 1 2.29 (1H, d, 13.4) 2.30 (overlap) 1.60 (1H, d, 13.4) 1.60 (1H, m) 2 2.38 (1H, m) 2.33 (overlap)a 1.14 (1H, ddd, 13.4, 13.4, 3.5) 1.13 (ddd, 13.3, 13.3, 4.2)b 3 2.57 (1H, d, 13.4) 2.56 (broad dt 13.3, 3.3) 4 5 2.22 (1H, d, 13.7) 2.22 (1H, d, 13.0) c 2.38 (1H, m) 2.34 (1H, m)d 6 2.91 (1H, d, 13.7) 2.90 (1H, broad d, 14.4) 7 4.65 (1H, s) 4.66 (1H, s) 8 9 10 11 7.12 (1H, d, 8.5) 7.12 (1H, d, 8.6) 12 7.16 (1H, d, 8.5) 7.16 (1H, d, 8.6) 13 14 15 3.50 (1H, m) 3.49 (m, overlap) 16 1.75 (3H, d, 6.9) 1.75 (3H, d, 6.9) 17 1.83 (3H, d, 6.9) 1.82 (3H, 6.9) 18 1.49 (3H, s) 1.48 (3H, s) 19 20 1.39 (3H, s) 1.38 (3H, s) e 7-OCH3 3.51 (3H, s) 3.51 (3H, s)

4.6. 4β-Carboxy-19-nortotarol (82) White amorphous solid; HRESIMS, positive mode, observed m/z: 339.1934 (calcd. for

+ + 1 13 C20H28O3Na , 339.1931 [M + Na] ) (Figure 34); H and C NMR (Table 6).

4.7. Makilactone G (84) White amorphous solid; HRESIMS, positive mode, observed m/z: 405.1144 (calcd. for

+ + 1 13 C18H22O9Na , 405.1156 [M + Na] ) (Figure 38); H and C NMR (Table 6).

Table 6. 1H (400 MHz) and 13C (100 MHz) NMR data of 82 and 84

4β-Carboxy-19-nortotarol (82) (in CDCl3) Makilactone G (84) (in CD3OD) Position δC δH δC δH 2.22 (1H, m, overlap) 1 40.1 72.1 4.10 (1H, d, 3.8) 1.33 (1H, overlap) 2.00 (1H, overlap) 2 20.0 72.9 3.97 (1H, dd, 5.3, 3.7) 1.62 (1H, m) 2.26 (1H, overlap) 3 37.3 71.7 3.89 (1H, d, 5.2, overlap) 1.09 (1H, m) 4 43.7 44.1 5 52.0 1.51 (1H, dd, 12.3, 1.5) 40.0 2.35 (1H, d, 4.3) 2.24 (1H, br dd, 12.3, 5.1)a 6 21.1 72.6 5.10 (1H, dd, 4.2, 1.6) 2.00 (1H, dddd, 12.3, 6.8, 5.1, 1.6)a 2.98 (1H, dd, 16.7, 4.8) 7 30.0 54.8 4.56 (1H, d, 1.6) 2.68 (1H, ddd, 16.7, 12.4, 6.5) 8 134.3 56.7 9 141.0 157.4 10 38.5 40.8 11 124.2 7.01 (1H, d, 8.6) 117.1 6.20 (1H, s) 12 114.6 6.55 (1H, d, 8.6) 164.2 13 130.9 14 152.0 82.3 4.39 (1H, 8.5) 15 27.2 3.31 (1H, septet, 7.1) 63.1 3.89 (1H, overlap) 16 20.4 1.36 (3H, d, 7.1) 19.1 1.33 (d, 6.2) 17 20.3 1.36 (3H, d, 7.1) 18 28.6 1.36 (3H, s) 21.2 1.52 (3H, s) 19 183.2 178.7 20 23.2 1.15 (3H, s) 22.9 1.46 (3H, s)

5. Biological evaluation As part of the established bioactivity-guided isolation protocol of program project

(P01-CA125066) described earlier in this chapter, in vitro biological evaluation of the initially collected plant sample and subsequent crude extracts and the fractions obtained from P. neriifolius root sample against HT-29 cells were conducted at The Ohio State

College of Pharmacy, while the purified compounds were evaluated by collaborators at the

University of Illinois at Chicago (UIC) using a panel of four human cancer cell lines as described below.

5.1. Cytotoxicity against the HT-29 cell line (OSU) Cytotoxicity assay against the human colon cancer cell line, HT-29, was performed on 96-well microplates, by Dr. Hee-Byung Chai at OSU, according to a procedure detailed previously (Pan et al., 2010b), and briefly described in Appendix B. The crude MeOH extract along with its partitioned fractions, the sub-fractions from the EtOAc partition (D3), and in some cases, the isolated precipitates (77 and 78) therefrom, were submitted for testing at a primary concentration of 4 mg/mL in 100% DMSO prior to serial dilutions at various concentrations (20, 4, 0.8, and 0.16 μg/mL in 10% DMSO). As a colorimetric assay, the absorbance values were measured on a Bio-Tek μQuant microplate reader, and tabulated for a nonlinear regression analysis using a TableCurve2Dv4 (AISN Software,

Inc., Mapleton, OR) software. The IC50 (concentration required to inhibit 50% cell growth) values of the test samples were then calculated from the obtained curve.

5.2. Cytotoxicity against the HT-29, MDA-MB-231, MDA-MB-435, and OVCAR3 cell lines (UIC) The procedure used to evaluate the in vitro cytotoxicity of the purified compounds

(77-84) from P. neriifolius against a panel of four cancer cell lines, namely, colon (HT-

29), breast (MDA-MB-231), ovarian (OVCAR3), and melanoma (MDA-MB-435), has also been described previously (Zhao et al., 2015; Ren et al., 2017). This biological evaluation was performed by collaborators (Dr. Wei-Lun Chen and Mr. Austin A.

Czarnecki) from the laboratory of Dr. Joanna E. Burdette at the UIC College of Pharmacy.

A brief description of the bioassay protocol is presented in Appendix C.

5.3. In vivo assay While inumakilactone A (78) was submitted for in vivo biological testing, this assay was conducted with additional amounts of this compound, obtained from a large-scale isolation described in Chapter 3 of this document. Therefore, the corresponding assay protocol will be described in detail in that chapter.

D. Results and Discussion

From the initial small-scale isolation study of the root of Podocarpus neriifolius D.

Don, eight compounds were obtained, of which one was new (83) and seven (77-82 and

84) have been reported previously (Kubo & Ying, 1991; Park et al., 2004a; Sato et al.,

2008; Sato et al., 2009a; Zheng et al., 2018). While the NMR (1 and 2D) spectroscopic and high-resolution MS data for the new compound (83) were obtained and used to determine 65

its structure, further characterization of 83 was performed following isolation of an additional amount of this compound. Moreover, compound 79, although a known compound, is being assigned as possessing a β-oriented hydroxy group at C-2 rather than the previously reported α-oriented derivative (Kubo & Ying, 1991). Therefore, the structural re-assignment of 79 will be discussed in more depth.

1. Structure determination of the isolated compounds 1.1. Makilactone E (77) Compound 77 was obtained as a crystalline solid. Its HRESIMS data showed a molecular ion peak at m/z 423.0806 [M+Na]+. An ion peak at m/z 425.0781 [M+Na+2]+ with an intensity of approximately a third of that of the molecular ion indicated the presence of a Cl atom (3:1 ratio of [M]+ and [M+2]+) (Figure 18). Thus, the molecular formula of

13 77 was determined to be C18H21ClO8. Inspection of the C NMR spectrum of 77 indicated the presence of two carbonyl and two olefinic carbon signals resonating at δC 177.9 (C-19),

163.3 (C-12), 155.5 (C-9), and 119.5 (C-11), respectively (Figure 19). In addition, three hydroxylated methine carbons were observed at δC 72.8 (C-2), 71.2 (C-3), and 63.6 (C-15), together with three methyl carbons [δC 20.8 (C-16), 22.1 (C-18), and 25.9 (C-20)]. HSQC and HMBC correlations (Figure 20) allowed for the assignment of the Cl-bearing carbon at C-1 (δC 63.1), as cross peaks were observed between the methyl protons of C-20 and this carbon, as well as between H-1 and carbons at C-9, C-10, and C-20.

Figure 18. HRESIMS [(full, (a) and zoomed with prediction (b)] spectra of makilactone E

(77)

AA06795D3F3.2MP_171116154423 #13 RT: 1.01 AV: 1 NL: 1.15E7 T: FTMS + p ESI sid=50.00 Full ms [150.00-1000.00] 423.08060 R=131100 100

95 90 (a) 85

RelativeAbundance 40 365.13519 35 R=139701

25 391.36508 447.42709 R=135201 R=126301 20 475.45834 R=123001 15 503.48956 R=118501 10 283.09131 531.52087 823.17346 629.23438 685.43439 727.48120 5 215.78882 R=158001 350.30234 R=115001 581.23450 775.10864 R=93100 907.77173 985.23236 R=144901 R=107301 R=101601 R=98701 R=183501 R=111401 R=94604 R=88501 R=77901 0 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000 m/z

(b) 423.08060 NL: R=131100 1.15E7 100 + [M+ Na] AA06795D3F3.2MP_171 90 116154423#13 RT: 1.01 AV: 1 T: FTMS + p ESI 80 sid=50.00 Full ms + [150.00-1000.00] 70 [M+2+ Na]

40 425.07813 R=130800 RelativeAbundance 30 Measured spectrum for 77 424.08401 425.44489 20 R=130304 R=134604 426.08096 10 421.40314 422.06854 423.83142 R=128704 427.09311 428.09631 429.31750 431.06769 432.07111 433.36380 434.36731 R=135104 R=118004 R=132104 R=128104 R=133304 R=131304 R=132101 R=99004 R=129001 R=129304 NL: 423.08172 6.11E5 100 C 18 H21 ClNaO 8+: 90 C 18 H21 Cl1 Na 1 O8 pa Chrg 1 80

40 425.07877 30 424.08507 20 Predicted spectrum for 77 10 426.08212

421 422 423 424 425 426 427 428 429 430 431 432 433 434 m/z

Further HMBC cross peaks between proton H-11 and the carbons at C-12 (δC 67

163.3), C-20 (δC 25.9), and C-10 (δC 42.6), and correlations between H-5 (δH 2.74) and carbons at C-19 further suggested that 77 is a bisnor-diterpene dilactone (Ito & Kodama,

1976). Finally, cross peaks of H-14 with C-15 and the methyl at C-16 indicated that the side chain is attached at C-14, as shown in the structure of makilactone E, and confirmed by comparison of the above data with the literature (Sato et al., 2009a).

Figure 19. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of makilactone E (77)

(in C5D5N)

(a)

(b)

Figure 20. HSQC (a) and HMBC (b) spectra of makilactone E (77) (400 MHz, in C5D5N)

(a)

(b)

1.2. Inumakilactone A (78) Compound 78 was isolated as a white amorphous powder, and its HRESIMS

displayed a sodiated ion peak at m/z 387.1041 [M+Na]+ (Figure 21), and suggested a nor-

1 13 diterpene with a molecular formula of C18H20O8. Inspection of its H and C NMR spectra

(Figure 22) revealed the presence of two carbonyl [δC 176.7 (C-19) and 163.3 (C-12)], two

hydroxy [(δC 68.1 (C-3) and δC 63.6 (C-15)], and two pairs of epoxy [δC 55.7 (C-1) and

50.8 (C-2); 57.0 (C-8) and 55.8 (C-7)] carbon signals. In addition, the presence of proton

resonances at δH 6.82 (s, C-11), 5.13 (dd, J = 5.2, 1.3 Hz, H-6) and 4.76 (d, J = 8.6 Hz, C-

14), typical for some B-type podolactones (Ito & Kodama, 1976), further indicated that 78

is a nor-diterpene dilactone belonging to this same sub-class. Interpretation of the HMBC

data of 78 led to confirmation of the position of the second epoxy group at C-1,C-2, and

the assignment of the hydroxy group-bearing carbons as C-3 and C-15 (Figure 23).

Figure 21. HRESIMS spectrum of inumakilactone A (78)

Podocarpus_171116154423 #14 RT: 1.10 AV: 1 NL: 2.63E6 T: FTMS + p ESI Full ms [150.00-1000.00] 387.10406 R=136001 C 18 H20 O8 Na 100

95 90 85

70 751.21948 R=98201 65

55 50 45 419.13004 RelativeAbundance 40 R=130704 C H O Na 35 19 24 9 30 25

20 447.42709 727.44952 783.24561 R=126801 15 343.07797 R=92901 R=95504 C 30 H55 O2 503.48956 R=145701 685.43396 215.78871 R=118404 10 C H O Na R=103301 R=181201 290.96851 16 16 7 C H O 559.55243 29 59 6 C 30 H62 O15 Na 811.54272 R=123704 R=103004 868.60950 907.77136 5 R=95201 985.23187 C 17 O4 Na C 29 H60 O8 Na R=90704 R=88301 R=84201 0 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000 m/z

Therefore, 78 was determined as the known compound, inumakilactone A, and both its 1H and 13C NMR data corresponded with previously reported data (Tables 1 and 2)

(Park et al., 2004a).

Figure 22. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of inumakilactone A

(78)

(a)

(b)

Figure 23. HSQC (a) and HMBC (b) spectra of inumakilactone A (78) (in C5D5N)

(a)

(b)

1.3. 3-Deoxy-2β-hydroxynagilactone E (79) Compound 79 was isolated as a white solid. Its HRESIMS exhibited a positive

molecular ion peak at m/z 371.1469, and its molecular formula was determined to be

1 C19H24O6, suggesting a nor-diterpenoid. Inspection of its H NMR spectrum showed that

79 shared many similar features to the two podolactones discussed above.

Figure 24. HRESIMS spectrum of 3-deoxy-2β-hydroxynagilactone E (79)

PodoLGD3C14M1-H17_2018-4-16_180321163227 #4-59 RT: 0.12-1.99 AV: 56 NL: 4.63E6 T: FTMS + p ESI Full ms [150.00-800.00] 371.14688 R=116167 100

RelativeAbundance 40 35 30

15 413.21180 R=110305 10 447.42839 R=105897 177.08889 357.20432 391.36599 503.49093 5 207.09946 274.27461 301.14165 531.52285 595.30538 639.33167 683.35810 719.30503 749.91344 787.77675 R=167491 R=117745 R=113310 R=99994 R=155848 R=135240 R=128986 R=93886 R=85517 R=90847 R=87614 R=70794 R=89733 R=74352 0 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

1 In the H NMR spectrum, the singlet resonance signal at δH 6.30 ppm, together with

proton signals at δH 5.14 (dd, J = 5.0, 1.4 Hz), 4.24 (d, J = 1.4 Hz), and 1.95 (d, J = 5.0

Hz), corresponding to protons at positions H-11, H-6, H-7, and H-5, respectively, indicated

a typical type-B podolactone skeleton (Figure 25). The HMBC data suggested the presence of lactone carbonyl carbons at C-12 (δC 163.6) and C-19 (δC 181.1), based on cross peaks with H-11 (δH 6.30, s) and H-5 (δH 1.95, d, J = 5.0 Hz), respectively (Figure 26).

Figure 25. 1H (400 MHz) (a) and 13C (175 MHz) (b) NMR spectra of 3-deoxy-2β- hydroxynagilactone E (79) (in C5D5N)

(a)

(b)

1 The H NMR data acquired in CD3OD revealed the presence of a hydroxy group on the A ring at position C-2, based on the splitting pattern of H-2, and that of both the adjacent protons attached to C-1 and C-3. The above spectroscopic data together with interpretation of HSQC and HMBC correlations were consistent with the previously reported 3-deoxy-2α-hydroxynagilactone E (Kubo & Ying, 1991). However, the analysis of another report by Addo and co-workers (Addo et al., 2015) of the structural revision of a type-C podolactone, 2α-16-hydroxynagilactone F, to its 2-epimer (74, Figure 16) led to a different conclusion. Through structural predictions based on bond angles and J values

[H-2 (dddd, J = 12.9, 9.1, 7.2, 5.1 Hz)], as well as verification by X-ray crystallography, the configuration of the hydroxy group at C-2 of this type-C podolactone isomer was reassigned as 2β-OH (Addo et al., 2015). Accordingly, it was suggested that compound 79, in having the same splitting pattern at H-2 (dddd, J = 13.5, 8.5, 7.4, 4.9 Hz) as the revised

2β-16-hydroxynagilactone F (74), should instead be assigned as 3-deoxy-2β- hydroxynagilactone E. A recent publication by Zheng and colleagues, reporting the isolation of 3-deoxy-2β-hydroxynagilactone E as a new compound (Zheng et al., 2018) further confirmed this assertion. As shown on Table 3, the present data and those reported by these authors closely match. Thus, 79 was determined as the known compound, 3- deoxy-2β-hydroxynagilactone E, first isolated by Kubo and Ying from Podocarpus nagi

(Kubo & Ying, 1991), and later reported as the corresponding 2-epimer and thus having a revised structure (Zheng et al., 2018).

Figure 26. HSQC (a) and HMBC (b) spectra of 3-deoxy-2β-hydroxynagilactone E (79) (in

C5D5N)

(a)

(b)

1.4. Inumakiol D (80) Compound 80 was obtained as a white amorphous solid. Its HRESIMS in the

positive mode showed a sodiated ion peak [M+Na]+ at m/z 355.1897, suggesting a

1 molecular formula of C20H28O4 (calcd. m/z 355.1880) (Figure 27). Inspection of its H

NMR spectrum measured in CD3OD (Figure 28), revealed the presence of two ortho-

coupled aromatic protons [δH 7.00 (H-12, d, J = 8.5 Hz) and 6.70 ( H-11, d, J = 8.5 Hz)],

and three signals appearing at δH 3.55 (H-15, m), 1.43 (H3-17, d, J = 6.8 Hz), and 1.35 (H3-

16, d, J = 6.8 Hz) corresponding to component functionalities of an isopropyl group, along

with two additional methyl groups at δH 1.29 (H3-18, s) and 1.09 (H3-20, s). In addition, a

downfield singlet methine proton resonating at δH 4.95 (H-7, br s) indicated that it was

attached to a hydroxy group-bearing carbon atom (δC 64.8), and this was confirmed by

HSQC spectroscopic cross peaks (Figure 29).

Figure 27. HRESIMS spectrum of inumakiol D (80)

AA06795D3.F3'.2_2 #168-299 RT: 1.33-2.37 AV: 132 NL: 1.55E7 T: FTMS + p ESI Full ms [50.00-2000.00] 413.21348 R=31745 100 355.18974 R=34743 95

45 RelativeAbundance 40 35

25 274.27536 R=39135 687.38942 20 443.22393 R=25702 369.18727 R=31255 R=34318 15 325.16116 R=35753 10 501.37761 545.40390 589.43024 727.38448 647.47233 177.66026 261.13214 R=28962 R=28118 R=26937 R=24382 R=25832 815.80799 5 R=24585 R=39918 R=22556 0 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000 m/z 77

Figure 28. 1H (400 MHz) (a) and 13C (175 MHz) (b) NMR spectra of inumakiol D (80) in

CD3OD

(a)

(b)

Analysis of the 13C NMR spectrum indicated the presence of six aromatic carbons

[δC 154 (C-13), 140.1 (C-9), 133.6 (C-8), 132.7 (C-14), 123.5 (C-11), and 116.3 (C-12)], of which C-13 was hydroxylated (Figure 28). The remaining protons and carbons were

assigned on the basis of 2D-NMR data, notably, HSQC and HMBC (Figure 29).

Figure 29. HSQC (400 MHz) (a) and HMBC (700 MHz) (b) spectra of inumakiol D (80) in CD3OD

(a)

(b)

Interpretation of the HSQC data suggested that 80 possesses four saturated methylene groups, although their respective proton signals could not be fully determined due to their significant overlap. HMBC correlations between methyl protons at H3-16 and

H3-17 and C-14 indicated that the isopropyl group is attached to the aromatic ring at C-14, while correlations between protons H-11, H-12, and H-15 with C-13, in addition to the above observations, confirmed the position of the hydroxy group at C-13. Moreover, a cross peak on the HMBC spectrum of 80 between H3-18 and a peak at δC ~182 ppm, confirmed the presence of a carboxylic acid group attached at C-4. Similarly, C-4 (δC

~42.5) was assigned on the basis of a HMBC cross peak to H3-18. The above evidence was in agreement with a totarane-type diterpene skeleton and corresponded to the known compound, inumakiol D (Sato et al., 2008).

1.5. Inumakiol E (81) Compound 81 was isolated as a pale yellow amorphous solid. Its molecular formula was determined to be C21H30O4 based on a molecular ion peak at m/z 369.2039 obtained from its HRESIMS [M+Na]+ (Figure 30). The spectroscopic data of 81 revealed a close similarity to those of 80, except that 81 possesses one methoxy carbon instead of a hydroxy group in addition to the acid carbonyl (δC 182.3, C-19), four methylene [δC 39.3 (C-1), 36.9

(C-3), 23.7 (C-6), and 19.9 (C-2)], and six aromatic carbon signals [δC 152.6 (C-13), 141.2

(C-9), 133.3 (C-14), 133.1 (C-8), 124.1 (C-11), and 117.3 (C-12)] (Figure 31), along with the isopropyl group attached to C-14.

Figure 30. HRESIMS spectrum of inumakiol E (81)

PodoD3-F3'-2-2-6_2018-6-17_run2_180321163227 #3-34 RT: 0.05-0.90 AV: 32 NL: 5.64E6 T: FTMS + p ESI Full ms [150.00-800.00] 369.20394 R=138699 100

75 70 65 391.18588 R=132699 60 413.21214 55 R=130601 50

45 301.14150 R=154713 RelativeAbundance 40 35 30

20 325.15999 281.13378 R=159720 R=147914 15 351.15713 447.42878 485.19488 715.41876 261.13127 R=143933 R=122182 R=98231 R=163798 R=127063 10 181.06232 217.10509 787.77465 R=200236 517.37146 R=181599 545.40285 589.42910 683.35667 R=92901 5 R=119415 R=114532 633.45537 R=109342 R=102505 R=99548 0 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

13 Figure 31. C NMR spectrum of inumakiol E (81) (100 MHz, in CDCl3)

These signals together with the 13C NMR resonances corresponding to the quaternary carbons at C-4 (δC 43.3) and C-10 (δC 38.7), and the methine proton H-5 (δH

1.98), indicated a totarane-type diterpene as for 80, and this observation was confirmed by

HMBC correlations (Figure 33).

1 Figure 32. H NMR spectra of inumakiol E (81) in CDCl3 (a) and in C5D5N (b) (400 MHz)

(a)

(b)

Figure 33. HSQC (a) and HMBC (b) spectra of inumakiol E (81) (400 MHz, in CDCl3)

(a)

(b)

As shown on Table 5, the 1H NMR data of 81 proved to be nearly identical to those reported for inumakiol E (Sato et al. 2008) (Spectroscopic comparison on Figure 32).

Thus, compound 81 was determined as inumakiol E based on the above evidence and

comparison with the reported phytochemical data (Sato et al., 2008).

1.6. 4β-Carboxy-19-nortotarol (82) Compound 82 was obtained as an amorphous solid. Its measured HRESIMS

displayed a sodiated positive ion peak at m/z 339.1934 [M+Na]+ (Figure 34), indicative of

a molecular formula of C20H28O3, which was similar to that of 80, except for the additional

oxygen in the latter compound (C20H28O4).

Figure 34. HRESIMS spectrum of 4β-carboxy-19-nortotarol (82)

PodoD3-F3'-2-3M-1_2018-6-17_run2_180321163227 #7-68 RT: 0.17-1.86 AV: 62 NL: 1.39E6 T: FTMS + p ESI Full ms [150.00-800.00] 339.19336 R=142207 100

85 80 75

65 329.17504 60 R=148486 413.21196 55 R=127583

50 397.41532 425.44660 R=130696 R=128462 45 301.14139 R=151110 353.17254 RelativeAbundance 40 R=142773 271.20601 655.39730 R=102052 35 245.07877 R=161020 R=169011 30 367.15179 447.42856 R=140022 25 189.12774 R=123913 R=189882 20 677.37927 475.45988 R=98844 15 204.06586 R=187429 R=120721 689.33517 793.82359 10 227.23729 R=100132 R=91133 178.50960 503.49128 R=177911 765.79231 R=118492 R=121446 641.48686 727.38257 5 531.52267 592.52903 R=93080 R=89019 R=111520 R=113817 R=101581 0 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

As with inumakiol D (80), six aromatic carbons signals resonating at δC 152.0,

141.0, 134.4, 130.9, 124.2, and 114.6, corresponding to carbons at positions 14, 9, 8, 13,11,

and 12, respectively, were present in the 13C NMR spectrum of 82 (Figure 35). Moreover,

inspection of its 1H NMR data showed two ortho-coupled aromatic protons (H-11 and H-

12), occurring at δH 7.01 (d, J = 8.6 Hz) and 6.55 (d, 8.6), respectively, along with four methyl proton signals, of which two overlapped doublets were part of an isopropyl moiety

[δH 1.36 (6H, d, J = 7.1 Hz)].

Figure 35. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of 4β-carboxy-19-nortotarol (82) (in CDCl3)

(a)

(b)

Complementing these 1D-NMR spectroscopic data, HSQC analysis revealed the presence of five methylene groups (Figure 37), confirming that 82 and 80 share the same skeleton, with the former having an additional methylene rather than a hydroxy group- bearing methine carbon at C-7. In addition, the resonance and splitting pattern of the protons at C-6 were verified by a selective 1D-TOCSY NMR experiment through irradiation of H-5 [δH 1.51 (dd, J = 12.3 Hz, 1.5 Hz)] (Figure 36).

Figure 36. Selective 1D-TOCSY (a) and 1H (b) NMR spectra of 4β-carboxy-19-nortotarol

(82) (400 MHz, in CDCl3)

Irradiating 1.51 ppm (H-5)

H-7a H-7b H-6a H-6b H-5 (irrad)

(a)

(b)

(a) 1D-TOCSY spectrum (Irrad. H-5, 1.51 ppm)

(b) 1H NMR spectrum

Figure 37. HSQC (a) and HMBC (b) spectra of 4β-carboxy-19-nor-totarol (82) (400 MHz, in CDCl3)

(a)

(b)

Further interpretation of the HSQC and HMBC data (Figure 37) allowed for the assignment of 82 as 4β-carboxy-19-nor-totarol, which matched reported data for this

compound (Park et al., 2004a).

1.8. Makilactone G (84) Compound 84 was obtained as a white amorphous solid. Its molecular formula was

determined as C18H22O9 on the basis of a sodiated molecular ion peak at m/z 405.1144

[M+Na]+, from its HRESIMS (Figure 38), suggesting a bisnorditerpene-type molecule.

This molecular formula was similar to that of makilactone E (77), except that 84 contained

a hydroxy group instead of the Cl atom in 77. Comparison of the 1H and 13C NMR data

(Figure 39) of these compounds indeed revealed that they only differ structurally by the

substituent at C-1, with one being chlorinated and the other hydroxylated, in agreement

with the high-resolution MS data.

Figure 38. HRESIMS spectrum of makilactone G (84)

AA06795D3.F3'.7_171116154423 #6-12 RT: 0.44-0.93 AV: 7 NL: 1.35E7 T: FTMS + p ESI Full ms [150.00-1000.00] 405.11442 R=127454 100

90 85 80

75 70 65

55 50 45

RelativeAbundance 40 787.24059 R=96025 35 30 25

20 393.02592 539.18709 15 R=133797 R=109893 10 437.14045 765.53185 R=123140 721.50554 809.55779 853.58407 175.03637 361.08848 505.12953 565.16640 897.61033 5 213.01309 290.97871 647.46888 R=97047 R=97398 R=93167 955.65219 R=198973 R=136739 R=114395 R=107310 R=90966 R=88431 R=183906 R=126068 R=103912 R=81930 0 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 1000 m/z

Analysis and interpretation of the 2D-NMR data, notably HSQC and HMBC

(Figure 40), led to the conclusion that 84 corresponded to the structure of makilactone G, and this was verified by comparison with previous data for this compound (Sato et al.,

2009a).

Figure 39. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of makilactone G (84)

(in CD3OD)

(a)

(b)

Figure 40. HSQC (a) and HMBC (b) spectra of inumakiol G (84) (400 MHz, in CDCl3)

(a)

(b)

2. Biological activity of the isolated compounds The cytotoxicity of the isolated compounds (77-84) was evaluated against a panel

of four human cancer cell lines, namely, colon (HT-19), breast (MDA-MB-231), melanoma

(MDA-MB-435), and ovarian (OVCAR3). As shown in Table 7, compounds 78 and 79 exhibited antiproliferative activity against these cell lines, whereas the remaining compounds were inactive (IC50 >10 µM). These data are in agreement with a previous cytotoxic activity report on this compounds against a murine leukemia cell line (Sato et al.,

2009a). However, this is the first report, to the best of the knowledge of the present author on a more comprehensive in vitro antiproliferative examination of these diterpenoids and podolactones against human cancer cells.

Table 7. Cytotoxic activity of isolated compounds

HT-29 MDA-MB-435 MDA-MB-231 OVCAR3 Compound (colon) (melanoma) (breast) (ovarian)

77 >10 >10 >10 >10

78 >10 3.7 6.6 5.2

79 6.3 2.4 4.2 2.9

80 >10 >10 >10 >10

81 >10 >10 >10 >10

82 >10 >10 >10 >10

83 >10 >10 >10 >10

84 >10 >10 >10 >10

E. Conclusion

The small-scale bioactivity-guided isolation of the root sample of Podocarpus neriifolius described in this chapter resulted in the isolation and structure determination of seven known compounds, including four type-B podolactones (77-79, 84) and three totarane-type diterpenes (80-82). Moreover, a new podolactone (83) was also obtained in the course of this study. However, since additional data were required for its complete assignment, a re-isolation of this compound was conducted with a larger amount of the same plant sample, and thus its structural elucidation will be described in the subsequent chapter of this dissertation (Chapter 3). Of these eight compounds, only the podolactones

78 and 79 exhibited cytotoxic activity against the panel of four human cancer cell lines used. Through this study, the biological properties of these compounds against these cell types is being reported for the first time. Therefore, the work described in this chapter resulted in a new contribution regarding the phytochemistry of P. neriifolius and its bioactivity. The discovery of the new compound, 83, sparked an interest on additional investigation of this plant sample, both in terms of its phytochemistry and bioactivity, and these aspects will be discussed in Chapter 3.

Chapter 3. 1D-NMR-guided isolation of further constituents from Podocarpus

neriifolius and monitoring of podolactone biotransformation in fungal cultures

A. Overview of dereplication methods in natural products research

1. General dereplication methods in natural products drug discovery The first chapter of this dissertation (Chapter 1) highlighted the essential role occupied by natural products in the field of drug discovery as valuable and continuous sources of promising lead compounds. While several natural sources have been investigated leading to the ample record of secondary metabolites reported thus far, only the “tip of the iceberg” has been scraped, referring to this vast natural product repository

(Paterson & Anderson, 2005; Cragg & Newman, 2013). Accordingly, numerous natural resources, including terrestrial and marine organisms remain to be investigated chemically and biologically, and thus tremendous opportunities for additional discovery have yet to be explored in the field of pharmacognosy.

1.1. Challenges in natural products drug discovery research and the need for dereplication methods Following a surge towards a “green Eldorado” of industrial pharmacognosy in the

1990s, interest in the field of natural product drug discovery has considerably declined in

the early 2000s in the pharmaceutical industry, as demonstrated by the downsizing and even complete termination of natural products-based programs within “big pharma”, which in turn has paralleled with a decrease in approved new drugs (Li & Vederas, 2009; David et al., 2015; Shen, 2015). Several reasons associated with this decline along with potential solutions to tackle the diverse limitations of natural product-based discovery have been discussed in various reviews (Koehn & Carter, 2005; Lam, 2007; Li & Vederas, 2009;

Kingston, 2011; Henrich & Beutler, 2013; Atanasov et al., 2015). Among these are reasons pertaining to the lack of compatibility of natural product extracts, composed of complex mixtures, for automated high-throughput screening (HTS), as compared to synthetic compound libraries, although the latter have been found to lack the chemical diversity offered by secondary metabolites from organisms. In the race to find new therapeutics, perhaps fueled by the pressure for profit-making, automated systems have been favored to increase the efficiency and “hit rate” during the discovery process. Other reasons include the often difficult access to certain natural resources due to legislation pertaining to ecological conservation and benefit-sharing considerations for bioprospecting in foreign countries (Cordell, 2010; Kingston, 2011), and reduced emphasis in therapeutic drug programs for certain conditions, such as infectious diseases, which do not represent highly profitable projects in comparison to treatments for chronic diseases (Koehn & Carter, 2005;

Paterson & Anderson, 2005; Li & Vederas, 2009). Additionally, natural product isolation and structure determination is usually a difficult and lengthy process, as many secondary metabolites have complex structural features requiring considerable expertise and time to

solve. Thus, the labor-intensive and time-consuming aspects of natural products-based drug discovery have caused a reduced popularity of this field in the pharmaceutical industry. Moreover, issues with the supply of naturally occurring lead compounds represent one of the main impediments for their further development as drug candidates. In most cases, bioactive compounds occur at low concentrations in the producing organisms

(Suffness & Douros, 1981), and thus further clinical development would require harvesting large amounts of raw material, the identification of additional natural sources, or synthetic and semi-synthetic routes to production, all of which representing challenging tasks. In addition to these difficulties, natural product isolation has a “high probability of duplication” (Li & Vederas, 2009), as many such compounds have been characterized over decades of research of agents of natural origin. Thus, the fruitful aspects of natural product research have also led to the frequent re-isolation of known secondary metabolites, which, consequently, has increased the difficulty in making new discoveries, and thus would prevent patenting and profitability.

Several technological advances have been made in the last few decades and continue to be improved to address these limitations, including biological and chemical analytical instrumentation along with the increased exploitation of genetic information, allowing natural products to remain a valuable and competitive source of lead compounds

(Hook et al., 1997; Koehn, 2008; Bugni et al., 2009; Li & Vederas, 2009; Henrich &

Beutler, 2013; Paddon et al., 2013; Atanasov et al., 2015; Harvey et al., 2015; Shen, 2015).

Among these developments, “dereplication” procedures were implemented to increase the

efficiency of the discovery of new and bioactive secondary metabolites (Gaudêncio &

Pereira, 2015; Hubert et al., 2017).

1.2. Definitions of the term “dereplication” As the term indicates, “dereplication”, particularly in natural product drug discovery, is aimed at preventing the redundant isolation (replication or rediscovery) of known compounds. According to a recent chronological account (Gaudêncio & Pereira,

2015), the concept of dereplication in the field of natural products drug discovery can be traced back to 1978 with the application of prescreening methods in the search for new fermentation-derived antitumor lead compounds (Bunge et al., 1978; Hanka et al., 1978).

In these early reports, in vitro bioactivity-guided prescreening of cultured broths through cytotoxicity and antimicrobial assays prior to further in vivo evaluation and secondary metabolite isolation was favored to random submission of samples for animal studies, and active broths were then subjected to fingerprinting steps to reduce the likelihood of rediscovery of known antitumor agents. It came as no surprise, therefore, that the term

“dereplication”, at the time, was defined as “the processes of determining the presence of a known agent in a culture broth” (Hanka et al., 1978), underlining the natural source being investigated. Since then, with the many technological advances in instrumentation and the extended array of natural products being explored, the term dereplication, in pharmacognosy, has comprised a variety of procedures or combination thereof, and has been defined subsequently in a more general manner according to the study material and

objectives (Hubert et al., 2017).

To rapidly prioritize phorbol dibutyrate (PDBu) receptor-binding bioactive plant extracts for the discovery of novel antibiotic compounds by by-passing the traditional bioassay-guided isolation procedure, Beutler and colleagues developed a method combining HPLC-UV detection with an online PDBu assay, and called dereplication “this process of quickly identifying known chemotypes” (Beutler et al., 1990). While emphasizing the importance of biological activity and novelty, dereplication has also been defined as the “process of determining whether an observed biological effect of an extract or specimen is due to known substance” (Clark, 2002). On the other hand, a more involved definition was provided by Harvey and co-workers, with dereplication being “the process of using spectroscopic methods to identify known metabolites during the preliminary screening stage and eliminating further isolation work on already well-studied natural products” (Harvey et al., 2015), specifying the analytical approach used. Another similar definition is one given by Lang and collaborators where “dereplication is a process of differentiating those natural product extracts that contain nuisance compounds, or known secondary metabolites, from those that contain novel compounds that are of interest” (Lang et al., 2008). Finally, Lam gave another and much inclusive definition of the term, stating that “dereplication is the process by which the chemical and biological characteristics of the unknown compounds are compared with the chemical and biological characteristics of known compounds from the databases to eliminate those that have been identified previously” (Lam, 2007). While the above definitions present some small variations, they

essentially share a common purpose, which is to enhance the efficiency of the natural product discovery process in terms of time and labor, as well as the prevention of unnecessary duplication through known compound re-identification.

1.3. Overview of the methods used in dereplication procedures and types of dereplication Dereplication is achieved through various procedures, and these can be classified according to the analytical methods applied, including separation, detection, or structure determination techniques, the biological properties being tested, and the genetic makeup of the organism being investigated (Gaudêncio & Pereira, 2015; Hubert et al., 2017). In some cases, these categories can overlap, as one technique may provide, for example, both detection and information on the structural composition of a test sample. Separation methods include chromatographic techniques such as TLC, HPLC, and flash chromatography (Gaudêncio & Pereira, 2015; Hubert et al., 2017), and thus, different samples can be compared according to their retention times or retention factors. While certain detection techniques are commonly coupled with the above separation methods for dereplication, including UV-vis detectors (e.g., PDA, DAD) and dyeing reagents for TLC development, other techniques, including mass spectrometry and NMR spectroscopy can be described as stand-alone detection methods for dereplication, and they can also provide partial structure determination of test compounds.

Different amalgamations within and between the above categories have been employed in tandem through on-line hyphenation or off-line sequences of individual 98

techniques (Bunge et al., 1978; Hanka et al., 1978; Koehn & Carter, 2005; Konishi et al.,

2007; Gaudêncio & Pereira, 2015; Harvey et al., 2015). For example, early dereplication studies reported involved the combination of biology and chromatography including off- line fingerprinting steps involving antimicrobial/antiproliferative, TLC, UV and HPLC profiles (Bunge et al., 1978; Hanka et al., 1978), the simultaneous use of TLC and biological testing through bioautography (Adhami et al., 2013; Potterat & Hamburger,

2013; Cabral et al., 2016), the use of affinity-bioassay (e.g. PDBu receptor binding assay) followed by TLC (Beutler et al., 1990), and later LC-MS and LC-MS-bioassay (Cordell et al., 1997).

Genetic information can also be used for dereplication as it facilitates the identification of and comparison between natural resources, and this practice has been most common in the taxonomic investigation of microbial strains (Hubert et al., 2017). Thus, genome mining and proteomics-guided approaches can serve as means of dereplication, and organisms most likely to produce unknown secondary metabolites, or other compounds of interest based on their gene clusters and genetic expressions, are given higher priority

(Gaudêncio & Pereira, 2015).

Since dereplication is based on comparison of unknown samples with previously obtained compounds, for the above techniques to be useful for dereplication purposes, the data obtained therefrom must be stored in accessible libraries or databases, which in turn can be consulted for assessment of the test samples. Accordingly, databases represent an essential part of the dereplication procedure. Several databases are available, either

publicly or commercially, and internal databases are also built by individual laboratories.

Depending on the type of data collected (e.g., genomic or spectroscopic), or the source of the natural compounds being characterized (e.g., plant, terrestrial bacteria, marine organisms), examples of databases include the Dictionary of Natural Products,

NAPRALERT, DEREP-NP, ACD/Labs NMR, the Marine Natural Products Database,

MARINLIT, AntiMarin, NIST, MassBank, the International Nucleotide Sequence

Database Collaboration (INSDC), NMRShiftDB, the Global Natural Products Social

Molecular Networking (GNPS) (Corley & Durley, 1994; Konishi et al., 2007; Gaudêncio

& Pereira, 2015; Hubert et al., 2017; Zani & Carroll, 2017).

Furthermore, the ever-evolving computational power has not only permitted the development and storage of the above-mentioned databases, but has also led to the exploitation of empirical data through statistical and algorithmic methods allowing for the development of additional techniques such as computer-assisted structure elucidation

(CASE) and automated DNA sequencing, all of which can assist in dereplication efforts

(Bouslimani et al., 2014; Gaudêncio & Pereira, 2015).

1.4. NMR spectroscopy as a tool for dereplication Nuclear magnetic resonance (NMR) spectroscopy is one of the most practical techniques that offers comprehensive and specific structural information unique to each compound (except enantiomers), and thus, it has been used extensively for dereplication.

Although this methodology has been criticized previously for its lack of sensitivity,

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improvements have been made to address this restriction, including the development of higher field magnets, cryoprobes and capillary NMR probes requiring only small sample volumes and providing high-quality spectra in a reduced time (Lang et al., 2008; Mitova et al., 2008; Bugni et al., 2009). Both one- and two-dimensional NMR spectroscopic techniques have been used for dereplication purposes, with 1H NMR spectroscopy being the most frequently used, owing to its throughput capability (van der Hooft & Rankin,

2016). However, when dealing with crude extracts and mixtures, 1H NMR spectroscopy would most likely be limited by a high degree of complexity due to overlapping signals

(Davis & Bax, 1985; Pauli et al., 2014). While this limitation can be partially addressed by further fractionation of the test samples or additional parameter changes, one may also turn to 13C NMR, or 2D-NMR techniques, such as COSY, TOCSY, HSQC, and HMBC as alternatives (Williamson et al., 2000; Schroeder et al., 2007; van der Hooft & Rankin,

2016; Bakiri et al., 2017; Markley et al., 2017). In addition, hyphenated techniques involving NMR spectroscopy (e.g., LC-SPE-NMR) have been reported as dereplication approaches in natural product isolation (Schefer et al., 2006; Lang et al., 2008; Appendino

& Pollastro, 2010; Pawlus et al., 2013; Wolfender et al., 2013).

2. Overview of dereplication methods for plant secondary metabolite isolation Considering the extensive studies conducted with plant sources, dereplication techniques represent useful, if not necessary, tools for the rapid and efficient discovery of new plant secondary metabolites. Dereplication techniques, thus, have been utilized widely

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in phytochemical investigations. Methods employed include chromatography, MS, NMR, bioassays, and their various combinations, and genome mining (Sumner et al., 2015;

Hubert et al., 2017). As mentioned previously, Beutler and co-workers used HPLC-UV coupled with a receptor-binding assay to identify new phorbol bioactives (Beutler et al.,

1990), while HPLC-MS combined with an estrogen-binding activity was used to prioritize samples from a plant extract library (Schobel et al., 2001). Furthermore, LC-ESI-MS linked to cytotoxic activity against the KB (human oral epidermoid carcinoma) cell line was performed during the isolation work on the stem bark of Artocarpus kemando

(Moraceae) and that of Acnistus arborescens (Solanaceae), resulting in bioactive flavonoids and withanolides, respectively (Minguzzi et al., 2002; Seo et al., 2003).

Similarly, a hyphenated HPLC-MS-bioassay method was used for the study of Aglaia perviridis (rocaglate derivatives) and Elaeocarpus chinensis (cucurbitacins) (Pan et al.,

2013). Dereplication tools have also been employed to target undesired interfering compounds, such as tannins and polysaccharides, as was the case with the screening of the ethanolic extracts of five South African medicinal plants for their potential anti-HIV related activities (Klos et al., 2009). The more recently developed concept of “molecular networking”, initially applied to the dereplication of microbial samples (Yang et al., 2013), has also been extended to applications in the study of plant natural products. These have included not only mapping by MS fragmentation patterns, but also bioactivity and taxonomic input (Allard et al., 2016; Cabral et al., 2016; Klein-Júnior et al., 2017; Olivon et al., 2017; Péresse et al., 2017).

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3. Selective 1D-TOCSY as a tool for natural products dereplication With the development of selective NMR spectroscopic methods over three decades ago (Davis & Bax, 1985; Kessler et al., 1986), one-dimensional analogs to proton correlation experiments, including 1D-total correlation spectroscopy (TOCSY), have proven useful in the structural determination of complex molecules and the identification of chemical entities within mixture preparations, especially those presenting convoluted 1H

NMR spectra. By irradiating selected signals from a 1H NMR spectrum and applying these selective gradient methods, simplified sub-spectra are obtained displaying either the correlation between directly coupled protons (1D-COSY), protons within the same spin network (1D-TOCSY), or those exhibiting spatial interactions (1D-NOESY and 1D-

ROESY). For instance, irradiation of each anomeric proton of the three-ribose containing trinucleotide A2′-P5′A2′P-5′A through 1D-TOCSY resulted in three sub-spectra showing protons from each ribose ring allowing for the respective resonance assignment previously impeded by the highly overlap region on the original 1H NMR spectrum (Davis & Bax,

1985). Selective 1D-TOCSY has also been used successfully for the analysis of structurally related compounds in a mixture (Sharman, 1999). As such, the 1D-TOCSY spectra derived from the irradiation of anomeric protons in a mixture of the agabrittonosides, saponins from the leaves of Agave brittoniana, allowed for the identification of their corresponding sugar moieties, while 1D-ROESY facilitated the assignment of the sugar sequences (Macias et al., 2007), a very similar application to that conducted by Davis and Bax mentioned above.

Moreover, gradient 1D-TOCSY and 1D-NOE were coupled to dereplication approaches

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employing 13C NMR, high-resolution MS data, and facilitated the identification of the marine sesterterpene lactones from two sponges, Lendenfeldia frondosa and a Hyrtios sp.

(Stessman et al., 2002).

Despite the extensive use of selective TOCSY experiments in plant natural product structure determination, application of this technique for dereplication purposes has only recently been reported in the identification of components in food and biofluids (Sandusky

& Raftery, 2005; Sandusky et al., 2011; Papaemmanouil et al., 2015; MacKinnon et al.,

2016).

B. Overview of the use of fungal biotransformation in natural product research

The use of fungi for the biotransformation or bioconversion of naturally occurring secondary metabolites, including various phytochemicals, has been documented in the literature, and this method has afforded new and bioactive molecules. Among the pioneering biotransformation studies was the 11α-hydroxylation of progesterone by a species of the fungus Rhizopus, reported in the early 1950s, which spurred the subsequent use of microbes for the semi-synthesis of steroids (Cannell et al., 2012). Natural products biotransformation describes the use of whole-cell organisms, or that of the enzymes isolated therefrom, to structurally modify previously obtained secondary metabolites.

Biotransformation is a widely used technique that has involved natural substrates belonging to a variety of structural classes and various catalyzing organisms, most of which are microorganisms. Several review articles have been published (Mahato & Garai, 1997; Das

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& Rosazza, 2006; Aldemir et al., 2014; Parshikov & Sutherland, 2014; Takahashi et al.,

2014; Xiao et al., 2014; de Oliveira Silva et al., 2015; Parshikov et al., 2015) highlighting the diverse reactions catalyzed and the respective products obtained, from steroidal hormones to various plant secondary metabolites and other organic compounds. As an alternative to conventional chemical synthesis, this technique has been applied widely in follow-up studies of natural products, ranging from the enzymatically mediated semi- synthesis, exploring the chemical diversity through the generation of analogs for structure- activity relationship studies, to gaining insights into the biosynthetic pathway of a target compound (Cannell et al., 2012). For instance, biotransformation of anthracyclinones, in cultures of wild type and mutant strains of Streptomyces spp., offered insight into the biosynthetic steps involved in the formation of their analogs, notably the glycosylation of these compounds (Blumauerova et al., 1979), while conversion of the water-insoluble ansamycin herbimycin A into its more polar 11-hydroxy-(11-demethoxy)-herbimycin C by

Eupenicillium spp. SD017 resulted in improved solubility and antiproliferative activity of the natural substrate (Xie et al., 2010). The microbial biotransformation of alkaloids from diverse classes has been reported, and these include the dealkylation and hydroxylation reactions, among others (Reighard et al., 1981). As such, O-demethylation by the fungus

Cunninghamella blakesleeana of the isoquinoline alkaloid 7,8-dimethoxy-2-methyl-1-(4′- methoxybenzyl)1,2,3,4-tetrahydroisoquinoline constituted a more efficient route to produce two of its derivatives, in comparison with chemical synthesis (Reighard et al.,

1981). Feeding retinoic acid to cultures of Cunninghamella spp. strains afforded oxidized

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metabolites, including two new compounds, namely, 2-hydroxyretinoic acid, and 2,3- dehydro-4-oxoretinoic acid (Hartman et al., 1990). Similarly, new derivatives were obtained from the fungal bioconversion of the diterpene lactone gelomulide G by

Aspergillus niger (Choudhary et al., 2005), while dimerization of the sesquiterpenoid herbertenediol to form mastigophorenes A and B through an aryl-aryl bond linkage by

Penicillium sclerotiorum offered insights in the biosynthesis of these compounds

(Harinantenaina et al., 2005). Moreover, Zhan and co-workers reported the unprecedented

N-glycosylation and the hydroxylation followed by glycosylation of commercially available aminoanthraquinone and hydroxyanthraquinone by the fungus Beauveria bassiana ATCC 7159 (Zhan & Gunatilaka, 2006). Recently, the same group reported the semi-synthesis of withanolide derivatives incorporating fungal biotransformation into the chemical synthesis process (Wijeratne et al., 2018). While natural products biotransformation has been most commonly conducted using bacterial and fungal material, several biotransformation studies have been performed in plant cell suspension cultures.

For example, biotransformation of digitoxigenin by cell cultures of Digitalis purpurea resulted in the production of digitoxigenone, epidigitoxigenin and its glucoside (Hirotani

& Furuya, 1980), whereas subjecting artemisinic acid and dihydroartemisinic acid to

Catharanthus roseus and Panax quinquefolium cell suspension cultures led to their hydroxylated derivatives (Zhu et al., 2010).

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C. Statement of problem

1. 1D-NMR dereplication-guided purification of P. neriifolius The aforementioned detection and isolation of a new B-type podolactone glucoside

(83) from the roots of Podocarpus neriifolius D. Don (Chapter 2), stimulated further investigation of this plant sample toward the isolation of a larger amount of this compound to enable its full characterization and the purification of additional secondary metabolites on the basis of the rationale presented below.

First, until recently (Zheng et al., 2018), only a very few B-type podolactone glucosides have been reported in the phytochemical literature (Barrero et al., 2003), and thus, the discovery of the above-mentioned glucoside derivative suggested the possible presence of other derivatives for which the structures have yet to be characterized.

Although several podolactones have been reported, their structural diversity mainly rests upon the configuration and the position of the substituents, essentially on the A ring

(positions C-1–C-3) and the side chain on the C ring of the podolactone core (Figure 41).

As illustrated by the newly discovered podolactone glucoside (83), there remains a strong possibility for the discovery of new derivatives. For instance, substituting a glycosidic side chain to any of the three positions, C-1, C-2, or C-3, on ring A of the known B-type podolactones, could give rise to new derivatives. Structural novelty could further be attained with various combinations of other substituents to the already isolated podolactones, and thus this provided a rationale to proceed to further isolation work on this plant sample.

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Figure 41. Overview of structural diversity on the B-type podolactone core.

R1 α-OH, β-OH

R1-R2 epoxy

R2 α -OH, β-OH, glycosyl R -R epoxy 2 3

R3 α -OH, β-OH, glycosyl

R4 alkyl, glycosyl, S-containing,

Second, while the novel nature of the podolactone, 3-deoxy-nagilactone E-2β-O-β-

D-glucoside (83) was not associated with any cytotoxicity against the cancer cell lines tested and at the threshold of concentrations used, its corresponding aglycone, 3-deoxy-2β- hydroxynagilactone E (79), exhibited antiproliferative potency (IC50 <10 µM) across all four cell lines used in the same bioassay. It is, therefore, speculated that a given new compound, while lacking a desired bioactivity, could represent a derivative of a more active new or known secondary metabolite. In this case, it can be stated that substitution within the new metabolite is “masking” the activity of its unsubstituted counterpart. Thus, bioactivity-guided isolation alone may not lead to such new compounds, since, for instance, new polar glycosidic podolactones may be concentrated in the non-cytotoxic aqueous fraction of P. neriifolius, and not be reflective of the activity of their corresponding aglycones. In this case, discarding these inactive derivatives not only reduces the likelihood

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of the discovery of new compounds, but concurrently removes a potential additional source of bioactive constituents. Therefore, exploring these inactive or less active fractions for their podolactone derivative content was undertaken.

To facilitate this endeavor and prioritize fractions in a timely manner, a 1H NMR- spectroscopy-based screening approach was utilized as a means of dereplication procedure using data gathered from the previous small-scale extraction, as described in the second chapter of this dissertation, to prioritize fractions of interest for compound isolation and structure determination. In addition, further detection and isolation of secondary metabolites from the cytotoxic ethyl acetate extract was conducted using this method.

2. Monitoring fungal biotransformation of isolated podolactone with 1D-NMR approach Although podolactone derivatives have been isolated from fungal strains previously

(Barrero et al., 2003), reports on their exogenous biological conversion were not found.

Described herein is one such bioconversion performed in two fungal strains as catalytic agents. During the 1H NMR-guided purification of the aqueous fraction from the large- scale extraction of P. neriifolius root sample mentioned in the previous section, a podolactone derivative, inumakilactone A-15-O-β-D-glucoside (89), was identified as the major component of this fraction (AA06795LG.D2) and as a nearly pure (>95% purity) constituent of its sub-fraction (AA06795LG.D2.2) obtained from Diaion® HP-20-based separation. Although this compound showed no cytotoxicity in the bioassay used, its aglycone, inumakilactone A (78) was active across most of the cancer cell lines tested. 109

Thus, the podolactone glucoside (89) could serve as a source for its active aglycone, which can then be utilized for further testing. However, a previous report of the hydrolysis of this glucosidic derivative, a method mainly applied for its structure elucidation resulted in the opening of the epoxide ring leading to an inactive product (Hayashi et al., 1972a).

Since fungal biotransformation has demonstrated a great potential in the production of bioactive and, in some cases, new secondary metabolite derivatives, and considering the large quantity of the aqueous extract (~ 60 g) in the present study, and the availability of two Penicillium strains, P. concentricum and P. expansum, compound 89, by means of its mother extract, was selected as starting material to be catalyzed in fungal cultures. 1D-

NMR spectroscopy was utilized as a tool for the rapid fingerprinting of the obtained fungal extracts, thus facilitating the detection of podolactones. Therefore, through this fungal- assisted biotransformation procedure, it is hypothesized herein that compound 89, would potentially yield its bioactive aglycone (78) as well as other similarly bioactive analogs through other fungal-catalyzed reaction mechanisms, and these reactions would be monitored through 1D-(1H and TOCSY) NMR spectroscopy-based dereplication.

With an emphasis veered more towards the development and application of this 1D-

NMR-spectroscopy-based scanning method, and while continuing to evaluate biologically the diterpenoid constituents, a large-scale (~2 kg) extraction coupled with phytochemical investigation of P. neriifolius roots as well as the fungal-assisted biotransformation of a podolactone glucoside (89) will be discussed in this chapter.

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D. Experimental

1. General experimental procedures In addition to all instruments and procedures (chromatographic materials, NMR,

MS, UV/vis, solvents) described in Chapter 2 (section C.1), the optical rotation was determined on a modular circular polarimeter (MCP)-150 (software version 1.50; Anton

Paar OptoTech GmbH, Seelze-Letter, Germany).

2. Fractionation and isolation coupled with 1D-NMR spectroscopy 2.1. Plant material The plant material used for the large-scale extraction described in the present chapter came from the same batch as that utilized for the investigation of this plant on a small-scale level. Accordingly, information regarding the plant sample collection can be found in section C.2 of Chapter 2.

2.2. Large-scale extraction of the roots of P. neriifolius D. Don A large-scale extraction of the remaining dried root sample (2.25 kg) of P. neriifolius was performed by extensive percolation in MeOH (5.0-9.0 L  5) in two separate vessels over a period of five days. The resulting percolate was then evaporated in vacuo affording ca. 130 g of a crude MeOH extract (AA06795LG). A portion of this extract

(132.8 g) was re-suspended in a 90% MeOH-H2O solution (550 mL), and partitioned with hexanes (500 mL  6). During this hexanes-aqueous liquid-liquid partition, a third insoluble emulsified layer was formed and collected. The hexane layer (AA06795LG.D1, 111

ca. 6.8 g) and the precipitated emulsified residue (AA06795LG.D0, ca. 4.5 g) were dried and stored. The aqueous residue was evaporated in vacuo, and resuspended in deionized water (500 mL). This aqueous solution was then divided into 100-200 mL aliquots, and each was further partitioned with EtOAc (500 mL  7) resulting in both an aqueous

(AA06795LG.D2, 67.7 g) and an EtOAc (AA06795LG.D3, ca. 40.0 g) fraction. As in the previous partition step, a precipitated residue, insoluble in water and EtOAc but soluble in

MeOH, formed during this separation, and was set aside (AA06795LG.D4, 2.4 g).

Illustrated in Figure 42 is a schematic summary of this large-scale extraction and liquid- liquid partition. Following the partitioning stage, the 1H NMR spectroscopic profile of extracts AA06795LGD1, -D2, and -D3 were acquired, and each partition was then fractionated as follows.

112

Figure 42. Large-scale solvent extraction and partition scheme of P. neriifolius root sample.

Dry powdered roots (2.25 kg)

Percolation in MeOH AA06795LG MeOH extract (~ 130 g) Dried, resuspended in 90% MeOH-H2O

Partitioned with hexanes

AA06795LG.D0 Aqueous precipitated residue residue (~ 4. 5 g) AA06795LG.D2 AA06795LG.D3 AA06795LG.D4 AA06795LG.D1 hexane extract aqueous extract EtOAc extract emulsion (~ 6.8 g) (67.7 g) (~ 40 g) (2.4 g)

1 H NMR spectroscopic scanning

Further fractionation

2.3. 1H NMR-spectroscopy-directed fractionation of the hexane partition (AA06795LG.D1) Following its 1H NMR spectroscopic profiling, the hexane partition

(AA06795LG.D1, 5.4 g), was fractionated on a normal-phase silica gel column (4.1 × 57 cm) in 1:1 hexanes-EtOAc, followed by a MeOH wash, to give 153 fractions combined into eleven fractions (D1.1 – D1.11) (Figure 43), based on similarities in their TLC and

1H NMR spectroscopic profiles. Fractions AA06795LG.D1.4 and AA06795LG.D1.6, according to their respective 1H NMR spectroscopic profiles, were prioritized for further 113

fractionation. Fraction AA06795LG.D1.4 (128.8 mg) was chromatographed on a silica gel column (2.2 × 28 cm) in an isocratic manner, with 9:1 hexanes-EtOAc, resulting in 176 sub-fractions. Following TLC and 1H NMR spectroscopic monitoring, the obtained sub- fractions were combined into nine fractions (AA06795LG.D1.4.1 – 9) (Figure 43).

According to their 1H NMR spectroscopic profiles, the major compounds detected in sub-fractions AA06795LG.D1.4.3, AA06795LG.D1.4.5, and AA06795LG.D1.4.8 were further investigated by additional 1D and 2D-NMR spectroscopic data interpretation, and they were structurally identified as totarol (85), totaral (86) (Ying & Kubo, 1991), and sandaracopimaric acid (87) (Muto et al., 2008), respectively. Fraction AA06795LG.D1.6

(1.0 g) was separated on a reversed-phase C18 column (2.2 × 47 cm) into 267 sub-fractions, and then recombined, in a similar manner as with previous fractions, into a total of twelve fractions (AA06795LG.D1.6.1–12) (Figure 43). Fractions AA06795LG.D1.6.2 and

AA06795LG.D1.6.10 afforded 4β-carboxy-19-nortotarol (82) (Park et al., 2004a), and macrophyllic acid (88) (Bocks et al., 1963; Amaro & Dignora Carros, 1989), respectively, following spectroscopic analysis.

2.4. 1H NMR-spectroscopy-directed fractionation of the aqueous partition (AA06795LG.D2) A portion of the aqueous extract (AA06795LG.D2, ca. 1.3 g) was eluted on Diaion®

HP20 adsorbent resin (3.5 × 50 cm), sequentially with 1:0, 1:1, and 0:1 H2O-MeOH, resulting in the generation of three corresponding fractions, AA06795LG.D2.1 (ca. 579.4 mg), AA06795LG.D2.2 (705.6 mg), and AA06795LG.D2.3 (70.8 mg) (Figure 44). 114

Figure 43. 1H NMR spectroscopy-guided fractionation scheme of the hexanes partition (AA06795LG.D1)

115

In addition, round pellets (AA06795LG.D2.2P, 88.1 mg) were recovered from the

AA06795LG.D2.2 fraction (Figure 44). All three fractions and the precipitated residue were analyzed by 1H NMR spectroscopy. Fraction AA06795LG.D2.2 was prioritized for structural determination resulting in the identification of inumakilactone A-15-O-β-D- glucoside (89, Figure 47) (Hayashi et al., 1972a) as its major component. Comparison of the 1H NMR spectrum of this fraction with that of the precipitated pellets showed an exact overlap.

2.5. 1H NMR-spectroscopy-directed fractionation of the EtOAc partition (AA06795LG.D3) The EtOAc partition was fractionated into different batches to optimize the method for detection of the targeted new podolactone glucoside (83), and other previously unidentified constituents. The dried fractions were then subjected to 1H NMR spectroscopic analysis as a means of dereplication.

2.5.1. Fractionation of AA06795LG.D3 on a reversed-phase C18 column Initially, a portion of the EtOAc partition (AA06795LG.D3; 282.6 mg) was fractionated on a reversed-phase C18 column using a MeOH/H2O step gradient mixture (40,

70, and 100%), affording three fractions, AA06795LG.D3.1 (108.2 mg), AA06795-

LG.D3.2 (39.5 mg), and AA06795LG.D3.3 (136.0 mg). The 1H NMR spectroscopic profile of each fraction collected was acquired, and subsequently, the most polar fraction

(AA06795LG.D3.1, 91.3 mg) was subjected to further separation on a silica gel column 116

(2.5 × 38.7 cm) with an isocratic 15:6:1 CHCl3-MeOH-H2O solvent system to target glycosidic analogs for purification. This chromatographic separation yielded seven sub- fractions (Figure 45). The 1H NMR spectroscopic profiling of fraction

AA06795LG.D3.1.2 (14.5 mg) revealed the presence of inumakilactone A (78) (Park et al., 2004a) as the major component, while its corresponding glucoside 89 was isolated from fraction AA06795LG.D3.1.5 (7.7 mg).

Figure 44. 1H NMR-spectroscopy-guided fractionation of the aqueous partition

(AA06795LG.D2)

(a) Fractionation scheme for the aqueous partition (AA6795LG.D2); (b) Round pellets obtained from AA06795LG.D2.2

(a) AA06795LG.D2 (b) Aqueous extract portion ( ~1.3 g)

Diaion® HP-20

column -1 -2 -3

100% H2O 50% MeOH/H2O 100% MeOH (579.4 mg) (705.6 mg) (70.8 mg)

1 H NMR scanning

117

1 Figure 45. H NMR spectroscopy-guided reversed-phase C18 fractionation scheme of

AA06795LG.D3

EtOAc extract AA06795LG.D3 RP-C 18

D3.1 D3.2 D3.3 (40% MeOH-H O) (70% MeOH-H O) (100% MeOH) 2 2

1 H NMR spectroscopic scanning

Si gel D3.1 sub-fractions (seven)

1 H NMR spectroscopic scanning

2.5.2. Fractionation of AA06795LG.D3 on Diaion® HP-20 resin A portion (2.7 g) of the bulk EtOAc extract was fractionated on Diaion® HP-20 resin (3.5 × 50 cm) with H2O/MeOH mixtures in a gradient fashion (1:0, 1:1, 0:1), followed by a 100% acetone wash, affording four fractions, AA06795LG.D3.Di1-Di4, respectively.

These fractions were analyzed by 1H NMR spectroscopy, and accordingly, fraction

AA06795LG.D3.Di2 was purified by HPLC, using a semi-preparative column (10 × 250 mm) and an increasing H2O/CH3CN gradient, 90:10 to 80:20 in 20 min, 80:20 for 3 min,

80:20 to 0:100 in 1 min, 0:100 for 6 min, 0:100 to 90:10 in 1 min, 90:10 for 3 min, at a wavelength of 234 nm. This afforded four known podolactones, inumakilactone A-15-O-

β-D-glucoside (89) (tR = 14.2 min, 6.2 mg), makilactone F (90) (tR = 14.9 min, 1.8 mg),

118

makilactone G (84) (tR = 16.3 min, 3.0 mg) (Sato et al., 2009a), and 78 (tR = 21.7 min, 5.7 mg).

2.5.3. Fractionation of AA06795LG.D3 through liquid-liquid partitioning An additional fractionation of the EtOAc partition from the large-scale extraction of P. neriifolius was conducted to locate and isolate a larger quantity of the new glucoside, nagilactone G-2β-O-β-D-glucoside (83) than previously obtained. The EtOAc extract (ca.

40 g) was initially re-suspended in MeOH to collect its major constituent, inumakilactone

A (78, 4.7 g), as a white precipitate from the extract. The supernatant was then dried in vacuo and suspended in water for further partitioning with CHCl3, resulting in an aqueous

(AA06795LG.D3.W, 6.8 g), a CHCl3-soluble (AA06795LG.D3.C, 4.9 g), and an emulsified (AA06795LG.D3.R, 9.1 g) fraction. A portion of the CHCl3-soluble fraction

(2.2 g) was chromatographed on a normal-phase silica gel column (2.7 × 55 cm) with

CHCl3/EtOAc mixtures in an increasing gradient (15:1, 4:1, 0:1) and then, with MeOH.

Fourteen fractions (AA06795LG.D3.C1-14) were obtained, and these were screened by 1H

NMR spectroscopy. The most polar fraction eluted with MeOH was then fractionated initially on a reversed-phase C18 column (2.5 × 37 cm) using MeOH/H2O (40, 70, 100%) and further separated by semi-preparative HPLC (10 × 250 mm) using mixtures of

H2O/CH3CN (80:20 to 50:50 in 20 min, 50:50 to 0:100 in 1 min, 0:100 for 5 min, 0:100 to

80:20 in 2 min, 80:20 for 4 min, at 234 nm). This led to the isolation and identification of inumakilactone A (78) (tR = 12.1 min, 6.2 mg), podolactone C (91) (tR = 13.6 min, 7.9 mg),

119

1-epi-makilactone E (92) (tR = 16.3 min, 2.7 mg), along with 3-deoxy-2β- hydroxynagilactone E (79) (tR = 22.6 min, 2 mg). Following dereplication by 1D-TOCSY

NMR spectroscopy (described below), the targeted glucoside (83) (tR = 16.4 min, 2.1 mg) together with 92 (tR = 16.1 min, 1.0 mg) were then obtained from the aqueous partition

AA06795LG.D3.W through HPLC purification in a gradient H2O/CH3CN solvent system

(80:20 to 60:40 in 20 min, 60:40 to 0:100 in 1 min, 0:100 for 5 min, 0:100 to 80:20 in 2 min, 80:20 for 4 min).

2.6. Use of 1D-TOCSY NMR spectroscopy for the localization of the targeted podolactone glucoside (83) in crude fractions The EtOAc extract was scanned by 1H and 1D-TOCSY NMR spectroscopy to confirm its content of nagilactone G-2β-O-β-D-glucoside (83). Using the 1H NMR spectroscopic data collected during the small-scale isolation and partial identification of this new glucoside, the spectral region corresponding to the resonance of its H-2 proton (δH

= 4.07-4.17 ppm, 80 Hz, in MeOD) was irradiated for every 20 Hz, resulting in four 1D-

TOCSY NMR spectra. The obtained spectra were then analyzed for the presence of protons

H-1a, H-1b and H-3a, H-3b, belonging to the same spin system, by comparison with the original glucoside 1H NMR spectrum. In the same manner, both the aqueous and chloroform-soluble partitions obtained from the above-mentioned EtOAc extract were subjected to this spectroscopic scanning, with the δH = 4.07-4.17 ppm region irradiated every 16 and 20 Hz, respectively. Finally, a small amount of the EtOAc extract was fractionated on a reversed-phase C18 column using MeOH-H2O (40, 70, and 100%) for 120

elution, affording fractions AA06795LG.D3.F1-F3. All three fractions were analyzed by

HPLC for the detection of nagilactone G-2β-O-β-D-glucoside (83) using a method by which this compound was initially obtained (Chapter 2. section C.3.). As its HPLC trace of fraction AA06795LG.D3.F1 suggested the presence of the compound of interest, this fraction was subjected to 1D-TOCSY NMR spectroscopy screening as described above as means of confirmation.

3. Characterization of compounds detected and isolated from NMR spectroscopy- based dereplication of P. neriifolius

3.1. Characterization of nagilactone G-2β-O-β-D-glucoside (83)

20 White-pink amorphous solid; [훼]퐷 +31 (c 0.10, MeOH); UV: (H2O) λmax nm (log ε) 201

(3.83), 279.5 (2.87); HRESIMS, positive mode, (m/z 511.2172 [M+H]+ (calcd. for

+ 1 13 C25H35O11 , 511.2174); H and C NMR (Table 8).

3.2. Detection of totarol, totaral, and sandaracopimaric acid (85-87) Detection and identification of compounds 85-87 in the corresponding hexane sub- fractions was performed on the basis of 1D and 2D-NMR spectroscopic data analysis and comparison with reported data.

121

Table 8. Comparison of the 1H and 13C NMR data of nagilactone G-2β-O-β-D-glucoside

(83) and its aglycone (79)

a [Measured in C5D5N at 400 MHz and 300 K. Appeared as dddd (13.5, 8.5, 7.4, 4.9) in

CD3OD]

Position 83 79 (present data)

δC δH δC δH 2.36 (dd, 13.8, 8.7) 2.42 (overlap, dd, 13.5, 1 38.4 1.81 (dd, 13.8, 7.6) 40.4 8.5) 1.80 (dd, 13.5, 7.4) 2 71.4 4.30 (overlap, m) 63.5 4.20 (m)a 3 2.29 (overlap) 2.14 (dd, 13.5, 4.9) 34.3 38.3 2.31 (overlap) 2.48 (overlap, t, 13.5) 4 41.7 42.0 5 41.9 1.77 (d, 4.9) 42.1 1.95 (d, 5.0) 6 72.7 5.07 (dd 4.9, 1.3) 72.8 5.14 (dd, 5.0, 1.4) 7 54.0 4.20 (d, 1.3) 54.1 4.24 (d, 1.4) 8 58.0 57.8 9 158.6 159.1 10 36.5 36.7 11 118.3 6.17 (s) 118.2 6.30 (s) 12 163.5 163.6 14 82.5 4.51 (d, 3.3) 82.6 4.58 (d, 3.3) 15 26.7 1.93 (m) 26.7 1.96 (overlap, m) 16 21.1 1.14 (d, 6.7) 21.1 1.16 (d, 6.7) 17 16.2 1.00 (d, 6.7) 16.3 1.02 (d, 6.7) 18 22.4 1.25 (s) 22.7 1.31 (s) 19 180.9 181.1 20 28.3 1.13 (s) 28.6 1.28 (s) 1′ 103.1 4.99 (d, 7.7) 2′ 75.1 4.07 (t, 8.2) 3′ 78.4 4.27 (overlap, dd, 8.8) 4′ 71.6 4.24 (overlap, dd, 8.8) 5′ 78.5 3.99 (ddd, 8.7, 5.8, 2.2) 6′ 4.36 (dd, 11.6, 5.8) 62.7 4.60 (br d, 11.6)

122

3.3. Characterization of macrophyllic acid (88) White amorphous solid; HRESIMS, positive mode, (observed m/z 653.3814; calcd. for

+ + 1 13 C40H54O6Na [M + Na] , 653.3813) (Figure 90); H NMR (Table 9); C NMR (Table 9).

3.4. Characterization of inumakilactone A-15-O-β-D-glucoside (89) White amorphous solid; HRESIMS, positive mode, (observed m/z 549.1579; calcd. for

+ + 1 13 C24H30NaO13 [M + Na] , 549.1579) (Figure 58); H and C NMR spectrum (Table 10).

3.5. Characterization of makilactone F (90) Amorphous solid; HRESIMS, positive mode, (observed m/z: 405.1156 [M + Na]+ (calcd.

+ 1 13 for C18H22O9Na , 405.1156) (Figure 70.); H NMR (Table 11), C NMR (Table 12).

3.6. Characterization of podolactone C (91) Amorphous solid; HRESIMS, positive mode, (observed m/z: 447.1084 [M + Na]+ (calcd.

+ 1 13 for C20H24O8SNa , 447.1084) (Figure 75); H and C NMR (Table 13).

3.7. 1-epi-makilactone E (92) Amorphous solid; HRESIMS, positive mode, (observed m/z: 423.0823 [M + Na]+ (calcd.

+ 1 13 for C18H21ClO8Na , 423.0817) (Figure 78); H NMR (Table 11); C NMR (Table 12).

123

Table 9. 1H (400 MHz) and 13C (100 MHz) NMR data of macrophyllic acid (88)

(Measured in CDCl3) a b Data for the monomer. Measured in CDCl3. NR: not reported.

Position 88 (Amaro & Dignora Carros, 1989)a 88 (present data) a

b b δC δH δC δH 1 2.12 (1H, d, 12.7) 38.4 NR 40.3 1.30 (1H, m overlap) 2 1.98 (1H, dd, 10.9, 3.1) 21.6 NR 19.9 1.58 (1H, br d, 14.1) 3 2.22 (1H, d, 13.2) 43.4 NR 37.0 1.08 (1H, dd , 13.3, 4.1) 4 40.2 43.6 5 51.9 NR 52.3 1.51 (1H, d, 11.9) 6 2.29 (1H, dd, 13.5, 5.8) 21.6 NR 21.1 2.02 (1H, dd, 13.5, 4.7) 7 3.04 (1H, dd, 16.9, 4.4) 30.1 NR 30.2 2.70 (1H, ddd,16.8, 11.6, 6.4) 8 132.1 135.1 9 134.4 140.7 10 36.8 39.9 11 124.4 6.89 (1H, s) 124.9 6.89 (1H, s) 12 120.5 120.2 13 149.8 150.1 14 140.8 131.9 15 27.8 3.30 (1H, sp, 7.0) 27.8 3.33 (1H, br m) 16 20.2 1.35 (3H, 7.0) 20.2 1.34 (3H, d, 6.9) 17 20.2 1.38 (1H, 7.0) 20.0 1.38 (3H, d, 6.9) 18 28.0 0.77 (3H, s) 28.2 1.37 (3H, s) 19 184.3 185.4 20 22.5 1.05 (3H, s) 22.3 1.02 (3H, s) OH-12 5.05 (1H, s) 5.04 (1H, s)

124

Table 10. Comparison of the 1H and 13C NMR data of inumakilactone A-15-O-β-D- glucoside (89) with reported data and inumakilactone A (78)

Present data recorded in C5D5N at 400 MHz and 100 MHz, respectively and at 300 K.

78 89 89 Position (present data) (Hayashi et al., (present data) 1972a), (in C5D5N )

δC δH δH δC δH 1 55.7 3.66 (1H, d, 4.2) 3.62 (d, 4.0) 55.7 3.62 (d, 4.3) 2 50.8 3.54 (1H, dd, 4.2, 6.2) 3.52 (dd, 4.0, 6.0) 50.8 3.52 (dd, 6.1, 4.2) 3 68.1 4.67 (1H, dd, 6.1, 4.7) 4.65 (d, 6.0) 68.0 4.64 (d, 6.1) 4 48.8 48.7 5 45.3 2.16 (1H, d, 5.2) 2.10 (d, 5.0) 45.2 2.10 (d, 5.2) 6 71.6 5.13 (1H, dd, 5.2, 1.3) Overlap with water 71.6 5.07 (dd, 5.1, 1.2) 7 55.8 5.16 (1H, br d, 1.2) Overlap with water 54.9 5.36 (d, 1.2) 8 57.0 56.6 9 158.0 158 10 37.7 37.6 11 119.0 6.82 (1H, s) 6.70 118.9 6.75 (s) 12 163.1 162.8 14 82.7 4.76 (1H, d, 8.6) 4.52 (d, 10.0) 81.2 4.99 (1H, d, 6.97) 15 63.6 4.37 (1H, m) unassignable 69.5 4.59 overlap 17 20.8 1.58 (3H, d, 6.2) 1.57 (d, 6.0) 15.8 1.57 (3H, d, 6.2) 18 25.0 1.39 (3H, s) 1.50 24.9 1.32 (3H, s) 19 176.4 176.5 20 20.5 1.58 (3H, s) 1.35 20.6 1.51 (3H, s) 1′ 101.6 4.99 (1H, d, 7.7) 2′ 74.7 3.98 (1H, dd, 8.3, 8.3) 3′ 78.2 4.20 (1H, dd, 8.8, 8.8) 4′ 71.6 4.14 (1H, d, 9.2, 8.9) 5′ 78.3 3.94 (1H, ddd, 9.0, 5.8, 2.4) 4.31 (1H, dd, 11.6, 5.8) 6′ 62.8 4.53 (1H, dd, 11.7, 2.6)

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Table 11. 1H NMR data of makilactone F (90) and 1-epi-makilactone E (92)

Measured in 400 MHz, in C5D5N, 300 K.

90 90 Makilactone E (77) Position 92 (present) (Sato et al., 2009a) (present) (Sato et al., 2009a)

δH δH δH δH

1 4.03 (1H, dd, 8.6, 2.8) 4.00 (1H, m, overlap) 4.97 (1H, d, 3.0) 5.28 (1H, overlap) 2 4.32 (1H, m, overlap) 4.32 (1H, m, overlap) 4.69 (1H, br m) 4.99 (1H, dd, 6.4, 2.0) 3 4.35 (1H, d, 6.2) 4.02 (1H, m, overlap) 4.00 (1H, m, overlap) 4.35 (1H, m)

5 2.19 (1H, d, 4.2) 2.18 (1H, d, 4.0) 2.75 (1H, d, 4.2) 2.8 (1H, d, 4.6) 6 5.27 (dd, 4.2, 0.9) 5.26 (1H, dd, 4.0, 1.2) 5.34 (1H, dd, 4.2, 1.2) 5.28 (1H, m, overlap) 7 5.13 (d, 0.9) 5.13 (1H, d, 1.2) 5.18 (1H, d, 1.3) 5.14 (1H, d, 1.3) 11 7.23 (s) 7.23 (s) 6.62 (1H, s) 6.72 (1H, s) 14 4.72 (d, 8.7) 4.71 (d, 8.9) 4.75 (1H, d, 8.5) 4.70 (1H, d, 8.7) 15 4.37 (m) 4.35 overlap 4.39 (1H, m) 4.32 (1H, m, overlap) 17 1.58 (3H, d, 6.0) 1.57 (d, 6.1) 1.57 (3H, d, 6.2) 1.55 (3H, d, 6.3) 18 1.68 (3H, s) 1.67 (s) 1.64 (3H, s) 1.33 (3H, s) 20 1.62 (3H, s) 1.61 (s) 2.00 (3H, s) 1.99 (3H, s)

126

Table 12. 13C NMR data of makilactone F (90) and 1-epi-makilactone E (92)

a Measured 100 MHz, in C5D5N, 300 K. Assigned from HMBC spectrum.

Positi 90 90 Makilactone E (77) 92 on (Sato et al., 2009a) (present) (Sato et al., 2009a) (present)

δC δC δC δC

1 74.0 73.77 63.6 62.3a

2 75.5 75.32 73.1 ~76.0 overlap

3 78.6 78.36 71.4 ~76.0 overlap

5 45.9 45.70 44.6 44.7a

6 44.6 44.31 40.6 40.4

7 72.9 72.64 72.6 72.1a

8 56.2 55.97 55.6 55.1

9 58.5 58.21 57.6 56.6

10 156.9 156.68 155.5 155.1

11 42.3 42.12 42.6 42.2

12 120.8 120.54 119.5 120.1

13 163.8 163.65 163.3 163

14 83 82.75 83.0 82.6

15 63.6 63.40 63.8 63.6

17 21.0 20.85 21.0 20.9

18 22.4 22.24 22.3 24.1

19 177.5 177.4 178.1 180.0

20 17.4 17.21 26.1 25.5

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Table 13. 1H and 13C NMR data of podolactone C (91) (400 MHz)

Position 91 91 (Cassady et al., 1984) (present)

δC (in CD2Cl2) δH (in C5D5N) δC (in C5D5N) δH (in C5D5N) 2.21 (1H, dd, 15.0, 2.0) 2.18 (1H, dd, 14.8, 2.2) 1 31.1 30.3 1.73 (1H, dd, 15.0, 1.0) 1.72 (1H, br d, 14.4) 2 53.0 3.37 (1H, ddd, 4.0, 2.0, 1.0) 51.7 3.36 (1H, overlap, m) 3 55.7 3.26 (1H, d, 4.0) 52.6 3.25 (1H, d, 3.8)

4 (not reported) 42.9 5 40.2 1.80 (1H, d, 5.0) 42.7 1.79 (1H, d, 4.6) 6 73.4 5.03 (1H, dd, 5.0, 1.2) 73.1 5.02 (1H, dd, 4.6, 1.2) 5.23 (1H, d, 1.2) 7 57.4 55.5 5.23 (1H, d, 1.2)

8 58.5 58.2 9 159.3 158.6 10 36.1 35.5 11 116.2 6.19 (1H, s) 116.7 6.20 (1H, s) 12 162.5 162.8 14 83.0 4.83 (1H, s) 83.0 4.84 (1H, s) 15 75.2 73.1 16 27.9 1.85 (3H, s) 27.5 1.84 (3H, s) 3.39 (1H, d, 14.0) 3.4 (1H, d, 13.9) 17 52.0 61.7 3.76 (1H, d, 14.0) 3.75 (1H, d, 13.9) 18 26.1 1.43 (3H, s) 20.8 1.40 (3H, s) 19 180.3 176.8 20 21.7 1.39 (3H, s) 25.4 1.37 (3H, s)

SOCH3 40.2 2.66 (3H, s) 40.1 2.65 (3H, s)

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4. Biotransformation of inumakilactone A-15-O-β-D-glucoside (89) and its aglycone (78) in fungal culture Two previously collected fungal strains, namely, Penicillum concentricum and

Penicillum expansum, were used in separate culture broths, for the biotransformation of inumakilactone A-15-O-β-D-glucoside (in the aqueous extract) and that of its aglycone.

The collection, identification, and culture protocol of these strains on potato dextrose agar are described in Appendices F and G. For each fermentation procedure, 6.5 g of DifcoTM potato dextrose broth were suspended in 250 mL of water, and the mixture was sterilized for 1 h at 250 °C in an EZ10 autoclave (Tuttnauer, Beit Shemesh, Israel). The sterilized broth was then cooled at room temperature for at least 3 h and further treated under UV light for 1 h. To this broth were then added P. concentricum hyphae on two 0.5 × 0.5 cm agar sections from a previously grown agar plate culture. The obtained fungal culture broth was then incubated on a MaxQ 3000 shaker (Thermo Fisher Scientific, Asheville, NC,

USA), continuously rotating at room temperature. After three days, fungal cells from the

P. concentricum culture broth (1 mL) were seeded into freshly made and sterilized potato dextrose broth to develop new cultures for the biotransformation experiments. These new cultures were left to incubate at room temperature on the orbital shaker. Cultivation and fermentation with P. expansum were performed in the same manner as that of P. concentricum described above.

4.2. Source of podolactone for fungal-assisted biotransformation For the biotransformation experiments, the aqueous extract of P. neriifolius 129

(AA06795LG.D2), its sub-fraction (AA06795LG.D2.2), and the aglycone inumakilactone

A (78) obtained from the EtOAc extract (AA06795LG.D3) were added to the appropriate fungal fermentations, as detailed in the following sections.

4.2.1. Pilot biotransformation of P. neriifolius aqueous fraction AA06795LG.D2.2 in Penicillium concentricum potato dextrose broth To a 3-day fermentation of P. concentricum culture (in a 500-mL Erlenmeyer flask, as mentioned above) were added 569.5 mg of the P. neriifolius aqueous extract fraction

® obtained from its 1:1 H2O-MeOH separation over Diaion HP20 resin (AA06795LG.D2.2, section 2.3), and the fermentation was resumed at room temperature. On day 4 following the addition of the plant aqueous fraction, a small amount of the fungal culture (ca. 10 mL) was transferred into a clean 20-mL vial under sterile conditions (in a fume hood and in the presence of an open flame), filtered, and evaporated in vacuo. This crude extract was then re-dissolved in water (5 mL) and partitioned in EtOAc (5 mL × 3), affording both an aqueous and a EtOAc partition, which were later analyzed by 1H NMR spectroscopy. Six days following the addition of AA06795LG.D2.2, the fermentation procedure was terminated. Then, EtOAc (800 mL) was added to the fungal culture (300 mL), and the mixture was filtered. The two layers of the resulting filtrate were separated, and the aqueous layer was further partitioned with EtOAc (500 mL × 3). Both partitions were dried, and their 1H NMR spectroscopic profiles were acquired.

130

4.2.2. Biotransformation of inumakilactone A (78) from the P. neriifolius EtOAc- soluble fraction in a P. concentricum potato dextrose broth A white precipitate (AA06795LG.D3.1P, 332.0 mg) from the EtOAc extract

(AA06795LG.D3), identified as inumakilactone A (78) by comparison of its 1H NMR spectrum to previously obtained data, was added to a 2-day fermentation of P. concentricum. Aliquots (15 mL) of the fungal mixture were collected on days 5 and 9, and the fermentation procedure was halted on day 16. The mixtures collected were filtered and partitioned with EtOAc, and the dried EtOAc partition from each collection was analyzed by 1H NMR spectroscopy.

4.2.3. Biotransformation of the P. neriifolius aqueous extract AA06795LG.D2 in Penicillium concentricum potato dextrose broth To a 2-day P. concentricum fermentation, was added a 10g-sample of the P. neriifolius aqueous extract (AA06795LG.D2). Small extractions (15 mL) of the fermentation were conducted at 2 and 6 days following the addition of AA06795LG.D2.

Each collection (15 mL) was partitioned with EtOAc (25 mL × 3), and the 1H NMR spectra of the obtained aqueous and EtOAc partitions were measured. On day 7 after addition of

AA06795LG.D2, the fermentation procedure was terminated, and the fungal mixture filtered and partitioned with EtOAc. The 1H NMR spectroscopic profile of the resultant

EtOAc partition was then acquired.

Using another batch of fraction AA06795LG.D2 for biotransformation in P. concentricum, two 500-mL Erlenmeyer flasks fermentations were seeded from a 3-day

131

culture. In order to minimize the concentration of fungal metabolites in comparison to that of the biotransformed products, the addition of AA06795LG.D2 samples was conducted one day post-seeding. Two ~250 mg samples of AA06795LG.D2 were transferred in vials and dissolved in 20 mL H2O. Both vials were placed under UV light for 1 h prior to addition to the fermentation flasks. To each flask, labeled I and II, were added 225.4 and 236.5 mg of AA06795LG.D2 dissolved in H2O (20 mL). Small collections (15 mL) of both fermentations were obtained at 2, 3, and 6 days following sample addition, and each fungal mixture was filtered and partitioned with EtOAc (20 mL × 3). The obtained partitions

(aqueous and EtOAc) were analyzed by 1H NMR spectroscopy. On day 8, the fermentation procedure was terminated, and each fungal culture (~250 mL) was extracted (1 L) and partitioned with EtOAc (300 mL × 2). As with the previous collections, 1H NMR spectroscopic data and HPLC profiles of both the aqueous and EtOAc partition from cultures I and II were collected. This fermentation procedure, including a description of the fractions obtained, is summarized in Figure 46.

132

Figure 46. Biotransformation procedure of 250 mg P. neriifolius aqueous extract in Penicillium concentricum culture

4. Aliquot (15 mL) EtOAc Extraction Day 2

5. Aliquot (15 mL) EtOAc 1. Plate to broth 2. Passage 3. Extract added Extraction (P. concentricum) (2 flasks, 3-day) (AA06795LGD2, 250 mg in H O) × 2 Day 3 2 1 (Day 2) H NMR Profile

6. Aliquot (15 mL) EtOAc Extraction Day 6

2 fractions (EtOAc and H2O per extraction) per flask total: 8 fractions per flask 7. Total EtOAc Extraction Day 8

133

4.2.4. Biotransformation of P. neriifolius aqueous fraction AA06795LG.D2 in Penicillum expansum potato dextrose broth A 2-day fermented colony of a P. expansum was sub-cultured into four 500 mL

Erlenmeyer flasks containing 250 mL potato dextrose broth. Three ~250 mg aliquots of

AA06795LG.D2 dissolved in 20 mL H2O were prepared and sterilized under UV light 0.5-

1 h. To flask I were added 240.2 mg of AA06795LG.D2 in solution on the day of sub- culture (day 0). The flask II and III were supplemented with 246.2 and 245 mg of

AA06795LG.D2 in solution, respectively, on day 1 post-sub-culture. A fourth fermentation flask (IV) was not supplemented with AA06795LG.D2, and served as a control. At 2 days post-sub-culture a portion (15 mL) of the fungal culture from each of flasks I and IV was collected and partitioned with EtOAc (20 mL × 3), in each case affording an aqueous and an EtOAc partition, and both were analyzed by 1H NMR spectroscopy. On day 3, a small collection from flask IV was partitioned with EtOAc, while the fermentations from flasks

I and II were terminated with the addition of EtOAc (500 mL) and further partitioned using the same solvent (250 mL × 2). The 1H NMR spectroscopic profiles of all six partitions were acquired subsequently. Finally, on day 4 following the sub-culturing, the remaining fermentations in flasks III and IV were terminated and partitioned as described for the previous cultures, and the 1H NMR spectra of their respective partitions were collected

(Table 14).

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Table 14. Biotransformation procedure of AA06795LG.D2 (250 mg) in Penicillum expansum fungal culture

Three study flasks (I-III) supplemented with PN-D2 (P. neriifolius aqueous extract, i.e.

AA06795LG.D2), and one flask (IV) used as a control. Partial and total extractions performed at various days, as indicated.

Fungus Fermentation flasks P. expansum freezer to potato I II III IV (control) dextrose broth PN-D2 added Subcultures made (250 mg in 20 No PN-D2 No PN-D2 No PN-D2 (2 days) mL H O) 2 PN-D2 added PN-D2 added (250 mg in 20 (250 mg in 20 No PN-D2 mL H O) mL H O) 2 2 Partial extraction Partial extraction

(15 mL, 2 days) (15 mL, 2 days) Total Total extraction Partial extraction extraction (3 days) (15 mL, 3 days) (2 days) Total Total extraction extraction (4 days) (3 days)

In a new fermentation batch, a 3-day fermented culture of P. expansum was seeded into two 500 mL Erlenmeyer flasks containing 250 mL potato dextrose broth. Two ~2 g samples of AA06795LG.D2 were dissolved each in 50 mL H2O, and allowed to stand under

UV light. These samples were added each to one of the fungal culture in flasks I and II on 135

days 0 and 1 of the sub-culture, respectively. The fermentation in flask I was monitored on days 2, 7 and 14 post-sample addition through 1H NMR spectroscopic profiling of the aqueous and EtOAc partitions from these small collections, and it was terminated on day

16. In turn, fermentation in flask II was left undisturbed until its termination on day 15.

Small collections (15 mL) from the fungal culture in flask I were each extracted with 50 mL EtOAc, filtered, and the aqueous layer was partitioned with the same solvent (20 mL

× 2). Upon termination, cultures from each of flasks I and II were extracted with 1-1.5 L

EtOAc during filtration, and further partitioned with the same solvent (500 mL × 2). The resulting partitions from each fermentation were then analyzed by 1H NMR spectroscopy.

4.3. 1H NMR spectroscopy monitoring of the biotransformation procedure As mentioned earlier, the progression of fungal biotranformation was monitored in each set of fermentation experiments at various time intervals, and this was accomplished through the measurement and analysis of the 1H NMR spectra of the partitions obtained from the fungal culture mixture collected at these time periods. The resultant 1H NMR spectra were compared with the corresponding in-house NMR data of previously isolated podolactones, specifically inumakilactone A glucoside (89) used as the “reactant” for the biotransformation, and its aglycone (78).

4.4. 1D-TOCSY-NMR-spectroscopy-based structural confirmation In addition to 1H NMR spectroscopic comparison, 1D TOCSY NMR spectroscopy

136

experiments were performed to confirm the presence of inumakilactone A (78) as a biotransformation product in two of the fungal fermentation partitions, namely, PN-D2-

PC-6d-E (EtOAc partition from the biotransformation of 10 g plant aqueous extract in P. concentricum at 6-day endpoint) and PN-D2-PE-15d-W (aqueous partition from the biotransformation of 2 g plant aqueous extract in P. expansum at 15-day endpoint). Key 1H

NMR resonance signals differentiating the podolactone glucoside (89) and its aglycone, notably the methyl doublet signals corresponding to H3-16, or the methine doublet analogous to H-14, were irradiated, and the obtained 1D-TOCSY NMR spectra were overlaid with the 1H NMR spectra of both podolactones, recorded in the same solvent

(CD3OD) as a means of comparison. Furthermore, 78 was isolated as a white precipitate from the fungal mixture, and its NMR and HRESIMS data were collected to further supplement the analysis above.

4.5. Identification and isolation of inumakilactone A (78) from fungal biotransformation Both the HRESIMS and the NMR spectroscopic data of the fungal biotransformation-derived aglycone, inumakilactone A (78), were identical to previously obtained data. HRESIMS, positive mode, observed m/z 387.1040 [M + Na]+ (calcd for

+ C18H20O8Na , 387.1050); NMR data, Tables 1 and 2.

137

5. Biological evaluation 5.1. Cytotoxicity assay in a panel of four cancer cell lines Fractions obtained from the liquid-liquid partitioning of P. neriifolius roots crude methanol extract (AA06795LG.D0-D4), along with their sub-fractions AA06795LGD1.4,

AA06795LGD1.6, AA06795LGD2.2, AA06795LGD2.3, and AA06795LGD3.1-3 were evaluated for their cytotoxic potential against a panel of four cancer cell lines, namely, the human melanoma (MDA-MB-435), ovarian (OVCAR3), colon (HT-29), and breast

(MDA-MB-231) cancer cells, as described in Chapter 2 section 5.2 and Appendix C.

Evaluated in the same bioassay were the water and chloroform-soluble fractions

(AA06795LGD3.W and AA06795LGD3.C) and sub-fractions obtained from the latter through silica gel separation (AA06795LGD3.C.1-14). In addition, compounds 85-89, along with 91 and 92 were tested in this in vitro cytotoxicity assay. This work was carried out by Mr. A. A. Czarnecki from the laboratory of Dr. J. E. Burdette at the College of

Pharmacy, University of Illinois at Chicago.

5.2. In vivo hollow fiber assay Following its isolation as a precipitate from the crude EtOAc extract of P. neriifolius, inumakilactone A (78) was submitted to the Burdette laboratory group at the

UIC College of Pharmacy, Chicago, IL, for testing in vivo in the hollow fiber assay. Prior to submission, the test sample was re-evaluated in vitro in three human cancer cell lines

(MDA-MB-435, OVCAR3, and MDA-MB-231), and based on the observed cytotoxic activity (Appendix D), these cancer cells were then inoculated into polyvinylidene fluoride 138

(PVDF) fibers, later implanted into athymic mice, according to procedures previously reported (Hollingshead et al., 1995; Mi et al., 2009; Ren et al., 2017) and briefly described in Appendix D.

5.3. Insecticidal activity evaluation

Inumakilactone A (78) and its 15-O-β-D-glucoside (89) were evaluated for their potential mosquito antifeedant property against the Aedes aegypti mosquito strain. This work was performed in the laboratory of Dr. Peter M. Piermarini, at the Department of

Entomology, The Ohio State University, Ohio Agricultural Research and Development

Center, Wooster, OH, USA, using the biological assay protocol detailed in a recent publication (Inocente et al., 2018).

5.4. Cytotoxicity of fungal fermentation extracts The aqueous extract of P. neriifolius together with the water and ethyl acetate- soluble fractions obtained from the fermentations procedures described in sections 4.2.3 and 4.2.4 were assessed for their cytotoxic activity against the ovarian and breast cancer cell lines A2780 and MCF-7, respectively. This biological testing was performed by Mr.

Choon Yong Tan and Ms. Fengrui Wang in the laboratory of Dr. L. Harinantenaina

Rakotondraibe, Division of Medicinal Chemistry and Pharmacognosy, The Ohio State

University. The protocol used for each bioassay is summarized in Appendix E.

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E. Results and Discussion

1. Structure elucidation of nagilactone G-2β-O-β-D-glucoside (83) Compound 83 was isolated as a white solid. Its HRESIMS obtained in the positive mode, gave a protonated molecular ion peak at m/z 511.2172 [M+H]+ (Figure 47), which

+ 1 suggested a molecular formula of C25H34O11 (calcd for C25H35O11 , 511.2174). The H and

13C NMR data of 83 were very similar to those of 79 (Table 8). The 13C NMR spectrum

(Figure 49) displayed signals at δC 180.86 (C-19) and 163.52 (C-12) corresponding to the

γ and δ-lactone carbonyls, along with the signals at δC 158.6 (C-9), 118.3 (C-11), 57.68 (C-

8), and 53.98 (C-7) distinctive of the conjugated 7α,8α-epoxy-9(11)-enolide, indicating that this compound has a B-type podolactone core (Hayashi et al., 1980; Sato et al., 2009).

In addition, inspection of the 1H NMR spectrum (Figure 49) revealed the presence of resonance signals at δH 6.17 (s), 5.07 (dd, J = 1.2, 5.0 Hz), 4.20 (d, J = 1.2 Hz), and 1.77

(d, J = 5.0 Hz), typical for protons H-11, H-6, H-7, and H-5 of this class of podolactone, respectively. The major differences between the NMR spectroscopic data of 78 and 83 were found in the additional resonances in both the 1H and 13C NMR spectra of the latter.

Six additional carbon signals characteristic of a β-glucopyranosyl moiety (C-1′-C-6′) resonated at δC 103.1, 75.1, 78.4, 71.6, 78.5, and 62.7, and this was confirmed by the large coupling value corresponding to its anomeric proton H-1′ (δH 4.99, J = 7.8 Hz) observed in the 1H NMR spectrum.

140

Figure 47. HRESIMS spectrum of nagilactone G-2β-O-β-D-glucoside (83)

Intens. 32100_AA06795D3_F2_1_H5_SID_4M_ESI_pos_000001.d: +MS x107 1+ 511.21718 HRESIMS full spectrum

1+ 285.27875 1

2+ 257.24746

1+ 1+ 419.08737 326.30531 1+ 767.45700 1+ 1+ 349.16452 637.30505 3+ 533.19916 2+ 1+ 187.20542 795.48835 10+ 391.22987 5+ 857.36948 663.45385 12+ 9+ 3+ 570.93007 714.55860 952.52151 133.23114 902.24963

0 100 200 300 400 500 600 700 800 900 m/z

Figure 48. MS/MS spectrum of nagilactone G-2β-O-β-D-glucoside (83)

Intens. 32100_AA06795D3_F2_1_H5_SID_4M_ESI_pos_QCID_511_5eV_000001.d: +MS2(qCID 511.21715) x108

1+ -162.05266 349.16449

1.50

1.25

1.00

0.75

0.50 Loss of glucose - C6H10O5

0.25 1+ +0.00000 511.21715

1+ 1+ 5+ -226.06868 -180.06328 -324.01173 285.14847 331.15387 187.20542

0.00 150 200 250 300 350 400 450 500 550 600 m/z

141

Figure 49. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of nagilactone G-2β-

O-β-D-glucoside (83)

(Measured in C5D5N)

(a)

(b)

142

Figure 50. 1H (400 MHz) NMR spectrum of nagilactone G-2β-O-β-D-glucoside (83) in

CD3OD

Moreover, the presence of a fragment peak at 349 [M- 162] in the MS/MS spectrum of 83, suggesting the loss of a glucopyranosyl unit (Zheng et al., 2018) (Figure 48), corroborated the above analysis. Based on the above evidence, it could be suggested that

83 is the 2-O-glucosylated derivative of 78, as the only possible points of substitution include the A ring, the B ring at C-7, or the side chain at C-14. Since C-7 was protonated,

C-14 bore an isopropyl group identical to that of the aglycone (78), and the A ring protons from both compounds shared a similar splitting pattern, the substituent was most likely attached to C-2. This was confirmed by key HMBC correlations between H-2 and the anomeric carbon (Figure 51 and 52). Finally, comparison of the 13C NMR spectroscopic data of 1 and 2 showed a D-glycosylation shift of +8 ppm at C-2 (63.45 ppm in 1 to 71.4

143

ppm in 2), and further shift values of -2.0 and -3.9 ppm for C-1 (40.38 ppm in 1 to 38.37 ppm in 2) and C-3 (38.26 in 1 ppm to 34. 32 ppm in 2), respectively (Kasai et al., 1977).

Therefore, the new compound 83 was fully assigned as nagilactone G-2β-O-β-D-glucoside.

Figure 51. HSQC (a) and HMBC (b) spectra of 83

144

Figure 52. Key HMBC (a) and NOESY correlation for 83

(a)

(b)

2. Detection and identification of compounds obtained using dereplication methods The large-scale extraction of P. neriifolius root sample reported in this dissertation was conducted to re-isolate the partially assigned new podolactone glucoside, nagilactone

G-2β-O-β-D-glucoside (83), allowing its complete structural characterization, and to identify potential new and bioactive podolactone derivatives from this plant. To facilitate this endeavor, data obtained from compounds isolated in the small-scale phytochemical study of this plant (Chapter 2) were used in the form of an in-house database serving as a

145

tool for dereplication. In particular, key 1H NMR spectroscopic signals corresponding to structural patterns germane to the B-type podolactone and totarane-type diterpene classes deduced from previously isolated compounds were utilized to screen extracts and fractions obtained from the large-scale isolation process. Hence, a 1H NMR spectroscopic screening method was applied starting with the extracts obtained from the liquid-liquid partitioning of the crude methanol extract of the plant sample in this study. The dereplication method expedited the prioritization of the plant fractions of interest, leading to the identification of targeted compounds, and the detection of secondary metabolites not present in the in-house database. Additionally, the large-scale extraction of P. neriifolius root sample conducted herein provided a means to implement a dereplication method using the in-house database generated from spectroscopic data, specifically from 1D-NMR spectroscopy, of compounds identified in the small-scale investigation of this plant sample described previously. Furthermore, by using 1D-NMR spectroscopy as a means of dereplication, this large-scale study also permitted this method to be validated towards these isolation purposes. The results obtained from the fingerprinting of the plant extracts and their respective subsequent purification are described in the following sections. First, the NMR- guided isolation followed by structural characterization and biological activities of isolated compounds (Figure 53) from the plant will be discussed. Then, the results of the fungal biotransformation experiments based on the application of the 1D-NMR spectroscopy dereplication method will be addressed.

146

Figure 53. Additional compounds isolated during the large-scale isolation of P. neriifolius

2.1. 1H NMR-guided fractionation of the P. neriifolius aqueous extract (AA06795LG.D2) for the identification of glycosidic podolactone derivatives Owing to its polar nature, the aqueous partition (AA06795LG.D2) obtained from the crude methanol extract of P. neriifolius root sample was selected as a candidate sample to screen for glycosidic podolactone derivatives, including the new podolactone glucoside, nagilactone G-2β-O-β-D-glucoside (83). The 1H NMR spectrum of this aqueous partition, measured in CD3OD, showed clear characteristic signals indicative of the presence of a podolactone analog as the major compound. These signals included resonances at δH 6.5

(s), 5.0 (dd, 5.2, 1.1), 4.8 (d, 1.1), 2.1 (d, 5.2 Hz), and 1.4 (d, 6.2 Hz), characteristic of H-

11, H-6, H-7, H-14, H-5, and the methyl proton signal at position C-16, respectively 147

(Figure 54). Furthermore, two overlapped doublet signals resonating at δH 4.5 ppm, both possessing similar J values (~ 7.7 Hz), indicated the probable presence of a β-anomeric glucosidic proton and a H-14 of B-type podolactone core, while an additional doublet appearing at δH 5.1 ppm (~3.6 Hz) suggested a potential α-anomeric proton of a glycosidic group. In addition, multiple resonances in the δH 3.0-4.1 ppm region could be attributed to glycosidic moieties present in this extract (Figure 54).

The 1H NMR fingerprinting of the fractions obtained from the Diaion® HP-20 resin purification of this aqueous partition revealed, not only a clear separation (Figure 55), but more importantly, confirmed the proposed structural composition mentioned above. As expected, the observed glycosidic constituents eluted in the most polar fraction

(AA06795LG.D2.1). This is illustrated by the corresponding resonance signals within the range δH 3.0-4.1 ppm, δH 4.5 (d, J = 7.8 Hz) and at δH 5.1 (d, J = 3.6 Hz) for potential α- and β-anomeric glycosidic protons, respectively, in the measured 1H NMR spectrum for this fraction (Figure 56a). On the other hand, the 1H NMR spectroscopic profiling of the least polar fraction (AA06795LG.D2.3) obtained from the aqueous partition showed no major distinctive peaks, and instead suggested the presence of minor aromatic peaks in the downfield spectral region (Figure 56b). Therefore, the second fraction (AA06795G.D2.2), seemingly containing the compound class of interest, was further investigated, as described in the following section.

148

Figure 54. 1H NMR spectroscopic profile of the P. neriifolius aqueous extract

(AA06795LG.D2)

Showing characteristic resonance signals for a B-type podolactone. Recorded in CD3OD at 400 MHz and at 300 K.

H3-16

H3-20 H3-18

H-7 H-11 H-6 H-5 Aqueous extract AA6795LG.D2

Figure 55. 1H NMR spectra of AA06795LG.D2 and its fractions from Diaion® HP-20 resin separation.

Recorded in CD3OD at 400 MHz and at 300 K.

AA06795LG.D2.3 (100% MeOH)

AA06795LG.D2.2 (50% MeOH-H2O)

AA06795LG.D2.1 (100% H O) 2

AA06795LG.D2

149

Figure 56. 1H NMR profiling of AA06795LGD2 with its first (D2.1) and last fractions

(D2.3) from Diaion® HP-20 resin separation

1 (a) Overlay of H NMR spectra of AA06795LG.D2 and AA06795LG.D2.1. *δH 4.5 (d, J = 7.8 Hz)

1 and **δH 5.1 (d, J = 3.6 Hz), potential α- and β-anomeric glycosidic protons, respectively. (b) H

NMR spectra overlay of AA06795LG.D2 and AA06795LG.D2.3 shows no distinct peaks in the

latter. Recorded in CD3OD at 400 MHz and at 300 K.

(a) (b) * **

AA06795LG.D2.3 AA06795LG.D2.1

AA06795LG.D2 AA06795LG.D2

2.1.1. Identification of inumakilactone A-15-O-β-D-glucoside (89) The mid-polarity fraction (AA06795G.D2.2) showed a nearly homogenous content

with an approximate purity of >95%, based on its 1H NMR spectroscopic profile, and the

spectrum displayed all the signals corresponding to the key characteristic peaks of a

podolactone core described previously (see section 2.1. above), in addition to observed

glucosidic signals (Figure 57). A solid residue precipitated out of this fraction in the form 150

of round pellets, and these were used, along with the mother liquor, to acquire the spectroscopic data for the structure determination of the corresponding podolactone derivative. While the pellets constituted an excellent sample for NMR spectroscopy measurement of compound 89, another batch of this compound, isolated at higher purity was obtained in pure form as a white amorphous solid (AA06795LG.D3.1.5), see Results

+ section 2.2.1 below). Its HRESIMS gave an ion peak at m/z 549.1579 ([M+Na] , calcd for

+ C24H30NaO13 , 549.1579), indicative of a molecular formula of C24H30O13 (Figure 58).

Figure 57. Overlay of the 1H NMR spectra of AA06795LGD2 and its fraction

AA06795LGD2.2

Recorded in CD3OD at 400 MHz and at 300 K.

AA06795LG.D2.2

AA06795LG.D2

151

Figure 58. HRESIMS spectrum of inumakilactone A-15-O-β-D-glucoside (89)

AA06795LGD3_1_5_inumaki15Glc_2018-1-27_180127161319 #83 RT: 1.99 AV: 1 NL: 8.17E7 T: FTMS + p ESI Full ms [150.00-800.00] 549.15790 R=28600 100 95 90

65 60 55

RelativeAbundance 40 557.91949 35 R=23904 30 25 20

15 581.18372 R=27801 10 5 177.89725 207.09914 283.15149 337.15973 365.12317 413.21198 443.22260 491.15268 535.17883 611.17651 651.32935 683.35675 761.24231 R=46604 R=46904 R=40904 R=36401 R=34801 R=33001 R=33104 R=32204 R=29101 R=27101 R=26401 R=24001 R=23601 0 150 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

The 1H NMR spectroscopic data collected for the pellets formed as precipitates

from this aqueous fraction matched that of its mother liquor, and the NMR data of a sample

of these pellets were then used for the structure determination of 89. These data were

closely similar to those recorded for inumakilactone A (78) (Ito et al., 1968; Park et al.,

2004a), as demonstrated by values in Table 10. However, like compound 83 (see section

1.), the NMR spectroscopic data of 89 possessed additional signals attributed to a

13 glucosidic moiety according to the characteristic signals in the C NMR spectrum (δC

101.6, 74.7, 78.2, 71.6, 78.3, and 62.8). Furthermore, the glucose unit was assigned as

having a β-configuration based on the coupling constant of the anomeric proton (δH 4.99,

d, J = 7.7 Hz) (assigned with HSQC data, Figure 59).

152

Figure 59. HSQC data for inumakilactone A-15-O-β-D-glucoside (89)

The observed chemical shift differences between 78 and 89 at C-15 (δC 63.6 ppm in 78 vs. δC 69.5 ppm in 89) and C-16 (δC 21.1 ppm in 78 vs. δC 15.8 ppm in 89) corresponding to the glucosylation shifts of +5.9 and -5.3 ppm, respectively, and suggested that the glucose moiety is attached to the C-15 on the podolactone core. Cross peaks between H-15 and the anomeric carbon of the glucopyranosyl moiety in the HMBC spectrum confirmed the position of this glucopyranosyl to be at C-15 (Figure 60).

Therefore, compound 89 was identified as the known compound, inumakilactone A-15-O-

β-D-glucoside (Hayashi et al., 1972a; Sato et al., 2009a). However, in the first report on the structure of this compound in 1972 by Hayashi et al., the 1,2-epoxy group on the A ring was assigned with an α-configuration, but this was later corrected by inference from the X- ray crystallographic structure of its aglycone, inumakilactone A (78) (Godfrey & Waters,

153

1975) to the structure later reported by Sato and colleagues (Sato et al., 2009a) and in the present study.

Figure 60. HMBC data for inumakilactone A-15-O-β-D-glucoside (89)

(a) HMBC spectrum recorded in C5D5N at 400 MHz and at 300 K. (b) Key HMBC correlations

(a)

(b)

154

It is worthy of mention that although the more recent publication (Sato et al., 2009a) reported this compound as a known compound, only the first isolation chemistry publication (Hayashi et al., 1972a) reported the 1H NMR spectroscopic data of 89 (Table

10). Hence, the 13C NMR spectroscopic data are reported herein for the first time.

The 1H NMR-guided investigation of the aqueous extract of the root sample of P. neriifolius allowed the prioritization of the second fraction as a source of podolactone glucoside, as predicted, and this led to the identification of a podolactone derivative, 89, which represented the major constituent of this extract.

2.2. 1D-NMR-dereplication-guided fractionation of the ethyl acetate-soluble fraction (AA06795LG.D3) As indicated in section 2.5., the large-scale fractionation of the ethyl acetate-soluble partition of P. neriifolius root sample was conducted in several batches using different chromatographic methods to identify the most efficient method toward the isolation of the newly discovered podolactone glucoside, and other derivatives belonging to this structural class not identified in the small-scale isolation study.

1 2.2.1. H NMR-complemented fractionation of AA06795LG.D3 by reversed-phase C18 chromatography

The polar fraction (AA06795LG.D3.1) eluted from in the 40% MeOH/H2O showed the presence of podolactone and glycosidic moieties as evidenced by its 1H NMR spectroscopic profile (Figure 61). As highlighted in Figure 61, major signals resonating

155

at 2.1, 5.0, 6.5, 1.3, 1.4, and 1.2 ppm and equivalent to protons at positions H-5, 6, 11, 16,

18, and 20 of a B-type podolactone core, in addition to a convoluted ~3.0-5.0 ppm spectral region, suggested the presence of podolactone and potential glucosidic derivative(s) in

AA06795LG.D3.1. 1H NMR fingerprinting of the sub-fractions obtained from its purification by silica gel allowed the isolation of inumakilactone A-15-O-β-D-glucoside

(89) in pure form in the fifth sub-fraction, as its 1H NMR spectrum perfectly overlapped with that of the AA06795LG.D2.2, except the former was devoid of detectable impurity

(Figure 62), and its HRESIMS equally matched with the previously acquired ion peak corresponding to that of 89. Similarly, the 1H NMR spectroscopic profile of fraction

AA06795LG.D3.1.2 also matched that of the previously isolated inumakilactone A (78)

(Figure 63) (Park et al., 2004a). While the use of NMR spectroscopy enhanced the isolation process through dereplication leading to the detection of a podolactone glucoside from this fraction, 1H NMR spectroscopic screening of the sub-fractions did not suggest the presence of 89. However, it is highly probable that the apparent absence of the target compound in these fractions was due to an insufficient amount of this compound for detection within the starting mother fraction or that it appeared below the limit of detection for the method used.

156

Figure 61. 1H NMR spectrum of fraction AA06795LG.D3.1 including key podolactone core signals

Recorded in CD3OD at 400 MHz and at 300 K.

H -16 (d, ~6) H-6 (dd, ~5, ~1.2) H -18 (s) 3 H -20 (s) H-5 (d, ~5) 3 3 H-11

(s)

Figure 62. Overlay of 1H NMR spectra of AA06795LG.D2.2 and AA06795LG.D3.1.5

(89)

Recorded in CD3OD at 400 MHz and at 300 K

AA06795LG.D2.2

AA06795LG.D1.3.5

157

Figure 63. Overlay of 1H NMR spectra of AA06795LG.D3.1.2 and 78

Recorded in CD3OD at 400 MHz and at 300 K

(78)

AA06795LG.D1.3.2

2.2.2. 1H NMR-spectroscopy and HPLC-guided purification of AA06795LG.D3Di2 As with previously described partitions and fractions from the large-scale fractionation of P. neriifolius crude MeOH extract, 1H NMR fingerprinting of the three fractions obtained from the Diaion® HP-20 separation of a second batch of the EtOAc partition (AA06795LG.D3) allowed the prioritization of fractions AA06795LG.D3.Di2 and AA06795LG.D3.Di3, as both their 1H NMR spectra contained signals characteristic of the podolactones, while fractions AA06795LG.D3.Di1 and AA06795LG.D3.Di4 were not investigated further (Figure 64).

In addition to the NMR signals, analysis of AA06795LG.D2.Di2 by HPLC confirmed the presence of peaks possessing UV spectral pattern comparable to

158

podolactones previously obtained (Figure 65). Peaks collected from the HPLC purification of this fraction were further examined by NMR spectroscopy and HRESIMS, and three of these corresponded to compounds identified in prior isolation conducted in this study, namely, AA06795LG.D2.Di2.H7 (inumakilactone A-15-O-β-D-glucoside, 89, Figures 65 and 66), AA06795LG.D2.Di2.H10 (makilactone G (84, Figures 65 and 67), and inumakilactone A (78, Figures 65 and 68) (Hayashi et al., 1972a; Park et al., 2004a; Sato et al., 2009a).

Figure 64. Overlay of 1H NMR spectra of AA06795LG.D3 and its Diaion® HP-20 fractions

Recorded in CD3OD at 400 MHz and at 300 K.

AA06795LG.D3.Di4 (100% acetone)

AA06795LG.D3.Di3 (100% MeOH)

AA06795LG.D3.Di2 (50% MEOH/H2O)

AA06795LG.D3.Di1

(100% H2O)

AA06795LG.D3

159

Figure 65. HPLC trace of fraction AA06795LG.D3.Di2 and UV pattern of selected peaks.

A: (89); B: (90); C: (84); D: (78).

A B

C D

D AA06795LG.D3.Di2 A C

160

Figure 66. Determination of AA06795LG.D3.Di2.H7 (89) by NMR spectroscopic dereplication

1 Overlay of H NMR spectra of AA06795LG.D3.Di2.H7 and 89. Recorded in CD3OD at

400 MHz and at 300 K.

AA06795LG.D3.1.5

AA06795LG.D3.Di2.H7

Figure 67. Determination of AA06795LG.D3.Di2.H10 (84) by NMR spectroscopic dereplication

1 13 Overlay of H (400 MHz) (a) and C (b) (100 MHz) NMR spectra of

AA06795LG.D3.Di2.H10 and previously obtained (84) Recorded in CD3OD and at 300 K.

(a)

AA06795LG.D3.Di2.H10

AA06795D3.F3′.7 (89)

Continued

161

Figure 67 Continued.

(b)

AA06795LG.D3.Di2.H10

AA06795D3.F3′.7

(89)

Figure 68. Determination of AA06795LG.D3.Di2.H14 (78) by NMR spectroscopic dereplication

Overlay of 1H NMR spectra of AA06795LG.D3.Di2.H14 and previously obtained

AA06795LGD3P (78). Recorded in CD3OD at 400 MHz and at 300 K.

AA06795LG.D3P

AA06795LG.D3.Di2.H14

The fourth peak (AA06795LG.D3.Di2.H8, Figure 65, peak B), also exhibited

162

spectroscopic features typical of the podolactones, and its 1H NMR spectrum was very similar to that of the above-mentioned peak C (Figure 65, makilactone G, 84). The NMR spectra of AA06795LG.D3.Di2.H8 did not match previously collected data from this study, and thus additional data were collected to determine its structure, as detailed in the section that follows.

2.2.2.1. Identification of makilactone F (90) Compound 90 was isolated as a brownish amorphous solid, and it was assigned the molecular formula of C18H22O9 on the basis of its HRESIMS that displayed an ion peak

[M+Na]+ appearing at m/z 405.1156 (Figure 70). This molecular weight and the ion peak observed were identical to those corresponding to makilactone G (84, Chapter 2, section

1.8), suggesting that these two compounds are isomers. Comparison of their 1H NMR data indicated that these compounds share the same planar structure. However, variations in the resonances of several signals in the 1H NMR spectrum suggested that differences between these two compounds resided in the configuration of the protons on their A ring. These chemical shift differences and the corresponding affected protons are highlighted in Figure

69. As exhibited in this Figure, the olefinic proton H-11 shifted downfield (6.63 ppm) in

90 as compared with the related makilactone G (84) (δH 6.21 ppm). On the other hand, both the singlet peak at δH 1.46 ppm and the doublet at 2.35 ppm corresponding to the methyl protons at C-20 and the methine at C-5 84, respectively, shifted upfield (H3-20 at

1.25 ppm, and H-5 at 2.03 ppm) in compound 90. Protons of the A ring followed similar

163

trends as shown by their significant upfield shifts [H-1 (4.1 ppm in 84 vs. 3.60 ppm in 90),

H-2 (3.97 ppm in 84 vs. 3.46 ppm in 90), and H-3 (~3.9 ppm in 84 vs. 3.59 ppm in 90)].

As the splitting pattern and coupling constants of the analogous protons in each compound remained relatively the same, the above observations, therefore, further supported configurational changes on the A ring distinguishing compound 90 from its isomer makilactone G.

Figure 69. Overlay of 1H NMR spectra of AA06795LG.D3.Di2.H8 (90) and

AA06795LG.D3.Di2.H10 (84)

Highlighted are the shifts in resonance of key signals suggesting differences in the A ring.

Recorded in CD3OD at 700 MHz and at 300 K.

H -20 3

H3-16 H-5 H-11 AA06795LG.D3.Di2.H10

H3-16 H-11 H3-20 H-5

AA06795LG.D3.Di2. H8

Continued

164

Figure 69 Continued.

H-3, H-15 H-1 H-2

AA06795LG.D3.Di2. H10

H-1, H-3

H-15 H-2

AA06795LG.D3.Di2.H8

Additional 1D and 2D NMR data were measured in C5D5N to enable comparison with reported data, and thus perform dereplication. Interpretation of the collected NMR spectroscopic data of 90 showed an agreement with the reported data (see Table 11), and thus this compound 90 was identified as makilactone F (Sato et al., 2009a).

165

Figure 70. HRESIMS spectrum of makilactone F (90)

PodoLGD3-Di2-H8-rerun2-2018-3-21_180321163227 #2-23 RT: 0.12-2.03 AV: 22 NL: 1.81E5 T: FTMS + p ESI sid=30.00 Full ms [150.00-800.00] 405.11585 R=131284 100

85 413.21204 80 R=127543 75

50 463.10353 45 274.27438 R=123030 RelativeAbundance 40 R=160644 35 391.36617 301.14139 R=136064 549.15810 30 R=150884 R=112990

25 177.08873 R=198950 20 369.18599 437.14208 557.20094 309.20401 R=136640 207.09928 R=124968 R=65952 15 R=179637 R=153832 256.82114 337.10502 10 R=167731 R=144135 475.46003 R=120346 581.18450 5 517.37136 639.33018 683.35639 727.38256 782.44296 R=113305 R=106704 R=101986 R=97267 R=92443 R=90333 0 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

1 Figure 71. H NMR spectrum of makilactone F (90) (700 MHz in C5D5N)

13 Figure 72. C NMR spectrum of makilactone F (90) (175 MHz in C5D5N)

166

Figure 73. HSQC spectrum of makilactone F (90) (in C5D5N)

2.2.3. 1D-NMR-dereplication-guided purification of the liquid-partitioned fractions of the EtOAc extract In a continued attempt to detect compound 83, the purification of an additional portion of the EtOAc extract was conducted. Since the glucoside of interest has been identified in the more polar sub-factions from an earlier separation of this extract, a shortcut of these fractionations was performed through liquid-liquid partition of the EtOAc extract with chloroform and water. The presence of 83 was not apparent in the 1H NMR spectra of the obtained, and therefore, further fractionation was performed initially.

167

Figure 74. Comparison of the HPLC traces of the CHCl3 (a) and aqueous (b) partitions of the EtOAc extract with that of the original glucoside-containing fraction (c)

Method: H2O/CH3CN (80:20 to 50:50 in 20 min, 50:50 to 0:100 in 1 min, 0:100 for 5 min,

0:100 to 80:20 in 2 min, 80:20 for 4 min)

(a) AA06795LG.D3.C.14 tR = 16

min

AA06795D3.F2’.1 (b)

The HPLC traces of both the aqueous partition (AA06795LG.D3.W) and the most polar sub-fraction from the silica gel purification of the CHCl3 partition

(AA06795LG.D3.C14) suggested the presence of the target compound based on comparison with its previous HPLC profile (tR = 16 min) (Figure 74). The CHCl3 sub- fraction was purified by HPLC, and four B-type podolactones, 78, 79, 91, and 92 were isolated, with the latter eluted at tR = 16 min.

168

2.2.3.1. Identification of podolactone C (91) Compound 91 was obtained as a white amorphous solid. Its HRESIMS gave a

sodiated molecular ion peak at m/z 447.1084 [M+Na]+ corresponding to a molecular

1 formula of C20H24O8S (Figure 75). The H NMR spectrum (Figure 76) displayed the

typical type-B podolactone proton signals attributed to the olefinic methine at H-11 (δH

6.20, s), as well as methine protons at H-5, H-6, and H-7, resonating at δH 1.79 (d, J = 4.6

Hz), 5.02 (dd, J = 4.6, 1.2 Hz), and 5.23 (d, J = 1.2 Hz), respectively. In addition, three

methyl singlet resonances were observed.

Figure 75. HRESIMS spectrum of podolactone C (91)

PodoLGD3C14M1H6-podolactoneC-2018-3-14_180127161319 #1-46 RT: 0.00-1.24 AV: 46 NL: 2.08E7 T: FTMS + p ESI Full ms [150.00-800.00] 447.10849 R=126069 100

90 85 80

75 70 65

55 50 45

RelativeAbundance 40 35 30

25 498.21572 R=119548 20

15 425.12679 10 R=128597 396.65146 515.09595 5 177.79880 207.09921 274.27437 325.06862 351.21447 R=135157 470.50449 561.39726 597.23060 633.45473 677.48091 727.38217 775.33336 R=114565 R=108379 R=134222 R=184485 R=164015 R=148079 R=143227 R=123193 R=112898 R=104128 R=101587 R=99679 R=93141 0 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

While two of these were characteristic to the diterpenoid isolated previously,

notably, H3-18 (δH 1.40, s) and H3-20 (δH 1.37, s), the more downfield signals at δH 1.80 169

and 2.65 were assigned to a tertiary alcohol methyl substituent (H3-16) and a sulfoxy- methyl group. The presence of a singlet methine at H-14 (δH 4.84) confirmed the quaternary nature of C-15.

Figure 76. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of podolactone (91)

Measured in C5D5N at 300 K.

(a)

(b)

Inspection of the 13C NMR and HSQC spectra suggested the presence of a pair of epoxide carbons linking C-2 (δC 51.7) and C-3 (δC 52.6) on the A ring, in addition to the

170

characteristic 7,8-epoxide ring. Moreover, two coupled methylene proton resonances occurring significantly downfield at δH 3.75 and 3.4 and having a large coupling constant

(J = 13.9 Hz) were assigned to C-7, adjacent to the sulfoxide group. Further analysis of the

HMBC spectrum confirmed the planar structure of 91. This planar structure also matched those of podolactone C and makilactone R (Cassady et al., 1984; Sato et al., 2009a), differing only at the configuration of the sulfoxide group. Comparison of the 1H and 13C

NMR data of 91 with the corresponding data for these known compounds led to the conclusion that 91 was indeed podolactone C (Cassady et al., 1984) (Table 13).

Figure 77. HSQC (a) and HMBC (b) spectra of podolactone (91)

Measured in C5D5N at 300 K

(a)

Continued

171

Figure 77 continued.

(b)

2.2.3.2. Identification of 1-epi-makilactone E (92) Compound 92 was isolated as a white amorphous solid with a molecular formula of C18H21ClO8 based on its HRESIMS spectrum displaying a sodiated molecular ion peak at m/z 423.0823 [M+Na]+ (Figure 78). This value was identical to that of makilactone E

(77), and following 1D and 2D-NMR spectroscopy-based structural assignment, 92 was assigned the same planar structure as 77. Comparison between the 1H and 13C NMR data of 77 and 92 revealed close similarities as demonstrated in Tables 11 and 12. Inspection of the carbon signals (obtained from HSQC and HMBC projections, Figure 80) of 92 indicated significant differences of the chemical shift values of the two compounds at C-2

(δC 73.1 in 77 vs. 76.0 in 92) and C-3 (δC 71.4 in 77 vs. 76.0 in 92) and suggested possible configurational differences between the two molecules in the A ring. Additionally, on

172

examining the 1H NMR spectrum (Figure 79), distinct variations in the resonance signals

corresponding to H-1 and H-2, as well as H3-18 were observed, where the former two

appeared significantly more downfield and the latter shifted upfield compared to the

respective proton signals in 77 (Table 11). Correlations between H-1 and H-2, H-2 and H-

3, and H-3 and H3-18 on the corresponding 1D-NOESY spectra (Figure 79) confirmed that

all methine protons on the A ring are α-oriented. Therefore, 92 was identified as the 1-

epimer of 77, and given the name 1-epi-makilactone E.

Figure 78. HRESIMS of 1-epi-makilactone E (92)

PodoLGD3C14M1H10-podolactoneC-2018-3-15-run2_180127161319 #1-54 RT: 0.02-1.47 AV: 54 NL: 1.20E7 T: FTMS + p ESI Full ms [150.00-800.00] 423.08231 R=129488 100

95 90 85

75 70 65 325.16011 R=144279 60

55 50 45

RelativeAbundance 40 413.21245 R=129711 35 30 25

15 309.20429 385.16279 581.23627 R=109142 R=149396 R=135720 443.22306 533.20000 10 195.09944 274.27466 R=125426 R=116158 496.04632 R=190439 R=158556 353.23047 R=117467 683.35693 727.38291 5 609.23144 639.33055 787.19860 239.12567 R=144475 556.95340 R=101663 R=99290 R=174676 R=112108 R=105435 R=102090 R=92928 0 200 250 300 350 400 450 500 550 600 650 700 750 800 m/z

173

Figure 79. 1H NMR spectrum (a) of and key NOESY correlations (b) for 1-epi- makilactone E (92)

(a) Measured in C5D5N at 400 MHz

(b) Data obtained from 1D-NOESY experiments in CD3OD

174

Figure 80. HSQC (a) and HMBC (b) spectra of 1-epi-makilactone E (92)

Measured in C5D5N at 300 K

(a)

(b)

175

2.2.3.3. 1D-TOCSY NMR spectroscopy-guided screening of the EtOAc extract and fractions

The peak eluted at tR = 16 min from the HPLC purification of AA06795LG.D3.C14

(section 2.2.3) did not correspond to the targeted compound, and instead was identified as

92. Moreover, the NMR resonance signals predicted to belong to those of the new podolactone glucoside 83 were later shown to be from its corresponding aglycone (79), which shares the same podolactone 1H NMR spectroscopic pattern. According to these results, one can postulate that either 83 was absent from this fraction or it hydrolyzed in its aglycone. To further investigate these hypotheses and to verify the presence of 83 in the large-scale EtOAc extract, a 1D-TOCSY NMR spectroscopy experiment was designed to target the 1H NMR spectroscopic differences between 83 and 79. In so doing, not only would the ambiguity with 79 would be avoided, but the presence of 83 would be confirmed through its specific resonance signals. Accordingly, the crude EtOAC extract and both its

CHCl3 and aqueous partitions were screened by 1D-TOCSY NMR spectroscopy, where the spectroscopic region corresponding to proton H-2 of 83 was irradiated. By irradiating

H-2 or any of the A ring protons (e.g., H-1a, Figure 81), the four remaining protons belonging to the same spin network would appear in the resulting 1D-TOCSY NMR spectrum (see Experimental Section for details on the 1D-TOCSY experiment). Therefore, the presence of 83 in the extract and fractions could be confirmed based on comparison between their respective 1D-TOCSY spectra and the 1H NMR spectrum of 83.

176

Figure 81. Overlay of 1H (a) and 1D-TOCSY NMR (b) spectra of 83

Measured in C5D5N. Irradiated H-1a ( δH 2.36, dd, J = 13.8, 8.7 Hz)

(a)

H-2 H-3a,b H-1a H-1b (b)

Of the four 1D-TOCSY NMR spectra obtained from the screening of the crude

EtOAc extract, two (spectra 2 and 3, Figure 83) exhibited resonance signals corresponding to the A ring proton spin network of 83. These spectra resulted from the selective irradiation of the 4.05-4.10 and 4.10-4.15 ppm range of the 1H NMR spectrum of this extract (Figure

82), and thus the target H-2 proton of 83 within this extract resonated within this region.

However, since spectrum 3 showed higher purity, as demonstrated by the lack of additional peaks in the 1D-TOCSY spectrum (Figure 83), it is more likely that an equally lower degree of overlap of similar spin network within that specific region. Resonances attributed to the A ring spin network of the aglycone 79, were also observed resulting from irradiation of the 4.15-4.20 ppm range (spectrum 4, Figure 83), confirming the presence of this compound in the extract and the similarity between its 1H NMR data and that of 83.

177

Figure 82. 1D-TOCSY NMR experiment for P. neriifolius EtOAc extract in CD3OD

1 (a) H NMR spectrum of 83 in CD3OD highlighting ring A protons. (b) Irradiated region for 1D-TOCSY experiment of the EtOAc extract

(a) H-2 H-1a H-3a H-3b H-1b

1 2 3 4

178

Figure 83. 1D-TOCSY NMR spectra for P. neriifolius EtOAc extract

(a) Full 1D-TOCSY spectra (1-4) overlaid with the original 1H NMR spectra of EtOAc extract and that of 83. (b) Detection of 83 from spectra 2 and 3. (c) Detection of 79 from spectrum 4

(a) 1D-TOCSY (4)

1D-TOCSY (3)

1D-TOCSY (2) 1D-TOCSY (1) 1 H NMR spectrum (83)

1H NMR spectrum EtOAc extract

(b) H-1a H-3a H-1b

1D-TOCSY (3)

1D-TOCSY (2)

1 H NMR spectrum (83) 1Hnagilactone NMR spectrum G-2-O -glc EtOAc extract

H-1a H-3a H-3b H-1b (c) 1 H NMR spectrum (79) 1D-TOCSY (4)

1D-TOCSY (3) 1D-TOCSY (2)

1 H NMR spectrum (83) 1H NMR spectrum EtOAc extract 179

The 1D-TOCSY NMR spectroscopic data obtained from the same experiment conducted with the aqueous partition (AA06795LG.D2.W) showed the same pattern as that of its extract of origin (EtOAc), confirming the presence of the targeted podolactone glucoside (83) in this fraction. However, no resonance corresponding to that of the aglycone (79) was apparent in any of the resulted spectra (Figure 84).

Figure 84.1D-TOCSY NMR spectra for the aqueous fraction (AA06795LG.D3.W)

(a) Full 1D-TOCSY spectra (1-5) overlaid with the original 1H NMR spectra of

AA06795LG.D3.W and that of 83. (b) Detection of 83 from spectrum 2.

(a) 1D-TOCSY (5)

1D-TOCSY (4)

1D-TOCSY (3)

1D-TOCSY (2)

1D-TOCSY (1)

1 H NMR spectrum (83) 1H NMR spectrum AA06795LG.D3.W

(b) H-1a H-3a H-1b

1D-TOCSY (2)

1 H NMR spectrum (83)

1 H NMR spectrum AA06795LG.D3.W

180

On the other hand, the CHCl3 partition did not seem to contain 83, based on its 1D-

TOCSY NMR spectra, and instead one of these spectra matched the corresponding signals for the A ring protons of 79 (Figure 85). These observations confirmed the earlier results from the HPLC profiling and purification of the polar sub-fraction of the CHCl3 partition.

Figure 85. 1D-TOCSY NMR spectra for the CHCl3 fraction (AA06795LG.D3.C)

(a) Full 1D-TOCSY spectra (1-4) overlaid with the original 1H NMR spectra of

AA06795LG.D3.C and that of 83 and 79. (b) Detection of 79 from spectrum 4.

(a) 1 H NMR spectrum (79)

1D-TOCSY (4)

1D-TOCSY (3)

1D-TOCSY (2) 1D-TOCSY (1)

1 H NMR spectrum (83) 1 H NMR spectrum AA06795LG.D3.C

(b) H-1a H-3a H-3b H-1b

1 H NMR spectrum (79)

1D-TOCSY (4)

1H NMR spectrum AA06795LG.D3.C

181

Based on the above observations, the aqueous partition was purified for the isolation of compound 83. Since two peaks were observed around the retention time at which the target compound eluted using the same method, both peaks were collected. As predicted, one of these peaks (tR = 16.3 min, peak 2), was identified as the target compound based on 1H NMR fingerprinting (Figure 86), thus validating the screening method performed through 1D-TOCSY-NMR spectroscopy. From this structural confirmation, a larger quantity of this peak was collected to complete the purposed full characterization and structural elucidation of (83), which was not previously possible due to the limited amount obtained. In addition, the second peak (tR = 16.0 min, peak 1) collected from the water-soluble partition of the EtOAc extract was identified as 1-epi-makilactone E (92), as its 1H NMR spectrum perfectly superimposed with that of AA06795LG.D3.C14.M1.H10.

Figure 86. Overlay of the 1H NMR spectra of (a) 83 and (b) that of peak 2 from

AA06795LG.D3.W

(a)

(b)

182

According to this body of evidence, the 1D-TOCSY NMR spectroscopy-based dereplication of the EtOAc represented a useful and user-friendly tool for the rapid detection of the target compound. Moreover, this experiment served as a confirmation of prior results obtained by HPLC, underlining the inefficiency of HPLC-UV alone during this dereplication study, as shown by the absence of compound 83 in the CHCl3 partition.

To further illustrate the importance of this NMR-based dereplication method for a more efficient fractionation, a portion of the original EtOAc extract was fractionated into three aliquots on a reversed-phase C18 column, and all three were subjected to both HPLC and

1D-TOCSY NMR spectroscopy.

Figure 87. Overlay of 1H NMR (a) and 1D-TOCSY (b) NMR spectra of AA06795LG-

D3.F1 with that of 83 (c) (zoomed 1.30-3.00 ppm region)

H-1a H-3a H-1b (c)

(b)

(a)

Hence, only the most polar fraction (AA06795LG.D3.F1) indicated the presence of

83 based on its HPLC profile, and this was confirmed by its second 1D-TOCSY spectrum

183

(irradiating δH 4.10-4.15 ppm) (Figure 87). Thus, an alternative effective and safer fractionation method could be performed toward further isolation of 83 through C18 fractionation in MeOH/H2O.

2.2.4. Screening of the hexane extract by 1H NMR dereplication The 1H NMR spectrum of the crude hexane extract (AA06795LG.D1) measured in

CDCl3 (Figure 88) displayed highly overlapping peaks resonating in the aliphatic region

(δH = 1.0-2.5 ppm). However, of interest was the olefinic-aromatic region of the spectrum with the presence of signals characteristic of the coupled aromatic protons of previously obtained totarane-type diterpenes [δH 7.01 (d, J = 8.6 Hz) and 6.55 (d, J = 8.6 Hz)], a singlet proton peak at δH 6.9 analogous to the H-11 signal of previously isolated podolactones, and three coupled olefinic protons [δH 5.8 (dd, J = 17.5, 10.7 Hz), 4.93 (dd, J = 17.5, 1.5 Hz) and 4.90 (d, J = 10.7, 1.5 Hz)] suggesting the presence of a vinyl group. In addition, a resonance signal at δH 9.85 was consistent with the occurrence of a formyl group- containing molecule. Constituents identified from P. neriifolius previously in this study have not possessed a comparable olefinic group or an aldehyde as a substituent, and this observation, by means of dereplication, prompted further purification of this extract towards the characterization of the corresponding compound(s), along with the identification of the suggested totarane and podolactone derivatives containing protons equivalent to the above-mentioned resonance signals.

The 1H NMR-guided targeted fractionation of the hexane extract led to the

184

identification of four diterpenoids, including three totarane (85, 86, and 88) and one pimarane-type (87) compounds. As illustrated in Figure 89, the pairs of aromatic proton doublets corresponded to the predicted H-11 and H-12 of 4β-carboxy-19-nor-totarol (82), totarol (85) and totaral (86), and the latter was also responsible for the formyl group proton signal observed in the mother extract. Moreover, the vinyl group proton signals were identified as corresponding to the side chain of sandaracopimaric acid (87). However, it was shown that the singlet methine proton at δH 6.90 corresponded to H-11 and H-11′ of macrophyllic acid (88), a dimer, and not that of a podolactone as predicted.

The assignment of compounds 85-87 was based solely on their respective NMR spectroscopic data, since they are previously known compounds and lacked cytotoxic activity when evaluated in the panel of four cancer cell lines used in this study.

185

Figure 88. 1H NMR spectroscopic profile of the hexane extract (AA06795LG.D1)

(a) Pair of aromatic ortho-protons. (b) H-1 singlet proton (c) Vinyl protons

(a) (b) (a)

7.2 7.0 6.8 6.6 6.4 6.2 6.0 5.8 5.6 5.4 5.2 5.0 ppm

7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 ppm

Figure 89. Overlay of 1H NMR spectrum of the hexane extract (AA06795LG.D1) with that of compounds 82, 87, and 88.

(87)

(88)

(82)

hexane extract

7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 ppm Scale: 0.1938 ppm/cm, 77.55 Hz/cm

186

2.2.4.1. Identification of macrophyllic acid (88) from a sub-fraction of the hexane extract Compound 88, a white solid, was obtained as a precipitate from a fraction

(AA06795LG.D1.6.10) of the hexane extract purified on silica gel. It was assigned its

+ molecular formula (C40H54O6) on the basis of the [M + Na] ion peak at m/z 653.1814 in

the HRESIMS (Figure 90), which suggested that 88 is a dimeric compound. Its 13C NMR

spectrum (Figure 91) displayed twenty peak signals (nineteen annotated and δC 135.1 also

present but at very low intensity) suggesting a diterpenoid and corresponding to the

monomeric unit.

Figure 90. HRESIMS spectrum of macrophyllic acid (88)

AA06795LG.D1 #26-198 RT: 0.21-1.60 AV: 173 NL: 4.53E7 T: FTMS + p ESI Full ms [100.00-1400.00] 653.38141 R=25103 100

90 85 80

65 60 55

45 RelativeAbundance 40 35

30 469.32853 1283.77466 R=17932 25 R=30069

20 15 675.36284 10 R=25203 1306.75933 413.21158 R=18004 5 R=32442

0 200 300 400 500 600 700 800 900 1000 1100 1200 1300 1400 m/z

When inspecting the 1H NMR spectrum (Figure 91), resonances characteristic to

the pair of isopropyl methyl at C-15 were observed [δH 1.34, d, J = 6.9 Hz (H3-16) and δH 187

1.38, d, J = 6.9 Hz (H3-17)], together with a singlet methine at δH 6.9, suggesting that the monomers were linked at C-12. This was confirmed by analysis of both HSQC and HMBC spectra (Figure 92). Hence 88 was identified as macrophylllic acid (Bocks et al., 1963;

Amaro & Dignora Carros, 1989).

Figure 91. 1H (a) (400 MHz) and 13C (b) (100 MHz) NMR spectra of macrophyllic acid

(88) (Measured in CDCl3 at 300 K)

(a)

(b)

188

Figure 92. HSQC (a) and HMBC (b) spectra of macrophyllic acid (88)

Measured in CDCl3 at 400 MHz and 300 K

(a)

(b)

2.2.4.2. Identification of compounds 85-87 from sub-fractions of the hexane extract Compounds 85-87 were identified as totarol, totaral, and sandaracopimaric acid,

189

respectively, from the semi-purified sub-fractions of the hexane extract through interpretation of their NMR spectroscopic data and comparison with the published literature (Figures 93-98) (Ying & Kubo, 1991; Muto et al., 2008). Due to ionization difficulties, the HRESIMS data in the positive mode of these fractions did not indicate the presence of their respective predicted sodiated molecular ions [85, m/z 309.2189 [M+Na]+

+ + + (calcd. C20H30ONa ); 86, m/z 323.1982 [M+Na] (calcd. C20H28O2Na ); 87, m/z 325.2138

+ + [M+Na] (calcd. C20H30O2Na )], but the corresponding HRESIMS data were obtained in the negative mode, confirming the structures of these compounds (Appendix H).

Figure 93. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra of totarol (85)- containing sub-fraction

(a)

(b)

190

Figure 94. HSQC (a) and HMBC (b) NMR spectra of totarol (85)-containing sub-fraction

(a)

(b)

191

Figure 95. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra totaral (86)-containing sub-fraction

(a)

(b)

Figure 96. HSQC (a) and HMBC (b) NMR spectra of totaral (86)-containing sub-fraction

(a)

Continued

192

Figure 96 continued.

(b)

Figure 97. 1H (400 MHz) (a) and 13C (100 MHz) (b) NMR spectra sandaracopimaric acid

(87)-containing sub-fraction

(a)

(b)

193

Figure 98. HSQC and HMBC NMR spectra of sandaracopimaric acid (87)-containing sub- fraction

(a)

(b)

3. Monitoring podolactone fungal biotransformation with 1D-NMR spectroscopy Several biotransformation procedures were conducted using the crude aqueous extract of P. neriifolius root sample in both P. concentricum and P. expansum fermentation

194

cultures, and in other cases, inumakilactone A-15-O-β-D-glucoside (89)

(AA06795LGD2.2) and its aglycone inumakilactone A (78) (AA06795LG.D3P).

Described in the following sections are the results from the bioconversion experiments, first using P. concentricum, then in P. expansum cultures. As the results illustrate, this study highlights some fermentation requirements for the occurrence and detection of bioconversion, and the effectiveness of 1H NMR spectroscopy as a method to monitor these. Moreover, some insights on the structural stability of the A-ring epoxy-group of the podolactones in this study are inferred. Finally, the biological data of the extractives obtained from the difference fungal fermentations conducted further confirmed the observations made from the 1H NMR spectroscopic analysis.

3.1. Biotransformation in Penicillium concentricum culture Initial biotransformation studies on podolactones from P. neriifolius in the present research were conducted using Penicillium concentricum as the fungal strain. Three podolactone sources constituted the starting material, namely, the aqueous extract

(AA06795LG.D2), its purified fraction (AA06795LG.D2.2), and the pure aglycone inumakilactone A (AA06795LG.D3P, 78). The observations derived from each of these procedures are described in the following paragraphs.

3.1.1. Monitoring P. concentricum-assisted biotransformation of AA06795LG.D2.2 through 1H NMR spectroscopy Following the detection, by 1H NMR spectroscopy, of inumakilactone A-15-O-β- 195

D-glucoside (89) as the main and nearly pure constituent of AA06795LG.D2.2 fraction, the latter was first subjected to fungal bioconversion in a P. concentricum fermentation culture as part of a pilot study. The reaction was analyzed at two endpoints, days 4 and 6, respectively. At the first endpoint, while the fungal culture was still ongoing, a small portion of the fermented mixture was aliquoted and extracted, partitioned with solvents, and analyzed by 1H NMR spectroscopy. The 1H NMR spectrum of the obtained aqueous partition (PN-D2.2-PC-4d-W) suggested the presence of the starting podolactone glucoside as evidenced by key resonance signals, such as H-11 (δH 6.75), the anomeric proton (δH

4.99), and the characteristic methyl signals at δH 1.57, 1.51, and 1.32, corresponding to methyl protons at positions C-17, C-20, and C-18, respectively. In addition, another set of key podolactone signals, resonating in close proximity to analogous peaks of the starting material appeared in the 4-day aqueous partition (PN-D2.2-PC-4d-W) of this pilot fungal biotransformation mixture. Overlapping this spectrum with that of the starting material 89 and its aglycone (78) confirmed the presence of both compounds in the aqueous partition

(Figure 99). However, the EtOAc partition obtained from the first endpoint of this pilot fungal biotransformation, when subjected to 1H NMR spectroscopic analysis, did not appear to contain any podolactones.

196

Figure 99. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-4d-W, (b) 89 and, (c) 78

Measured in CD3OD, at 400 MHz.

(c)

(b)

(a)

While the formation of the aglycone analog (78) was anticipated through P. concentricum-assisted hydrolysis of the glucopyranosyl moiety, the presence of this non- readily water-soluble metabolite in the aqueous partition rather than its less polar EtOAc counterpart was at first rather intriguing. A plausible explanation to this unexpected distribution can, however, be provided by a closer evaluation of the extraction and partition processes conducted on the 4-day fungal fermentation aliquot. The addition of EtOAc and subsequent extraction and partition of the 4-day mixture were performed prior to removal of the fungal cells through filtration. Therefore, it is possible that the remaining aqueous

197

layer still containing most of the fermentation products also retained the aglycone (78).

Nevertheless, fermentation was maintained to further analyze additional reaction(s) or the completion of the podolactone glucoside hydrolysis toward the formation of the aglycone.

At the second endpoint (day 6) and termination of the pilot biotransformation procedure, the fermentation mixture was extracted with EtOAc and filtered simultaneously prior to further partition of the obtained aqueous filtrate. In this manner, a more exhaustive extraction of EtOAc-soluble compounds was achieved. Accordingly, the 1H NMR spectroscopic profiling of the 6-day aqueous partition (PN-D2.2-PC-6d-W) did not result in an apparent detection of either the podolactone glucoside 89 or its aglycone 78 (Figure

100). On the other hand, the 1H NMR spectrum of the 6-day EtOAc partition shared some similarities with the above-described 4-day aqueous mixture, as both 78 and 89 were detected in this ethyl-acetate soluble fraction (PN-D2.2-PC-6d-E) (Figure 101).

Figure 100. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-6d-W, (b) 89 and, (c) 78

Measured in CD3OD

(c)

(b)

(a)

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Figure 101. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-6d-E, (b) 89 and, (c) 78

Measured in CD3OD

H -20, H -16, and H -18 H-11 3 3 3

(c)

(b)

(a)

Comparison of the 1H NMR profiles of both the 4-day aqueous and the 6-day

EtOAc partitions revealed some differences, and these were mainly attributed to signals associated with the fungal metabolite(s). First, as mentioned above, the longer fermentation time, as predicted, caused the production of greater amounts of fungal metabolites.

However, comparison between an aliquot and a total extraction at two different endpoints may not be the most reliable method for quantitative analysis of the metabolite content, as the latter would be more concentrated by nature. Nevertheless, a more unbiased

199

comparison of resonance signal intensities would require homogeneity, in terms of concentrations, of the samples used for the NMR spectroscopy experiment. For example, an aliquot from a dilute EtOAc partition from the second endpoint would contain less metabolite than a sample from a concentrated extract from the first endpoint. A more desirable condition for such analytical comparison would, therefore, necessitate the usage of known concentrations of the fermentation mixtures (partitions) for NMR spectroscopic analysis.

This pilot fungal biotransformation of 89 using AA06795LG.D2.2 in a P. concentricum fermentation culture resulted in the successful hydrolysis of this glucoside into its corresponding aglycone, inumakilactone A (78), confirming the earlier hypothesis regarding the use of this method towards this result, and fulfilling one of the goals of this procedure. Thus, the selective hydrolysis of the glucoside derivative left intact the epoxy groups on both the A and B rings, and hence conserving the bioactivity of the resulting aglycone. This biotransformation, though not complete, occurred by the second day as displayed by the 1H NMR spectroscopic analysis, and this relatively fast reaction can be explained by the higher concentration of fungal cells in the culture and the amount of starting material used (nearly 600 mg). This 1D-NMR spectroscopic method constituted a tool for the fingerprinting the fungal fermentation extracts, as it provided structural information on the composition of the analyzed extracts, as well as a means to compare these. Moreover, in this study, 1D-NMR spectroscopy, through the usage of previously obtained in-house spectroscopic data, represented a rapid and reliable method of

200

dereplication that resulted in the identification of a biotransformation product without undergoing extensive chromatographic purification.

However, comprehensive quantification, purification and identification of metabolites from the obtained partitions were not conducted as such analytical procedures would fall somewhat outside the scope of the current study. Additional biotransformation experiments were conducted for further reproducibility and investigation of possible variations with respect to the time and starting material required, and the metabolite(s) produced.

3.1.2. Monitoring P. concentricum-assisted biotransformation of AA06795LG.D2 through 1H NMR spectroscopy Following the successful pilot biotransformation study described in the preceding section, conducted with the aqueous sub-fraction AA06795LG.D2.2, the original aqueous extract (AA06795LG.D2) was used, in two subsequent batches, for additional biotransformation experiments. Considering the highly similar 1H NMR spectroscopic profiles of both these extracts, notably in term of their content in the podolactone glucoside of interest (89), the more crude AA06795LG.D2 was used as an alternative to its further purified fraction. Moreover, using this original extract allowed the study to be expedited as it did not require additional fractionation, thus constituting a means for dereplication.

In one of the two sets of P. concentricum-assisted biotransformation experiments using AA06795LG.D2 as the starting material, two ~250-mg samples of this extract were used, in separate fermentation flasks with the same conditions as with the pilot 201

biotransformation study. At the 2-day endpoint, aliquots from both flasks were obtained, filtered, partitioned as with previous study with EtOAc, and analyzed by 1H NMR spectroscopy. The resulting spectra of the water-soluble partitions (PN-D2-PC-I-2d-W and

PN-D2-PC-II-2d-W) showed the presence of fungal metabolites and that of inumakilactone

15-O-β-D-glucoside (89) at lower intensity (Figure 102). On the other hand, 1H NMR spectra obtained from the 2-day EtOAc partitions from both flask I and II did not show any distinctive resonances associated with the occurrence of podolactones (PN-D2-PC-I-2d-E and PN-D2-PC-I-2d-E).

Figure 102. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-2d-W, (b) PN-D2.2-PC-

II-2d-W, (c) 89 and, (d) 78. Measured in CD3OD, 400 MHz

(d) (c)

(b)

(a)

According to these observations, it was inferred that biotransformation in this study

202

required a longer fermentation period. However, despite the absence of biotranformation, the nearly identical spectra obtained from each set of partition (aqueous and EtOAc) from each flask at this endpoint revealed that the experiment was reproducible across both flasks.

Analysis of partitions from the 3-day endpoint aliquots also led to the above conclusion.

As with the 2-day endpoint, each set of partitions from both flasks displayed identical NMR spectra. While the 1H NMR spectra of the 3-day aqueous partitions (PN-D2-PC-I-3d-W and PN-D2-PC-II-3d-W) from both flasks were similar to those acquired for the analogous

2-day partitions (Figure 103), the 1H NMR spectra of the EtOAc partitions obtained at the

3-day endpoint (PN-D2-PC-II-3d-E and PN-D2-PC-II-3d-E) showed the presence of fungal metabolites unlike the spectra of the 2-day EtOAc partitions (Figure 104). These results, therefore, suggest the start of fungal metabolite production to occur within the 2 to

3-day period.

Figure 103. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-2d-W, (b) PN-D2.2-PC-

I-3d-W Measured in CD3OD, 400 MHz

(b)

(a)

203

Figure 104. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-2d-E, (b) PN-D2.2-PC-

I-3d-E

Measured in CD3OD, 400 MHz

(b)

(a)

Figure 105. Overlay of the 1H NMR spectra of (a) PN-D2.2-PC-I-3d-E, (b) PN-D2.2-PC-

I-6d-E, and (c) PN-D2.2-PC-II-6d-E

(c)

(b)

(a)

Spectroscopic data obtained from the 1H NMR profiling of the partitions of the 6- day fermentation aliquots were similar to those obtained from the 3-day partitions, with the

204

EtOAc partition (PN-D2.2-PC-I-6d-E and PN-D2.2-PC-II-6d-E) containing higher concentration of the fungal metabolites as deduced from the intensities of the resonances signals (Figure 105). Analysis by 1H NMR spectroscopy of the extractives from the aliquot fermentation of each culture at days 2, 3, and 6 and from the total culture at day 8 did not reveal the presence of the desired aglycone product (78), but only the starting podolactone 89. Therefore, while the results from each culture flask in this set of biotransformation procedures showed the reproducibility of the study and that of the 1D-

NMR-spectroscopy-based dereplication, it seemed that a higher quantity of the starting material was required, either for the reaction to occur or for the detection of possibly trace amounts of the biotransformed podolactone, had it taken place.

The second set of fungus-assisted biotransformation of the Podocarpus neriifolius aqueous extract (AA06795LG.D2) was conducted with a 10 g-sample of this extract in one fermentation flask of P. concentricum culture in 250 mL potato dextrose broth. Two aliquots of the fermentation were collected at 2 and 6 days, respectively, and at day 7 the cultivation was terminated. As with the previous 1H NMR spectroscopy-based biotransformation monitoring processes, the spectroscopic profiles of both the water- and ethyl acetate-soluble partitions from each endpoint were collected. As observed in the resulting NMR spectra, the usage of a high amount of starting material in this batch resulted in a rapid and detectable hydrolysis of 83 to its aglycone, inumakilactone A (78).

According to the 1H NMR dereplication of the 2-day EtOAc partition of the fermented mixture, both podolactone derivatives were present in this mixture, along with

205

some fungal metabolites. The observed overlapped singlets at δH ~ 6.46 ppm corresponding to the resonance signals of the proton at H-11 of each of the podolactones of interest, along with the overlapped doublets in the δH 2.12 ppm region (H-5), and the presence of two overlapped sets of the characteristic methyl groups at C-20, C-16, and C-18 resonating within the δH 1.43-1.45 ppm, 1.34-1.40 ppm, and 1.20-1.22 ppm regions, respectively.

Figure 106. Overlay of the 1H NMR spectra of (a) PN-D2-PC-2d-E, (b) 89, and (c) 78

H-5 H3-20, H3-16, and H3-18 H-11

(c)

(b)

(a)

Overlaying the 1H NMR spectra of the inumakilactone A (78) and its glucoside derivative (89) with that of the 2-day EtOAc partition (PN-D2-PC-2d-E) confirmed this

206

observation (Figure 106). However, as seen in this figure, while the peak patterns correspond to both the podolactones as observed in the pilot study (section 2.1.1), they did not always show an exact overlap. This shift can be explained, first, by the mixed nature of the partition analyzed, as the presence of various metabolites at high concentrations within the same sample can cause stronger intermolecular interactions leading to a slight shift of their corresponding resonance signals. Moreover, in some cases, the presence of solvent (EtOAc) in the partitions can also cause peak shifting. The use of 1D-TOCSY spectroscopy in the equivalent EtOAc partition obtained at the 6-day endpoint undoubtedly confirms the presence of both podolactones, as described in more details in the subsequent paragraph.

Figure 107. Overlay of the 1H NMR spectra of (a) PN-D2-PC-2d-W and (b) PN-D2-PC-

6d-W

Measured in D2O at 400 MHz

(b)

(a)

207

1 The water-soluble extract, of which the H NMR data was measured in D2O due to its poor solubility at high concentration in deuterated methanol seemed to only contain the starting material (89) (Figure 107).

The EtOAc partition obtained at the 6-day endpoint (PN-D2-PC-6d-E), similarly to its 2-day analogous partition, also contained both compounds 78 and 89. As mentioned previously, in some cases, the resonance signals corresponding to the characteristic peaks used to identify these compounds in the mixtures tended to shift, thus making the 1H NMR spectroscopy-based dereplication method somewhat ambiguous in these instances. To further confirm the presence and identity of the podolactones of interest, the mixture being investigated, the 6-day fermented fungal mixture EtOAc partition, was scanned using 1D-

TOCSY NMR spectroscopy, a method previously used for the detection of nagilactone G-

2β-O-β-D-glucoside (83) in various P. neriifolius extracts (section 2.6).

The 1H NMR spectra of compound 89 mainly differs from that of its aglycone at the H-15 proton chemical shift, due to the glucopyranosyl moiety at C-15, and the resulting shifting of signals corresponding to its coupled protons at positions H-14 and H3-17, respectively. Thus, irradiating the peak signals corresponding to any of the protons at these positions in a 1D-TOCSY NMR experiment would result in a spectrum showing all these protons from the same spin network (Figure 108).

208

Figure 108. Overlay of the 1H NMR spectra of (a) 89 and (b) 78

H-15 H3-17

(b)

(a)

However, the methyl protons at H3-17 would be the best choice for selective irradiation to measure a 1D-TOCSY NMR spectrum with the least ambiguity, since both peaks appeared clearly in the 6-day EtOAc partition (PN-D2-PC-6d-E) of the fungal fermentation. Moreover, both the H-14 and H-15 protons of inumakilactone A (78) resonated at distinct chemical shifts when compared to the analogous resonances in its glucoside derivative (89). These resonances were dispersed enough so that each proton would not overlap with its equivalent proton from the other derivative when both are present in the same mixture.

Therefore, the doublet signals at δH 1.36 ppm and 1.38 ppm corresponding to the methyl protons at H3-17 of each podolactone derivative were selectively excited, and the resulting 1D-TOCSY NMR spectra showed clear signals corresponding to their respective

H-14 and H-15 protons. Overlapping these 1D-TOCSY NMR spectra with the 1H NMR 209

spectrum of the original mixture (6-day EtOAc), that of compound 78 and 89 each confirmed unambiguously the identity and presence of both compounds in this EtOAc partition, as all peaks of interest exactly overlapped with their equivalent signals from the purified compounds (Figure 109).

Figure 109. Detection of 89 and 78 in PN-D2-PC-6d-E fraction by 1D-TOCSY NMR spectroscopy

1 Measured in CD3OD, 400 MHz. (a) H NMR spectrum of PN-D2-PC-6d-E. (b) 89. (c) 78.

H-14 H-15 H3-17

(c)

H-14 H-15 H3-17

(b)

(a)

The fermentation procedure was then terminated at day 7, and the partitions from the resulting extract were subjected to 1H NMR spectroscopic analysis. As with the previous endpoints (2 and 6 days), the fermented mixture still contained the starting material 89 in both its water- and ethyl acetate-soluble partitions. The intense appearance of this compound in the aqueous partition is not surprising, as a very high concentration 210

was used in this study, in comparison to previously conducted biotransformation experiments mentioned above (250 and 600 mg), representing approximately 16-40 fold the quantities used. Thus, using such a large amount of compound 89 in the same volume of fungal fermentation broth and analyzing at the same endpoints resulted in the above observation. On the other hand, while both compounds 78 and 89 were also present in the

EtOAc partition of the 7-day fermentation as seen in the corresponding 6-day partition, the intensity of the signal corresponding to the methyl proton attached to C-17 of the glucoside

(89) was much lower compared to its equivalent resonance in the aglycone congener 78.

Therefore, based on these observations, the biotranformation of 89 to 78 continued at day

7 and either a higher concentration of the latter compound was present overall in the 7-day fermentation or the 7-day EtOAc (PN-D2-PC-7d-E) sample utilized for 1H NMR spectroscopic profiling was more concentrated than that used in the 6-day partition. A similar conclusion could be inferred regarding the concentration of the fungal metabolites, for which the spectroscopic signals were much more intense in the last endpoint than the one preceding it.

The biotranformation procedures conducted with the second batch of the starting material (10 g) allowed the replication of the pilot study, as the successful production of the aglycone was observed from the 1H NMR data analysis of the EtOAc partitions at each fermentation endpoint. Furthermore, this study gave some insights on the quantity of the podolactone glucoside required for the occurrence and/or the detection of this type of biotransformation. Thus, using the same conditions for the fungal culture broth (250 mL),

250 mg of the plant aqueous extract were found to be insufficient to afford a successful

211

detection of biotransformation, if it occurred, through the 1H NMR spectroscopy-based monitoring method.

3.1.3. Monitoring P. concentricum-assisted biotransformation of inumakilactone A (78) through 1H NMR spectroscopy Following the successful hydrolysis of 89 in P. concentricum fungal culture to 78, the latter was subjected to the same biotransformation procedure to gain some insights on further fungal-assisted structural modifications of this compound. While one may argue that a similar analysis could have been carried out by extending the fermentation period in cultures where the aglycone was formed, notably in the 10 g-glucoside-biotranformation study, the use of a fresh culture constituted a better alternative as the culture was not contaminated by high concentration of fungal metabolites, facilitating the 1H NMR-based fingerprinting used. In addition, subjecting P. concentricum directly to the cancer cell cytotoxic aglycone (78) allowed for the assessment of possible fungal vulnerability toward this plant secondary metabolite independently of the lack of or reduced levels of nutrients through the observation of fungal growth and metabolite production. Since the fermentation culture was kept constant (with no potato dextrose broth added), as the fungus proliferates through the progression of the fermentation, the concurrent reduction of this nutrient would eventually produce a stressful environment, impeding fungal growth. In that case, if 78 also affected fungal growth, this causal agent would be less apparent in an already nutrient-deficient environment. Thus, performing the biotransformation of this compound in a freshly made fungal culture allows for both a facilitated detection of its effect on the P. concentricum and for a longer duration of the procedure. Accordingly, the

212

fungal-assisted biotransformation of 78 was performed using the material precipitated from a fraction from the EtOAc extract of the plant (AA06795LG.D3.1P) and monitored through

1H NMR spectroscopy at days 5, 9 and 16. Surprisingly, no biotransformation was apparent at either of the 5- and 9-day endpoints, or at the termination of the procedure at day 16, according to the respective 1H NMR spectroscopic profiles obtained from the partitioning of the fungal extracts at these endpoints and comparison of the spectra with the 1H NMR spectrum of the starting material. As illustrated in Figure 110, the only podolactone present in the EtOAc partitions of the fungal extracts obtained was the starting material, and thus, the fungus did not structurally modify this phytochemical. Moreover, from the physical observation of the fermentation, inumakilactone A (78) did not seem to negatively affect fungal growth. This is perhaps one of the causes of the absence of biotransformation in the current procedure. Furthermore, it can be speculated that unlike its glucosidic derivative

(89) from which the fungus could harness glucose as nutrient, 78 does not seem to possess a structural feature “needed” by the fungus, thus resulting in the inert fermentation.

Another explanation of the current observation can be related to the solubility parameters of this podolactone. Since the fermentation environment is constituted primarily of water, it is possible that the sparingly water-soluble starting material did not fully dissolve in the fungal culture broth, despite constant shaking, and, consequently, only a small amount, if any, would have been accessible to the fungus. As a result, although compound 78 may have been derivatized by the fungus, the yield of such reaction(s) would have been too low to allow for the detection of the product through the current 1D-NMR spectroscopy-based approach. Hence, the need, as a further direction, for the additional analysis and purification of the extracts obtained from these fungal fermentations. 213

Figure 110. Overlay of the 1H NMR spectra from the biotransformation of 78 in P. concentricum

1 Measured in CD3OD, 400 MHz. (a) H NMR spectrum of PN-D3-PC-5d-E. (b) PN-D3-

PC-9d-E. (c) PN-D3-PC-16d-E. (d) 78.

(d)

(c)

3.2. Biotransformation in Penicillium expansum culture To further evaluate the reproducibility of the above-reported fungal-assisted hydrolysis of inumakilactone A-15-O-β-D-glucoside (89) to its aglycone derivative, a second Penicillium species (strain), P. expansum, was cultivated under similar conditions as P. concentricum, and fermented in two sets of biotransformation procedures with the

214

crude aqueous extract (AA06795LG.D2) of the Podocarpus neriifolius root sample used as starting material. The results from these procedures shared many similarities with the P. concentricum-aided hydrolysis, and these will be discussed in the following paragraphs.

3.2.1. Monitoring P. expansum-assisted biotransformation of the aqueous extract AA06795LG.D2 through 1H NMR spectroscopy As described earlier, using a 250 mg-sample of the crude aqueous extract of P. neriifolius did not lead to its successful and/or detectable P. concentricum-aided biotransformation. In the current fungal fermentation, the same quantity of this starting material was added to different culture broths in the first set of biotransformation procedures. In contrast to the parallel study conducted in the previous Penicillium species, this procedure involved the use of a control fermentation flask composed of the fungus only, which facilitated detection of differences within the podolactone-containing fungal cultures through comparison of their 1H NMR spectra, and hence the detection of the compounds of interest, the podolactones. Thus, aliquots from this control flask were collected and partitioned at each fermentation endpoint of the study flasks. The first flask

(I), with the plant aqueous extract added on the same day as the start of the fermentation, was sampled on day 2 and terminated on the following day. Spectroscopic profiling of the aqueous (PN-D2-PE-I-2d-W and PN-D2-PE-I-3d-W) and ethyl acetate-soluble partitions

(PN-D2-PE-I-2d-E and PN-D2-PE-I-3d-E) resulted from each endpoint, but did not suggest the occurrence of the targeted hydrolysis. The EtOAc partitions from both endpoints of this fermentation were comparable to the control EtOAc-soluble partition at the corresponding endpoints (PN-D2-PE-IV-2d-E and PN-D2-PE-IV-3d-E), suggesting the

215

absence of podolactones or any metabolites foreign to P. expansum (Figure 111). In contrast, the 1H NMR spectroscopic profiles of the water-soluble partitions of flask I at the

2- and 3-day endpoints still showed the presence of the starting material 89.

Figure 111. Overlay of 1H NMR spectra of the EtOAc partitions from flasks I and IV at days 2 and 3.

Measured in CD3OD at 400 MHz, (a) PN-D2-PE-I-2d-E, (b) PN-D2-PE-IV-2d-E, (c) PN-

D2-PE-I-3d-E, (d) PN-D2-PE-IV-3d-E

(d)

(c)

(b)

(a)

Analysis of the 1H NMR spectra of the partitions obtained from flasks II and III led to the same observations as in the above-described fermentation (flask I). Accordingly, the

1H NMR spectra of the EtOAc partitions from both fermentations matched the corresponding partitions from the control (flask IV, days 3 and 4) (Figure 112), as did the

EtOAc-soluble fractions from flask I. Moreover, overlay of the spectra from the analogous 216

partitions from the first and second flasks further confirmed their similarities. Likewise, the starting material remained in the aqueous partitions of both flasks II and III. From these observations, it can be concluded that the quantity of the starting material used in this first set of biotransformation procedures, as was the case with the similar study conducted in P. concentricum, was not sufficient for detectable hydrolysis to the cytotoxic podolactone 78.

Figure 112. Overlay of 1H NMR spectra of the ETOAc partitions from flasks I and IV at days 2 and 3.

Measured in CD3OD at 400 MHz, (a) PN-D2-PE-II-2d-E, (b) PN-D2-PE-IV-3d-E, (c) PN-

D2-PE-III-3d-E, (d) PN-D2-PE-IV-4d-E

(d)

(c)

(b)

(a)

However, these results suggest some similarities between the two Penicillium species regarding their interactions with the addition of the exogenous podolactone glucoside 89. It appeared that a greater quantity of the plant aqueous extract was necessary 217

to prompt fungal biotransformation, as observed in the P. concentricum fermentation procedures. Hence, in a second set of P. expansum fermentation experiments, 2 g-samples of the plant material were added to each of two fungal culture broths. The fermentation procedure from the first flask (I) was monitored at three endpoints, on days 2, 7, and 14, respectively, while that from the second flask was only analyzed upon termination of the procedure, on day 15. As predicted, increasing the amount of starting material was necessary for the targeted biotransformation to take place, as demonstrated on the 1H NMR spectra obtained at day 7 and subsequent endpoints of the fermentation in the first flask and the spectroscopic profiles of partitions from the second flask. However, the formation of the aglycone was not fully apparent on the second day (2-day endpoint).

Figure 113. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-I-2d-W and (b) 89

Measured in CD3OD at 400 MHz.

(b)

(a)

Apart from the signals in the methyl region resonating at δH 1.44 ppm (s), 1.36 ppm

(d, 6.4), and 1.21 ppm (s), equivalent to the methyl protons at positions H-18, 17, and 20 218

of the aglycone 78, respectively, the remaining key characteristic resonance signals observed in the 1H NMR spectrum of the EtOAc-soluble partition of this endpoint only suggested the presence of the starting material 89. Furthermore, analysis of the aqueous partition of the fungal mixture extracted at this 2-day endpoint (PN-D2-PE-I-2d-W) indicated that the reaction had not fully progressed, owing to the 1H NMR spectroscopic profile displaying intense peaks corresponding to the starting plant material (Figure 113).

Figure 114. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-I-7d-W, (b) 89, and (c)

Measured in CD3OD at 400 MHz.

(c)

(b)

(a)

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At day 7 of the same fungal culture, however, the 1H NMR spectroscopic profile of the EtOAc partition exhibited similar patterns to those observed in the analogous partition from the P. concentricum-assisted biotranformation described in section 3.1.2. As seen in

Figure 114, the 1H NMR spectrum of the 7-day EtOAc partition, though mainly constituted of fungal metabolites, displayed overlapped peaks indicating the presence of both the podolactone glucoside 89 and its aglycone derivative 78. For instance, two overlapped peaks at δH 6.18 ppm and 1.20 ppm were equivalent to protons at the H-11 and H3-20 positions, respectively, of both compounds.

Moreover, the two doublets (J = 6.4 Hz) resonating at 1.35 and 1.38 ppm, exactly overlapped with the analogous peaks corresponding to the H3-17 of 78 and 89, respectively.

To confirm this observation, both these methyl protons were each irradiated for a selective

1D-TOCSY NMR spectroscopy experiment. The resulting 1D-TOCSY NMR spectra each revealed the spin network composed of the protons at positions H-14, H-15, and H3-16 corresponding to the above compounds.

At the following endpoint (day 14), the 1H NMR spectrum of the EtOAc-soluble fraction from the fungal mixture (PN-D2-PE-2g-14d-E) contained inumakilactone A (78) as the only podolactone constituent (Figure 116), while the aqueous partition (PN-D2-PE-

2g-14d-W) no longer contained any of the starting podolactone glucoside 89, as seen in its 1H NMR spectrum (Figure 115). Hence, these spectra suggested that by the 14-day endpoint, the biotransformation of compound 89 into its aglycone 78 had come to completion.

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Figure 115. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-14d-W and (b) 89

Measured in CD3OD at 400 MHz (δH 0.90-1.60 ppm region).

(b)

(a)

Figure 116. Detection of 79 as biotransformation product in PN-D2-PE-2g-14d-E by 1H

NMR spectroscopy

1 H NMR spectra (δH 0.93-1.98 ppm region) of (a) PN-D2-PE-2g-14d-E, (b) 78, and (c) 89.

Measured in CD3OD at 400 MHz.

(c)

(b)

(a)

Consequently, the fermentation procedure in the second flask was terminated at day

15. A white precipitate, later identified as 78 by 1H NMR fingerprinting (Figure 117), was 221

collected from the EtOAc-soluble mixtures of both flasks at days 14 (PN-D2-PE-2g-14d-

EP), and 15 (PN-D2-PE-2g-15d-EP), respectively.

Figure 117. Overlay of the 1H NMR spectra of (a) PN-D2-PE-2g-14d-EP, (b) PN-D2-PE-

2g-15d-EP, and (c) 78

(c )

(b)

(a)

From these results, one can conclude that both Penicillium species used in the biotransformation procedures conducted in this study could carry out the targeted hydrolysis of compound 89 into its derivative 78. As with P. concentricum, 250 mg of the plant aqueous extract did not result in a measurable production of the podolactone of interest, possibly for reasons related to the accessibility of the fungus to the podolactone glucoside, or simply the low concentration of the product, rendering its detection laborious if not impossible, through the current NMR spectroscopic method. Furthermore, the fermentation from the first flask of this batch exemplifies a practical application of the 1H 222

NMR spectroscopy-based fingerprinting method as a tool to monitor the progression of a chemical reaction, or more precisely in this case, that of a fungal-assisted biotransformation of a phytochemical. This method allowed the observation of the podolactone composition of the partitioned fungal mixtures, notably in the EtOAc extract, as it progressed from only the glucoside at the 2-day endpoint, to a mixture of both podolactone derivatives at day 7, and finally to the aglycone alone at the last endpoint (day 15), all based on the respective

1H NMR spectroscopic profiles.

3.2.2. 1D-NMR spectroscopy for the structural identification of inumakilactone A (78) as a fungal-assisted biotransformation product As mentioned previously, the production of inumakilactone A (78) was detected in the 7-day EtOAc partition of the fungal mixture, and it was confirmed by 1D-TOCSY

NMR obtained from the selective excitation of the H3-17 doublet protons. This 1D-TOCSY spectrum revealed resonance signals at δH 3.90 ppm and 4.47 ppm, corresponding to the adjacent H-15 and H-14 protons, respectively. Selecting this spin network permitted a differentiation between compound 78 and its glucoside 89 when both were present in the fungal mixture, and confirmed by comparison with the 1H NMR spectrum of 78, that despite the shifting of the NMR signals in the mixture, the compound was indeed the hydrolyzed glucoside. This product later precipitated out of the fungal mixture EtOAc partitions, and the white solid collected was analyzed by 1H NMR spectroscopy, revealing a pure compound, the spectrum of which exactly matching that of the previously isolated

78. Therefore, the isolation of this podolactone from the fungal mixture confirmed the predicted biotransformation reaction.

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4. Biological activity 4.1. In vitro cytotoxicity evaluation of components obtained from the large-scale extraction of P. neriifolius root sample 4.1.1. Antiproliferative activity of the main partitions of the methanol crude extract in a panel of four cancer cell lines The methanol extract generated from the large-scale extraction of P. neriifolius was evaluated in a panel of four human cancer cell lines, constituted of melanoma, ovarian, colon, and breast cancer cells (Appendix C). As detailed in Table 15, this crude extract exhibited comparable antiproliferative activity across all four cell lines, with the breast cancer cells (MDA-MB-231) being the least susceptible. While the equivalent cytotoxic activity of the emulsified precipitate (AA06795LG.D0) formed during the hexanes-MeOH liquid-liquid partition was at first surprising, this biological property was later explained through the identification of its main constituent, inumakilactone A (78) previously shown to be one of two cytotoxic isolates in the small-scale investigation of P. neriifolius. As expected, of the four samples (D1-D4) obtained from the partition of the above-mentioned extract, the EtOAc partition (AA06795LG.D3) in this large-scale separation displayed the highest activity, notably against the melanoma and ovarian cancer cells with estimated IC50 values below 2 µg/mL (IC50 <2 µg/mL) (Table 15). Moreover, the antiproliferative components of the EtOAc partition seemed to have been concentrated in its D3.1 fraction.

Considering the above-mentioned activity of 78, and its identification from this fraction, it is pertinent to imply that the observed bioactivity may be due mainly to this compound.

Unlike its hexane-MeOH counterpart, the emulsified product of the EtOAc-H2O partition did not negatively affect the growth of the cancer cells at the concentrations used in this study. 224

4.1.2. Antiproliferative activity of the identified compounds (85-89, 91) from the large-scale extraction of P. neriifolius in a panel of four human cancer cell lines Compounds 85-88, identified from the dereplication of the hexanes partition

(AA06795LG.D1), 89 obtained from the aqueous partition, and 91 from the EtOAc extract, were evaluated for their potential cytotoxicity in the same panel of cancer cells as their original fractions, and none of these were active in this bioassay (IC50 >10 µM). On the other hand, compound 91 displayed weak cytotoxicity with IC50 values of 7.7 and 8.1 µM against the melanoma (MDA-MB-435) and breast (MDA-MB-231) cancer cell lines, respectively, whereas the growth of the ovarian (OVCAR3) and colon (HT-29) cell subtypes were not affected by this compound at the concentrations used.

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Table 15. Antiproliferative activity of fractions from the large-scale extraction of P. neriifolius in a panel of four human cancer cell lines

Values in % viable cells.

MDA-MB-435 (melanoma) OVCAR3 (ovarian) HT-29 (colon) MDA-MB-231 (breast) 20 μg/mL 2 μg/mL 20 μg/mL 2 μg/mL 20 μg/mL 2 μg/mL 20 μg/mL 2 μg/mL AA06795LG 39 75 30 79 42 69 38 86

AA06795LG.D0 19 44 21 55 19 70 45 56 AA06795LG.D1 84 95 84 96 71 87 48 93 AA06795LG.D1.4 94 96 81 93 82 87 59 90 AA06795LG.D1.6 88 95 79 92 71 85 45 88 AA06795LG.D2 91 100 81 96 79 82 72 92 AA06795LG.D2.2 83 100 79 93 78 88 64 86 AA06795LG.D2.3 72 100 70 89 78 94 55 88 AA06795LG.D3 21 39 22 41 21 60 50 50 AA06795LG.D3.1 23 71 27 73 34 78 51 53 AA06795LG.D3.2 41 93 39 94 57 87 50 82 AA06795LG.D3.3 59 86 47 100 59 83 52 81 AA06795LG.D4 85 100 95 99 85 95 77 88

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4.2. Biological activity of inumakilactone A (78) in vivo Inumakilactone A (78) exhibited antiproliferative activity across three of the four cancer cell lines used in the in vitro bioassay, and hence, it was evaluated at the College of

Pharmacy, University of Illinois at Chicago in a xenograft murine model of the hollow fiber assay using fibers implanted with the human colon (HT-29), ovarian (OVCAR3), and melanoma (MDA-MB-435). However, compound 78 did not significantly affect cell survival when compared to the vehicle control in all three cell lines tested. Therefore 78 was deemed inactive in this in vivo assay.

4.3. Insecticidal potential of inumakilactone A (78) and inumakilactone A-15-O-β- glucoside (89) Considering the report on the potential insecticidal and antifeedant activities associated with plants from the genus Podocarpus (Barrero et al., 2003), such as the termiticidal property of P. neriifolius and the podolactones derivatives, inumakilactone A

(78) and its glucosidic derivative (89) were evaluated for their antifeedant activity against the Aedes aegypti mosquito strain. However, these podolactones did not show activity in this assay.

4.4. Cytotoxic activity of extracts obtained from Penicillium concentricum biotransformation procedures The partitioned P. concentricum fermentation mixtures from the second batch were evaluated for their cytotoxicity against two human cancer cell lines, namely, MCF-7

227

(breast) and A2780 (ovarian). As presented in Tables 15 and 16, in both assays, only the ethyl acetate-soluble fractions exhibited antiproliferative activity. This can be explained by the presence of the aglycone, inumakilactone A (78) in all these partitions and not in their aqueous counterparts. This conclusion was better supported by the trend in the cytotoxicity activity against the ovarian A2780 cell line, as even the original starting material was inactive (IC50 >20 µg/mL), while the fungal EtOAc partitions were moderately cytotoxic.

Furthermore, the 7-day EtOAc partition was more active than its 2- and 6-day counterparts, probably due to higher concentrations of 78 in the former partition. Interestingly, the starting podolactone glucoside 89 was somewhat active in the MCF-7 human breast cancer cell line (IC50 = 9.9 µg/mL). It was, thus, not surprising that the resulting EtOAc partitions containing both 78 and 89 exhibited more potent cytotoxicity against these cells (2.8 µg/mL

< IC50 <3.8 µg/mL), compared to the A2780 cells. However, the trend remained the same in both cell lines, still relating the antiproliferative activity of the fermented extracts to their content in compound 78.

4.4. Cytotoxic activity of extracts obtained from Penicillium expansum biotransformation procedures The aqueous and ethyl acetate-soluble fractions obtained from the partitioning of the fungal mixtures in both sets of the P. expansum-facilitated biotransformation procedures were evaluated for their antiproliferative activities against the human breast and ovarian cancer cell lines, MCF-7 and A2780, respectively. Unlike their P. concentricum counterparts, the potency of which positively correlated with the production of 228

inumakilactone A 78 in both cell lines, the extracts from P. concentricum display some variability in their cytotoxic effect. As documented in Tables 16 and 17, all the aqueous partitions from the first biotransformation batch (250 mg starting plant material) were non- cytotoxic (IC50 >20 µg/mL) against both cell lines. In contrast, all the EtOAc partitions from this batch were potent against the MCF-7 cell line, with IC50 values ranging from 18.8 to 1.8 µg/mL. Since the EtOAc partitions collected from the control fungal culture were cytotoxic against these cancer cells, their activity is, thus, attributed to the fungal metabolite(s). While the cytotoxic activity observed for the plant extract-supplemented fungal extracts (EtOAc partitions, flasks I-III) was consistent with fungal growth, with IC50 values decreasing from 13.1 µg/mL on day 2 to 4.8 µg/mL on day 3 for flask I, and from

2.6 µg/mL for flask II (day 2) to 1.8 µg/mL for flask III (day 3), results observed in the control slightly deviated from this trend, notably with the unexpected low potency of the

EtOAc extract obtained at day 3 from this control. As seen in the above tables, the partitions from the first biotransformation batch exhibited only weak activity against the more robust

A2780 ovarian cancer cells, as their corresponding IC50 values ranged from 12.7-19.5

µg/mL, suggesting that this cell line was less vulnerable to the P. concentricum metabolites. However, the trend in potency followed that observed using the MCF-7 cells

(Table 17).

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Table 16. Antiproliferative activity fungal biotransformation fractions from two

Penicillium strains against the A2780 human ovarian cancer cell line

PN-D2: Podocarpus neriifolius aqueous extract; PC: Penicillium concentricum; PE:

Penicillium expansum; I, II, III and IV: flask numbers; d: day; E: EtOAc partition; W: aqueous partition

Sample IC50 (μg/mL) Sample IC50 (μg/mL) PN-D2-PE-I-2d-E >20 PN-D2-PE-7d-E 0.9

PN-D2-PE-I-2d-W >20 PN-D2-PE-7d-W 2.5

PN-D2-PE-I-3d-E 18.0 PN-D2-PE-14d-E 0.6

PN-D2-PE-I-3d-W >20 PN-D2-PE-14d-W >20

PN-D2-PE-II-2d.E 17.0 PN-D2-PE-16d-E 0.2 PN-D2-PE-II-2d.W >20 PN-D2-PE-16d-W 11.9 PN-D2-PE-III-3d.E 13.0 PN-D2-PE-15d-EP 0.3 PN-D2-PE-III-3d-W >20 PN-D2-PE-15d-EM 0.4 PN-D2-PE-IV-2d-E 19.5 PN-D2-PE-15d-W >20 PN-D2-PE-IV-2d-W >20 PN-D2-PC.2d-E 12.8 PN-D2-PE-IV-3d-E >20 PN-D2-PC.2d-W >20 PN-D2-PE-IV-3d-W >20 PN-D2-PC.6d-E 13.8 PN-D2-PE-IV-4d-E 12.7 PN-D2-PC.6d-W >20 PN-D2-PE-IV-4d-W >20 PN-D2-PC.7d-E 9.3 PN-D2-PE-2d-E 3.8 PN-D2-PC.7d-W >20 PN-D2-PE-2d-W >20 PN-D2 >20 Taxol (positive control) 0.01

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Table 17. Antiproliferative activity fungal biotransformation fractions from two

Penicillium strains against the MCF-7 human breast cancer cell line

Sample IC50 (μg/mL) Sample IC50 (μg/mL) PN-D2-PE-I-2d-E 13.1 PN-D2-PE-7d-E <0.16

PN-D2-PE-I-2d-W >20 PN-D2-PE-7d-W 1.8

PN-D2-PE-I-3d-E 4.8 PN-D2-PE-14d-E <0.16

PN-D2-PE-I-3d-W >20 PN-D2-PE-14d-W 18.6

PN-D2-PE-II-2d.E 2.6 PN-D2-PE-16d-E <0.16 PN-D2-PE-II-2d.W >20 PN-D2-PE-16d-W 5.4 PN-D2-PE-III-3d.E 1.8 PN-D2-PE-15d-EP 0.2 PN-D2-PE-III-3d-W >20 PN-D2-PE-15d-EM <0.16 PN-D2-PE-IV-2d-E 6.3 PN-D2-PE-15d-W >20 PN-D2-PE-IV-2d-W >20 PN-D2-PC.2d-E 3.1 PN-D2-PE-IV-3d-E 18.8 PN-D2-PC.2d-W >20 PN-D2-PE-IV-3d-W >20 PN-D2-PC.6d-E 3.8 PN-D2-PE-IV-4d-E 2.0 PN-D2-PC.6d-W >20 PN-D2-PE-IV-4d-W >20 PN-D2-PC.7d-E 2.8 PN-D2-PE-2d-E 0.6 PN-D2-PC.7d-W >20 PN-D2-PE-2d-W >20 PN-D2 9.9 Camptothecin (positive control) 0.004

The ethyl acetate-soluble partitions obtained from the second biotransformation batch conducted in P. expansum cultures were significantly more potent than those from

231

the previous batch, with IC50 values ranging from <0.16 to 0.6 µg/mL and 0.2 to 3.8 µg/mL against the MCF-7 and A2780 cells, respectively (Tables 16 and 17). As these partitions have been shown to contain the hydrolyzed podolactone, inumakilactone A (78), their observed potency in the MCF-7 cell line is, therefore, due simultaneously to the presence of both cytotoxic fungal and plant metabolites. On the other hand, the poor potency of the

P. expansum secondary metabolites against the A2780 cells suggest that the strong cytotoxic activity observed in the EtOAc partitions from the second biotransformation batch is mainly caused by their content in the cytotoxic phytochemical, 78. Owing to the very low IC50 values observed in the MCF-7 bioassay, no clear trend in cytotoxic activity could be deduced from the results. However, consistent with previous data, potency against the A2780 cell line increased with the progression of the biotransformation, which is with the production of inumakilactone A. Thus, the IC50 values decreased from 3.8 µg/mL at the 2-day endpoint to 0.2 µg/mL at the last endpoint (16-day), corresponding to the complete hydrolysis of compound 89 to 78. However, while most aqueous-soluble partitions from this second batch were inactive in both cell lines, those obtained at the 7- and 16-day endpoints displayed some unexpected activity. This odd result may have arisen from erroneous mishandling of the samples prior or during the bioassay procedure, and thus, will be repeated prior to publication. It is also worth mentioning that despite this strong potency against the cancer cell lines used in the present in vivo bioassays, compound

78 did not seem to affect the fungal growth, as was observed by the growth trend during the fermentation procedures.

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E. Conclusion

The dereplication method described in this dissertation work, though grounded on the basic concepts for more effective discovery of novel secondary metabolites, inclusive of polarity-based pre-fractionation of extracts, followed by chromatographic and spectroscopic profiling (e.g. HPLC-UV, MS, NMR), proved to be a useful tool for an accelerated detection and isolation of compounds from the conifer Podocarpus neriifolius, and facilitated a new application of selective total correlation spectroscopy (1D-TOCSY) in a plant natural products isolation procedure.

First, subjecting extracts and fractions to 1H NMR spectroscopy, both for fingerprinting against previously isolated secondary metabolite from the early part of this study (Chapter 2) led to the dereplication of known compounds through spectral data comparison, and the detection of “unfamiliar” proton signals resulted in the isolation and structural determination of compounds not previously isolated in this study (85-92, Figure

47). Spectroscopic profiling of the hexane extract was a good example of a “proof of concept”, as its further purification was solely 1H NMR spectroscopic-scanning-based, for the isolation and determination of the compounds corresponding to the proton signals of interest. The 1H NMR spectroscopy-based fingerprinting of the aqueous extract of this plant to search for glycosidic podolactone derivatives, was successful as it resulted in the isolation of compound 89, but more importantly, the screening of the fractions from this extract allowed for the discovery of this compound as the major component of the plant

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(nearly 90% of the aqueous extract), which resulted in the use of this non-cytotoxic extract as a starting material for fungal-assisted biotransformation studies affording its active aglycone, inumakilactone A (78). The 1H NMR spectroscopy-guided fractionation of the

EtOAc extract of P. neriifolius was equally useful, as it accelerated the prioritization of fractions of interest towards the detection and isolation of podolactone derivatives. While

NMR spectroscopy was the main method highlighted here, the role of HPLC-UV was extremely significant, especially for the fingerprinting of the pre-fractionated EtOAc extract for the detection of the new podolactone glucoside (83). However, this chromatographic method coupled with UV spectra was not sufficient, and even led to a false lead (92) possessing a similar retention time and UV pattern as the target compound.

Thus, it is important to combine these methods for best results. Moreover, though not heavily used in this study, mass spectrometry would be a necessary tool to complement the above methods (HPLC-UV-NMR) to increase the effectiveness of the overall dereplication procedure.

Similar aspects of the selective 1D-TOCSY NMR method has been used recently in the profiling and dereplication of microbial mixtures (MacKinnon et al., 2016), and these have required the use of algorithms, leading to a seemingly more complicated process, especially for application in more complex plant extracts. However, unlike this reported method aiming at the total metabolomic profile of the natural source investigated, the current study was focused on specific compounds and compound classes, the podolactones.

Thus, the development of a selective 1D-TOCSY NMR spectroscopy-based fingerprinting

234

of plant extract, in the present study, resulted in facilitated and time-effective fractionation and detection approaches for the targeted isolation of a newly identified secondary metabolite, namely, the B-type podolactone, nagilactone G-2β-O-β-D-glucoside (83).

An additional application of both the 1H and 1D-TOCSY NMR spectroscopy-based dereplication method was performed in monitoring the progress of the fungal-assisted hydrolysis of 89 to 78, and this procedure further validated the usefulness of this dereplication. Equally, if not more, significant is the fungal biotransformation procedure itself, which, according to the consulted literature, is the first of such reaction performed with any class of podolactone.

In conclusion, this dissertation work, initiated with the search for new potential anticancer leads from the Vietnamese Podocarpus neriifolius D. Don resulted in the discovery of a new podolactone glucoside (83), and the isolation or detection of a several

B-type podolactones along with other diterpenoids. Furthermore, this study reported the in vivo evaluation of inumakilactone A 78 using the murine hollow fiber assay. Although 78 was proven inactive, the in vivo evaluation of this compound in this bioassay has not been reported previously, thus adding new knowledge for this compound, as well as providing a full spectrum of the preliminary natural product drug discovery process. Moreover, additional contributions of this work constitute a new application of 1H and 1D-TOCSY

NMR spectroscopy in the dereplication of plant natural products, and the first report of

Penicillium-assisted biotransformation of a podolactone glucoside (89) of plant origin to its aglycone derivative (78) using two fungal strains. Therefore, this procedure has the

235

potential to expand the possibilities of new discoveries, not only with the podolactones and their plant sources, but also with other natural sources as well.

236

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Appendix A: Permission to Reproduce Published Materials

Permission to extract and reproduce Figure 6 from (Tropicos.org, 2018)

“Citation: Cite this page: Tropicos.org. Missouri Botanical Garden. 07 Jun 2018

Name: Podocarpus neriifolius D. Don; Long Description: Flora of China Illustrations vol.

4, fig. 93, 1-4…

License Creative Commons Attribution-NonCommerical-ShareAlike 3.0 Unported (CC

BY-NC-SA 3.0) https://creativecommons.org/licenses/by-nc-sa/3.0/

…You are free to share (copy and redistribute the material in any medium or format).

License terms: …You must give appropriate credit, provide link to license, and indicate of changes were made. You may do so in any reasonable manner, but not in any way that suggests the licensor endorses you or your use…” (Tropicos.org, 2018)

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Appendix B: HT-29 Cytotoxicity Using an SRB Assay Protocol (OSU)

Evaluation of the cytotoxic activity of the plant crude extract, fractions and pure compounds against the HT-29 human colon cancer cell line (ATCC HTB-38) was performed by

Dr. Hee-Byung Chai (former Research Scientist, College of Pharmacy, OSU) using an established sulfhorodamine B (SRB) assay protocol (Skehan et al., 1990) in which cell viability was measured in 96-well plates based on cellular protein content. In this assay HT-29 cells were seeded in 96- well microtiter plates in 190 µL of cell culture medium per well, to which 10 µL of either the test samples, the positive control (paclitaxel) or 10% DMSO as the negative control, were added at suitable concentrations. The plates were then incubated for 72 h at 37 °C in a humidified 5% CO2 environment. Following this incubation phase, the cells were fixed with 100 µL/well of cold 20% trichloroacetic acid (TCA), and were further incubated at 4 °C for 30 min. Then, the plates were washed four times with tap water and air-dried. The obtained fixed cells were further stained with

100 µL/well of 0.4 % (wt/v) SRB in 1% acetic acid, incubated for 30 min at room temperature, and the unbound dye was removed by rinsing the plates four times with 1% acetic acid. To the air-dried plates were added 200 µL/well of 10 mM Tris base (pH 10), and the plates were placed on a gyratory shaker for 5 min to solubilize the protein-bound dye. The optical density was measured on a microtiter plate reader at 515 nm. Cell viability was expressed as IC50 values calculated using a nonlinear regression analysis (TableCurve 2DV4; AiSN Software Inc., Mapleton, OR, USA).

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Appendix C: Cytotoxicity Assay Using a Panel of Four Human Cell Lines (UIC)

Fractions and pure compounds obtained from the extraction of Podocarpus neriifolius were evaluated for their in vitro cytotoxic activities against a panel of four human cancer cell lines, namely, MDA-MB-435 (melanoma), MDA-MB-231 (breast), OVCAR3 (ovarian), and HT-29

(colon). The various sets of bioassays were performed by Dr. Wei-Lun Chen and Mr. Austin A.

Czarnecki at the University of Illinois at Chicago (UIC) College of Pharmacy under the asupervision of Dr. Joanna E. Burdette. Described below is the protocol provided by these colleagues:

“Human melanoma cancer cells MDA-MB-435, human breast cancer cells MDA-MB-231, human ovarian cancer cells OVCAR3 and human colon cancer cells HT-29 were purchased from the American Type Culture

Collection (Manassas, VA). The cell line was propagated at 37°C in 5% CO2 in RPMI 1640 medium, supplemented with fetal bovine serum (10%), penicillin (100 units/ml), and streptomycin (100 µg/ml). Cells in log phase growth were harvested by trypsinization followed by two washing to remove all traces of enzyme.

A total of 5,000 cells were seeded per well of a 96-well clear, flat-bottom plate (Microtest 96®, Falcon) and incubated overnight (37°C in 5% CO2). Samples dissolved in DMSO were then diluted and added to the appropriate wells. The cells were incubated in the presence of test substance for 72 h at 37°C and evaluated

® for viability with a commercial absorbance assay (CellTiter 96 AQueous One Solution Cell Proliferation

Assay, Promega Corp, Madison, WI) that measured viable cells. IC50 values are expressed in µM relative to the solvent (DMSO) control.”

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Appendix D: In vivo Evaluation of inumakilactone A (78) in the Murine Hollow

Fiber Assay

Prior to the submission of inumakilactone A (78) for in vivo antitumor evaluation in the hollow fiber assay, this compound was tested against the human melanoma (MDA-MB-435) and ovarian (OVCAR3) cancer cell lines, according to the protocol summarized in Appendix C.

Compound 78 exhibited cytotoxicity with IC50 values of 0.5 and 1.4 μg/mL against the MDA-MB-

435 and OVCAR3 cell lines, respectively. This bioassay was performed by Mr. Austin A.

Czarnecki, a former member of Dr. Joanna E. Burdette research group at UIC College of Pharmacy.

Compound 78 was then evaluated for its in vivo antitumor activity against the above two cell lines and the HT-29 human colon cancer cell line. This bioassay was performed by Mr. Daniel

D. Lantvit under the supervision of Dr. J.E. Burdette. The method employed was provided by Mr.

Lantvit as follows:

“Human ovarian cancer cells OVCAR3 (4x106 cells/mL) are cultured in hollow fibers on day -2. Human colon cancer cells HT29 (1x106 cells/mL) and human breast cancer cells MDA-

MB-435 (2.5x106 cells/mL) are cultured in hollow fibers on day -1. All cell lines were used to evaluate inumakilactone A. Seven-week-old immunodeficient NCr nu/nu mice were purchased from

Taconic Laboratory and housed in microisolation cages at room temperature and a relative humidity of 50–60% under 12:12 h light–dark cycle. All animal work was approved by University of Illinois at Chicago Animal Care and Use Committee (protocol number 16-035), and the mice

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were treated in accordance with the institutional guidelines for animal care. Inumakilactone A was dissolved initially in DMSO and subsequently diluted with 60% PEG 300 and 30% water. Hollow fibers were implanted into the abdominal cavity on day 0 (n= 6 control, 61 mg/kg, and 32 mg/kg).

The mice were injected i.p. once daily for 4 days (days 3-6) with vehicle, inumakilactone A or the positive control (paclitaxel). Each mouse was weighed daily during the study. One animal from each treatment group died and the rest showed signs of toxicity after the second injection. Doses were reduced to half. All the remaining mice were sacrificed on day 7. The fibers were retrieved, and viable cell mass was evaluated by a modified MTT [3-(4,5-dimethylthiazol-2-yl)-2,5- diphenyltetrazolium bromide] assay”.

In addition, Dr. Xiaoli Zhang (Center for Biostatistics, OSU) provided the biostatistics analysis of the obtained in vivo data, with the following comments:

“ANOVA was used for analysis, and the results showed that Inum had no significant effect on cell survival comparing to vehicle control for all the three tested cell lines as seen in the following table and the plot.

Inum comparison to positive and negative controls

Effect Compounds Compounds Estimate Standard p-values 95% CI Error

HT29 Inumakilactone A Taxol 0.5095 0.2212 0.0468 0.009044 1.0100

HT29 Inumakilactone A Vehicle 0.02000 0.1649 0.9061 -0.3530 0.3930

HT29 Taxol Vehicle -0.4895 0.2086 0.0435 -0.9613 -0.01767

MDA-MB-435 Inumakilactone A Taxol 0.8977 0.2673 0.0073 0.3022 1.4932

MDA-MB-435 Inumakilactone A Vehicle 0.2055 0.1934 0.3130 -0.2255 0.6365

MDA-MB-435 Taxol Vehicle -0.6922 0.2608 0.0242 -1.2733 -0.1110

OVCAR3 Inumakilactone A Taxol 0.5037 0.2231 0.0475 0.006689 1.0007

OVCAR3 Inumakilactone A Vehicle -0.2033 0.1614 0.2365 -0.5630 0.1564

OVCAR3 Taxol Vehicle -0.7070 0.2177 0.0088 -1.1920 -0.2220

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Appendix E: Cytotoxicity Evaluation of Fungal Biotransformation Fractions against

a Human Ovarian and a Breast Cancer Cell Line Using the SRB Assay

The ethyl acetate and water soluble fractions obtained from the liquid-liquid partition of fungal fermentation cultures (Penicillium concentricum and P. expansum) used for the biotransformation of the aqueous extract of P. neriifolius were evaluated in vitro for their antiproliferative activity against a human breast (MCF-7) and an ovarian (A2780) cancer cell line, using a sulforhodamine B (SRB)-based assay. These bioassay procedures were performed by Ms.

Fengrui Wang (Rae) and Mr. Choon Yong Tan, two colleagues from Dr. L. H. Rakotondraibe research group, and Mr. Tan provided the corresponding assay protocol, described below.

The test samples were dissolved in 100% DMSO, and then subjected to a ten-fold serial dilutions with sterile H2O. This was followed by a five-fold serial dilution in 10% DMSO. Using

96-well microtiter plates, 10 μL of the test compounds, positive control (Taxol®for A2780, or camptothecin for MCF-7), or the negative control (10% DMSO), at the appropriate concentrations were added in duplicate to each well. The wells containing the test compound and the positive control were then seeded with 190 μL of harvested A2780 cancer cells at 25 × 104 cells per well, whereas 190 μL of fresh culture medium was added to the wells containing the negative control.

The microtiter plates were then incubated for three days in 5% CO2 and at 37°C. Following the incubation period, the cells were fixed with 100 μL cold trichloroacetic acid (20%) and further incubated at 4°C for 30 min. Then, the plates were washed with tap water and dried at room

275

temperature. Once dry, the fixed cells were stained with 100 μLper well of sulforhodamine B (SRB) in acetic acid (1%) for 30 min at room temperature, and then the excess SRB was washed with 1% acetic acid.. Following a 30 min drying period, 200 μL 0f 10 mM unbuffered Tris base (pH10) were added to each well. The plates were then placed on a shaker for 1 h to solubilize bound SRB stain.

Subsequently, the absorbance of the stained wells was measured at 515 nm on a microplate reader, and the IC50 values were calculated using a non-linear regression analysis (Table Curve2Dv4; AISN

Software, Inc.; Mapleton, OR, USA).

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Appendix F: Penicillium concentricum Source, Identification, and Culture

The endophytic fungus Penicillium concentricum was isolated from the liverwort,

Trichocolea tomentella (Trichocoleaceae), collected in Newport, VA, USA. Following sterilization and growth on agar plate, single colonies of the fungus were streaked onto potato dextrose agar plates. Identification of the fungus was conducted by Dr. Chad

Rappleye (Department of Microbiology, OSU), based on morphological properties and sequencing of its internal transcribed spacer (ITS). Upon comparison with accessions from the GenBank database, this endophytic fungus showed 100% similarity with P. concentricum, and thus was positively identified. The specimen of P. concentricum was further incubated on potato dextrose agar plates, or stored in cryogenic tubes in -80°C freezer. Colonies from either the agar plates or the frozen samples were then fermented in potato dextrose broth, and these fermentation cultures were used in the biotransformation studies conducted in the current dissertation work. Detailed collection, treatment, identification, and culture of P. concentricum are described in a recent publication authored by members of this group (Ali et al., 2017). Depicted below are the liverwort, T. trichocolea and an agar plate culture of P. concentricum.

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Appendix G: Penicillium expansum Source, Identification, and Culture

Colonies of the fruit pathogen, Penicillium expansum, were collected from contaminated plums by Dr. L. Harinantenaina Rakotondraibe, and cultivated on potato dextrose agar plates similar to P. concentricum (see Appendix F). The fungal species was identified by Dr. Chad

Rappleye, at the Department of Microbiology, The Ohio State University, similar to P. concentricum. A specimen of P. expansum was stored on agar plates or in cryogenic tubes (at -

80°C) at the OSU College of Pharmacy, Division of Medicinal Chemistry and Pharmacognosy.

Fermentations for biotransformation procedures were conducted using previously grown colonies in potato dextrose broth. (Photo courtesy: Dr. L. H. Rakotondraibe)

Plums contaminated by P. P. expansum

expansum

P. expansum agar plate culture

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Appendix H: HRESIMS data of compounds 85-87

The HRESIMS data for totarol, totaral, and sandaracopimaric acid (85-87) in the negative ion mode were provided by Dr. Arpad Somogyi (Campus Chemical Instrumentation Center, OSU).

These data corresponded to the respective calculated values, as shown in the following table.

Compound Molecular formula (structure) Calculated m/z [MH] Measured m/z [MH] 85 (totarol) C20H30O

285.22239 285.22241

86 (totaral) C20H28O2

299.20165 299.20165

87 C20H30O2 (sandaracopimaric acid) 301.21730 301.21733

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