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Isolation, synthesis, and biological target identification of natural products from terrestrial and marine… Centko, Ryan Matthew 2014

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ISOLATION, SYNTHESIS, AND BIOLOGICAL TARGET IDENTIFICATION OF NATURAL PRODUCTS FROM TERRESTRIAL AND MARINE ENVIRONMENTS by  RYAN MATTHEW CENTKO  B.Sc., The University of South Florida, 2008  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Chemistry)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  August 2014  © Ryan Matthew Centko, 2014   ii Abstract  Natural products offer an unparalleled resource for the discovery and development of chemical tools to be used by humans.  The terrestrial and marine environments contain unique niches where organisms chemically adapt to produce compounds that have proven useful medicinally and beyond.  In the following chapters, several classes of novel natural products from terrestrial fungi and marine sponges will be presented. In some cases, synthetic methodology, biological activity, and enzymatic target identification will also be presented.  Chapters 2 and 3 highlight ramariolides A–D (2.18–2.21) and dhilirolides A–N (3.9–3.22), two new fungal derived compound classes isolated from terrestrial sources. The structure elucidation of these compounds will be presented alongside their biological activities as antimycobacterial and insecticidal agents, respectively.  Chapter 4 contains the structure elucidation of three new members of the xestoquinone family of compounds, xestolactone A (4.19), xestosaprol O (4.20), and xestosaprol P (4.21) along with their potent inhibitory effect on human indolamine 2, 3-dioxygenase (hIDO).  A new method for synthetic access to derivatives of these compounds is demonstrated in Chapter 5 along with a brief structure activity relationship (SAR) study.    Lastly, Chapter 6 discusses latonduine A (6.9), a sponge-derived alkaloid, which has shown promise as a lead structure for the correction of cystic fibrosis (CF). Probes derived from latonduine A (6.9) have led to identification of poly (ADP-ribose) polymerase (PARP) as the enzymatic target. Methodology for the probes’ construction and SAR studies resulting in simplified synthetic analogues of latonduine will be presented.    iii Preface A version of Chapter 2 has been published. Centko, R. M.; Ramon À. García, S.; Taylor, T.; Patrick, B. O.; Thompson, C. J.; Miao, V. P.; Andersen R. J. Ramariolides A–D, Antimycobacterial Butenolides Isolated from the Mushroom Ramaria cystidiophora. J. Nat. Prod. (2012), 75, 2178-2182. I conducted all of the work involving isolation and structure elucidation/ characterization recorded therein. Dr. Brian O. Patrick provided the single crystal X-ray analysis. Dr. S. Ramon À. García, Dr. V. Miao, or T. Taylor conducted all biological assays and mushroom identification.  My supervisor Professor Raymond J. Andersen wrote the majority of the original manuscript. A version of Chapter 3 has been published in two separate manuscripts:  de Silva, E. Dilip; Williams, David. E.; Jayanetti, Dinith R.; Centko, Ryan M.; Patrick, Brian O.; Wijesundera, Ravi L. C.; Andersen, Raymond J. Dhilirolides A−D, Meroterpenoids Produced in Culture by the Fruit-Infecting Fungus Penicillium purpurogenum Collected in Sri Lanka. Org. Lett. (2011), 13, 1174–1177.  Centko, Ryan M.; Williams, David. E.; Patrick, Brian O.; Akhtar, Yasmin; Garcia Chavez, Miguel A.; Wang, Yan A.; Isman, Murray B.; de Silva, E. Dilip; Andersen Raymond J.  Dhilirolides E–N, Meroterpenoids Produced in Culture by the Fungus Penicillium purpurogenum Collected in Sri Lanka: Structure Elucidation, Stable Isotope Feeding Studies, and Insecticidal Activity J. Org. Chem. (2014), 79, 3327–3335. Isolation and structure elucidation of dhilirolides A–N (3.9–3.22) was a collaborative effort. D. Jayanetti and L. Ravi isolated the fungus in Sri Lanka. The original isolation of dhilirolide A–D (3.9–3.12), L (3.20) and M (3.21) was accomplished by Dr. David E. Williams or Dr. E. Dilip de Silva, while characterization and structure elucidation were carried out by myself (A (3.9), D (3.12), and L (3.20)) and Dr. David E. Williams (A–C (3.9–3.11), L (3.20), and M (3.21)). I isolated and characterized dhilirolides E–K (3.13–3.19) and N (3.22) in full. Dr. Brian O. Patrick provided the X-ray analyses. My supervisor Prof. Raymond J. Andersen authored the majority of the published prose for both papers. Chapters 4 and 5 are based on work conducted in collaboration with members of Prof. Grant Mauk’s laboratory, Dr. Anne Steino and Dr. Fred Rosell.  I was responsible for isolation and characterization of all of the natural products as well as the design and   iv completion of the synthetic work presented therein. Dr. Brian O. Patrick provided the X-ray analyses. A version of Chapter 6 has been published. Carlile, Graeme W.; Keyzers, Robert A.; Teske, Katrina A.; Robert, Renaud; Williams, David E.; Linington, Roger G.; Gray, Christopher A.; Centko, Ryan M.; Yan, Luping; Anjos, Suzana M.; Sampson, Heidi M.; Zhang, Donglei; Liao, Jie; Hanrahan, John W.; Andersen, Raymond J.; Thomas. David Y. Correction of F508del-CFTR Trafficking by the Sponge Alkaloid Latonduine Is Modulated by Interaction with PARP Chemistry & Biology. (2012), 19, 1288–1299. I performed the synthesis of latonduine A probe and also designed and synthesized all of the synthetic derivatives shown except where it is explicitly mentioned otherwise.                  v Table of Contents  Abstract .......................................................................................................................................... ii	  Preface ........................................................................................................................................... iii	  Table of Contents ...........................................................................................................................v	  List of Tables ............................................................................................................................... xii	  List of Figures ............................................................................................................................. xiii	  List of Schemes ........................................................................................................................ xxvii	  List of Abbreviations ............................................................................................................. xxviii	  Acknowledgements ................................................................................................................ xxxiii	  Dedication ............................................................................................................................... xxxiv	  Chapter 1: Introduction to Natural Products Chemistry ............................................................. 1	  1.1	   Natural Products Chemistry ........................................................................................... 1	  1.2	   A Brief History of the Field ........................................................................................... 1	  1.3	   Producers of Natural Products ....................................................................................... 4	  1.4	   Biosynthetic Route to Natural Products ......................................................................... 5	  1.4.1	   Polyketide Biosynthesis .......................................................................................... 6	  1.4.2	   Terpene Biosynthesis .............................................................................................. 7	  1.4.3	   Shikimate Biosynthesis ........................................................................................... 8	  1.4.4	   Non-Ribosomal Peptide and Ribosomal Peptide Biosynthesis .............................. 9	  1.5	   Methods Used for Natural Products Chemistry ........................................................... 12	  1.5.1	   Isolation ................................................................................................................. 12	  1.5.2	   Characterization .................................................................................................... 13	  1.6	   Modern Natural Products in Drug Design and Discovery ........................................... 14	  1.7	   Conclusion ................................................................................................................... 15	    vi Chapter 2: Ramariolides A–D, Novel Antimycobacterial Butenolides From the Pacific Northwest Native Coral Mushroom Ramaria cystidiophora. ................................................... 16	  2.1	   Tuberculosis ................................................................................................................. 16	  2.1.1	   Mycobacterium tuberculosis (Mtb): ..................................................................... 17	  2.1.2	   Tuberculosis Drugs: Drugs Involving the Cell Capsule of Mtb ........................... 19	  2.1.3	   Tuberculosis Drugs: Alternative Mechanisms of Action ...................................... 20	  2.2	   Mushrooms as a Source for Drug Leads ...................................................................... 22	  2.2.1	   Ramaria: Description, Distribution and Known Chemistry ................................. 23	  2.3	   Isolation of Ramariolides A–D .................................................................................... 24	  2.3.1	   Structure Elucidation of Ramariolide A ............................................................... 25	  2.3.2	   Structure Elucidation of Ramariolide B ................................................................ 28	  2.3.3	   Structure Elucidation of Ramariolide C ................................................................ 31	  2.3.4	   Structure Elucidation of Ramariolide D ............................................................... 33	  2.4	   Biological Activity of Ramariolide A .......................................................................... 35	  2.5	   Conclusions .................................................................................................................. 36	  2.6	   Experimental section .................................................................................................... 37	  2.6.1	   General Experimental Procedures ......................................................................... 37	  2.6.2	   Mushroom Sample ................................................................................................ 37	  2.6.3	   Extraction and Isolation ........................................................................................ 38	  2.6.4	   Mosher’s Analysis of Ramariolide A76 ................................................................. 39	  2.6.4.1	   Synthesis of MTPA Esters of Ramariolide A ................................................ 39	  2.6.5	   2D NMR Spectra of Ramariolide A–D ................................................................. 41	    vii Chapter 3: Dhilirolides A–N, Meroterpenoids Produced in Culture by the Fruit-Infecting Fungus Penicillium purpurogenum Collected in Sri Lanka ..................................................... 48	  3.1	   Chemical Prospecting: Structure Guided Isolation of Natural Products ..................... 48	  3.2	   Global Fungal Diversity ............................................................................................... 48	  3.2.1	   Fungal Diversity in Sri Lanka ............................................................................... 49	  3.3	   The Relationship Between Fungi and Plants ............................................................... 49	  3.4	   Diversity of Fungal Secondary Metabolites: The C25 Meroterpenoids ....................... 50	  3.4.1	   Diversity and Biosynthetic Origins of C25 Meroterpenoids From 3,5-Dimethylorsellinic acid ..................................................................................................... 50	  3.4.2	   Biological Activity ................................................................................................ 53	  3.5	   Isolation of Dhilirolides A–N ...................................................................................... 54	  3.6	   Structure Elucidation of Dhilirolides A–N .................................................................. 56	  3.6.1	   Structure Elucidation of Dhilirolide A .................................................................. 56	  3.6.2	   Structure Elucidation of Dhilirolide B .................................................................. 62	  3.6.3	   Structure Elucidation of Dhilirolide C .................................................................. 63	  3.6.4	   Structure Elucidation of Dhilirolide D .................................................................. 66	  3.6.5	   Structure Elucidation of Dhilirolide E .................................................................. 70	  3.6.6	   Structure Elucidation of Dhilirolide F .................................................................. 73	  3.6.7	   Structure Elucidation of Dhilirolide G .................................................................. 75	  3.6.8	   Structure Elucidation of Dhilirolide H .................................................................. 78	  3.6.9	   Structure Elucidation of Dhilirolide I ................................................................... 81	  3.6.10	   Structure Elucidation of Dhilirolide J ................................................................. 85	  3.6.11	   Structure Elucidation of Dhilirolide K ................................................................ 86	    viii 3.6.12	   Structure Elucidation of Dhilirolide L ................................................................ 89	  3.6.13	   Structure Elucidation of Dhilirolide M ............................................................... 91	  3.6.14	   Structure Elucidation of Dhilirolide N ................................................................ 93	  3.7	   Dhilirolide Feeding Studies and Biosynthetic Proposal .............................................. 95	  3.8	   Dhilirolide Biological Activity .................................................................................... 98	  3.9	   Conclusion ................................................................................................................... 98	  3.10	   General Experimental Procedures .............................................................................. 99	  3.10.1	   Fungal Material ................................................................................................... 99	  3.10.2	   Extraction of P. purpurogenum and Isolation of Dhilirolides A–N ................... 99	  3.10.3	   Stable Isotope Feeding Study ............................................................................ 101	  3.11	   2D NMR Spectra for the Dhilirolides A–N ............................................................. 102	  Chapter 4: Xestolactone, Xestosaprol O and Known Xestoquinones Identified as Potent Indolamine 2, 3-Dioxygenase (IDO) Inhibitors ...................................................................... 125	  4.1	   IDO ............................................................................................................................ 125	  4.1.1	   IDO in Disease and Immune Evasion ................................................................. 127	  4.1.2	   IDO Inhibitors ..................................................................................................... 128	  4.2	   Xestoquinones ............................................................................................................ 130	  4.2.1	   Xestoquinone Biogenesis .................................................................................... 131	  4.2.2	   Known Xestoquinone Biological Activity .......................................................... 133	  4.3	   Isolation of Xestolactone A, Xestosaprol O and P .................................................... 134	  4.4	   Structure Elucidation of Novel Xestoquinone Analogues ......................................... 135	  4.4.1	   Structure Elucidation of Xestolactone A ............................................................ 135	  4.4.2	   Structure Elucidation of Xestosaprol O .............................................................. 142	    ix 4.4.3	   Structure Elucidation of Xestosaprol P ............................................................... 145	  4.5	   Proposed Biosynthetic Origin of Xestolactone .......................................................... 149	  4.6	   Structure Revision of Adociaquinone A and B Based on Isolated Material .............. 150	  4.7	   The Xestoquinones as Inhibitors of IDO ................................................................... 153	  4.8	   Conclusion ................................................................................................................. 154	  4.9	   Experimental Section ................................................................................................. 155	  4.9.1	   IDO Inhibition Assays ........................................................................................ 155	  4.9.2	   General Experimental Procedures ....................................................................... 155	  4.9.3	   Extraction and Isolation ...................................................................................... 156	  4.10	   2D NMR Spectra ...................................................................................................... 157	  Chapter 5: Synthesis of Analogues of Xestosaprol N and the Adociaquinones; Structure Activity Relationship Studies and Novel Inhibitors of Indolamine 2,3-Dioxygenase. ........... 161	  5.1	   Previous Syntheses of Xestoquinone and Analogues ................................................ 161	  5.2	   Synthesis of Xestosaprol N Analogues ...................................................................... 162	  5.2.1	   Photocyclization Towards the Carbon Framework of Xestosaprol O ................ 162	  5.2.2	   Retrosynthetic Analysis ...................................................................................... 163	  5.2.3	   Initial Synthetic Trials ......................................................................................... 163	  5.2.4	   Halogen Effects on Photocyclization .................................................................. 165	  5.2.5	   Synthetic Efforts Continued ................................................................................ 166	  5.3	   Further Synthesis of Xestoquinone Analogues .......................................................... 169	  5.3.1	   Attempted Synthesis of 5.28 and 5.29 ................................................................ 170	  5.3.2	   Attempted Synthesis of 5.30 ............................................................................... 171	  5.3.3	   Thiazine Containing Analogues .......................................................................... 172	    x 5.4	   Biological Activity of Adociaquinone and Xestosaprol O (4.20) Derivatives .......... 173	  5.5	   Conclusion ................................................................................................................. 174	  5.6	   Experimental Section ................................................................................................. 176	  5.6.1	   General Experimental Parameters for the IDO Assay ........................................ 176	  5.6.2	   General Experimental Procedures ....................................................................... 176	  5.6.3	   Synthetic Procedures and Methodology ............................................................. 177	  Chapter 6: Chemical Probe Synthesis, Cellular Target Identification, and Structure Activity Relationship Studies of Latonduine A: A Trafficking Corrector of the Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) for the Treatment of Cystic Fibrosis. ....... 235	  6.1	   Cystic Fibrosis ........................................................................................................... 235	  6.1.1	   The CFTR Protein ............................................................................................... 235	  6.2	   Cystic Fibrosis Treatments ........................................................................................ 237	  6.2.1	   Current Clinical Treatments ................................................................................ 237	  6.2.2	   CFTR Trafficking Correctors ............................................................................. 238	  6.3	   Elucidation of the Enzymatic Target of Latonduine A for the Correction of F508del-CFTR   ................................................................................................................................ 240	  6.3.1	   Construction of Latonduine A ABPP Probes...................................................... 243	  6.4	   PARP Inhibitors As Trafficking Correctors of F508del-CFTR ................................. 247	  6.4.1	   The PARP Family of Enzymes ........................................................................... 247	  6.4.2	   PARP/ ARTD-3 .................................................................................................. 248	  6.4.3	   PARP 16 and Other PARP Family Members ..................................................... 249	  6.4.4	   Synthetic PARP Inhibitors .................................................................................. 250	  6.5	   Synthesis of Novel Trafficking Correctors of F508del-CFTR .................................. 251	    xi 6.5.1	   Pyrrole Azepine SAR .......................................................................................... 251	  6.5.2	   Benzazepinone Trafficking Corrector Synthesis ................................................ 252	  6.5.3	   Ethyl Silyl Tethered Phthalimides Produce Dihydrobenzazepinone .................. 253	  6.5.4	   Phthalimide Photoreactions with Styrene Derivatives ........................................ 254	  6.5.5	   Other Alkene Phthalimide Photoreactions .......................................................... 255	  6.5.6	   Schmidt Rearrangement ...................................................................................... 256	  6.5.7	   First Generation Correctors with Combined Functionality ................................ 256	  6.6	   Combining Selective PARP Inhibitors ...................................................................... 258	  6.7	   Assay Results ............................................................................................................. 259	  6.8	   Conclusions ................................................................................................................ 259	  6.9	   Experimental .............................................................................................................. 260	  6.9.1	   General Experimental ......................................................................................... 260	  Chapter 7: Conclusion ............................................................................................................. 345	  References ...................................................................................................................................348	  Appendices ..................................................................................................................................364	             xii List of Tables Table 2.1 NMR data for ramariolide A (2.18) and ramariolide B (2.19). .................................... 28	  Table 2.2 NMR Data (C6D6, 1H 600 MHz, 13C 150 MHz) for ramariolides C (2.20) and D (2.21)........................................................................................................................................................ 33	  Table 3.1 1H and 13C NMR chemical shifts for dhilirolides A–D. ............................................... 57	  Table 3.2 1H NMR data for dhilirolides E–I (3.13–3.17) recorded in DMSO-d6. ........................ 71	  Table 3.3 13C NMR data for dhilirolides E–G (3.13–3.22) recorded in DMSO-d6. ..................... 72	  Table 3.4 1H NMR data for dhilirolides J–N (3.18–3.22) recorded in DMSO-d6. ....................... 85	  Table 4.1 1H and 13C NMR chemical shifts for 4.19, 4.20 and 4.21. ......................................... 139	  Table 6.1 Conditions screened for Heck coupling using 6.14 or 6.15 as starting materials. ...... 245	  Table 6.2 Photoreaction products and reduction products .......................................................... 254	  Table 6.3 Schmidt reaction products and isolated yields. ........................................................... 256	  Table 6.4 delF508-CFTR trafficking correction and PARP inhibition. ...................................... 259	  	      xiii List of Figures  Figure 1.1 Morphine (1.1) and salicylic acid (1.3) and their acetyl derivatives ............................. 2	  Figure 1.2 Penicillin G (1.4) and amoxicillin (1.6) ......................................................................... 3	  Figure 1.3 Spongothymidine (1.7) with Ara-C (1.8) and -A (1.9) .................................................. 3	  Figure 1.4 Plant and sponge derived natural products likely produced by microorganisms21–23 ... 5	  Figure 1.5 Type II PKS product tetracycline (1.15) ....................................................................... 6	  Figure 1.6 Type I PKS biosynthetic route for monensin (1.16).25 .................................................. 7	  Figure 1.7 Terpene biosynthesis25 ................................................................................................... 8	  Figure 1.8 Shikimic acid formation and examples of economically important shikimate products25 ........................................................................................................................................ 9	  Figure 1.9 RP and NRP products Prialt® (1.25) and dolastatin 10 (1.26).27–29 ............................. 10	  Figure 1.10 Lysergic acid (1.27), quinine (1.28) and staurosporine (1.29) .................................. 11	  Figure 1.11 Oroidin proposed biosynthetic pathway based on labeling studies33, 34 .................... 12	  Figure 1.12 Rapamycin (1.35) and trapoxin (1.36)35, 37 ................................................................ 15	  Figure 2.1 Streptomycin (2.1) and the current first-line antibiotics 39, 42 ...................................... 17	  Figure 2.2 A diagram of the cell capsule’s underlying structure in Mtb and an example of a mycolic acid. 46, 47 .......................................................................................................................... 19	  Figure 2.3 Several drugs used to inhibit Mtb cell wall biosynthesis or mycolic acid production42....................................................................................................................................................... 19	  Figure 2.4 Mtb drugs with mechanisms of action other than inhibition of cell wall biosynthesis.39, 60 .................................................................................................................................................... 22	  Figure 2.5 Clinically relevant mushroom natural products61–66 .................................................... 23	  Figure 2.6 Ramaria sp. in situ. ...................................................................................................... 23	  Figure 2.7 Structures of ramariolides A–D. .................................................................................. 24	  Figure 2.8 1H NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in methylene chloride-d2. .................................................................................................................................... 25	  Figure 2.9 13C NMR spectrum of ramariolide A (2.18) recorded at 150 MHz in methylene chloride-d2. .................................................................................................................................... 26	    xiv Figure 2.10 Selected 2D NMR correlations for ramariolide A (2.18). Fragments A, B and C were constructed using gHMBC and gCOSY data. ............................................................................... 26	  Figure 2.11 ORTEP diagram for ramariolide A (2.18). ................................................................ 27	  Figure 2.12 Relevant COSY and HMBC correlations for ramariolide B (2.19). ......................... 29	  Figure 2.131H NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. ..... 29	  Figure 2.14 13C NMR spectrum of ramariolide B (2.19) recorded at 150 MHz in benzene-d6 .... 30	  Figure 2.15 1D NOESY NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. .................................................................................................................................... 31	  Figure 2.16 1H NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6. .... 32	  Figure 2.17 13C NMR spectrum of ramariolide C (2.20) recorded at 150 MHz in benzene-d6. ... 32	  Figure 2.18 1H NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6. .... 34	  Figure 2.19 13C NMR spectrum of ramariolide D (2.21) recorded at 150 MHz in benzene-d6. ... 34	  Figure 2.20 Anti-mycobacterial activity of ramariolide A (2.18). ................................................ 36	  Figure 2.21 Hygrophorone F (2.22) and G (2.23) and parthexetine (2.24).72–74 ........................... 37	  Figure 2.22 Mosher’s Analysis: Showing Δδ(δS-δR) values from Mosher’s ester analysis of ramariolide A76 (2.18). .................................................................................................................. 39	  Figure 2.23 1H NMR spectrum of ramariolide A MTPA-R ester (2.25) recorded at 600 MHz in CDCl3 ............................................................................................................................................ 40	  Figure 2.24 1H NMR spectrum of ramariolide A MTPA-S (2.26) ester recorded at 600 MHz in CDCl3. ........................................................................................................................................... 41	  Figure 2.25 gCOSY NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in CD2Cl2. . 41	  Figure 2.26 gHSQC NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in CD2Cl2. . 42	  Figure 2.27 gHMBC NMR spectrum of ramariolide A(2.18) recorded at 600 MHz in CD2Cl2. . 42	  Figure 2.28 DEPT 135 NMR spectrum of ramariolide A (2.18) recorded at 150 MHz in CD2Cl2........................................................................................................................................................ 43	  Figure 2.29 gCOSY NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6........................................................................................................................................................ 43	  Figure 2.30 gHSQC NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6........................................................................................................................................................ 44	    xv Figure 2.31 gHMBC NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. .................................................................................................................................................. 44	  Figure 2.32 gCOSY NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6........................................................................................................................................................ 45	  Figure 2.33 gHSQC NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6........................................................................................................................................................ 45	  Figure 2.34 gHMBC NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6. .................................................................................................................................................. 46	  Figure 2.35 gCOSY NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6........................................................................................................................................................ 46	  Figure 2.36 gHSQC NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6........................................................................................................................................................ 47	  Figure 2.37 gHMBC NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6. .................................................................................................................................................. 47	  Figure 3.1 Peramine and ergotamine alkaloids and meroterpenoid clinical candidates. .............. 50	  Figure 3.2 Representatives of each novel fungal C25 meroterpenoid carbon skeleton.103, 104, 106–119....................................................................................................................................................... 52	  Figure 3.3 Biosynthetic route overview to C25 meroterpenoidsAdapted from 103, 123–126 ..................... 53	  Figure 3.4 The dhilirolides A–N (3.9–3.22) and the dhilirane (I), isodhilirane (II), 14,15-dinordhilirane (III), and 23,24-dinorisodhilirane (IV) meroterpenoid carbon skeletons. ............. 55	  Figure 3.5 1H NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6 showing numbering. .................................................................................................................................... 58	  Figure 3.6 13C NMR spectrum of dhilirolide A (3.9) recorded at 150 MHz in DMSO-d6. .......... 58	  Figure 3.7 Selected gHMBC and gCOSY 60 correlations for dhilirolide A (3.9). ....................... 59	  Figure 3.8 gHMBC spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6. ............ 60	  Figure 3.9 gCOSY 60 NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6........................................................................................................................................................ 60	  Figure 3.10 Selected tROESY correlations of dhilirolide A (3.9) ................................................ 61	  Figure 3.11 tROESY NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6. 61	  Figure 3.12 ORTEP diagram of dhilirolide A (3.9) ...................................................................... 62	    xvi Figure 3.13 1H NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6. ....... 63	  Figure 3.14 13C NMR spectrum of dhilirolide B (3.10) recorded at 150 MHz in DMSO-d6. ...... 63	  Figure 3.15 1H NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4. ........ 65	  Figure 3.16 13C NMR spectrum of dhilirolide C (3.11) recorded at 150 MHz in MeOD-d4 ........ 65	  Figure 3.17 1H NMR spectrum of dhilirolide D (3.12) recorded at 600 MHz in DMSO-d6. ....... 66	  Figure 3.18 13C NMR spectrum of dhilirolide D (3.12) recorded at 150 MHz in DMSO-d6 ....... 67	  Figure 3.19 Selected gHMBC and gCOSY correlations of dhilirolide D (3.12) .......................... 68	  Figure 3.20 gCOSY spectrum of dhilirolide D (3.12) in DMSO-d6. ............................................ 68	  Figure 3.21 gHMBC spectrum of dhilirolide D (3.12) in DMSO-d6. ........................................... 69	  Figure 3.22 Selected tROESY correlations for dhilirolide D (3.12) ............................................. 69	  Figure 3.23 tROESY spectrum of dhilirolide D (3.12) recorded in DMSO-d6 ............................ 70	  Figure 3.24 1H NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6 ......... 72	  Figure 3.25 13C NMR spectrum of dhilirolide E (3.13) recorded at 150 MHz in DMSO-d6 ....... 73	  Figure 3.26 1H NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6 ......... 74	  Figure 3.27 13C NMR spectrum of dhilirolide F (3.14) recorded at 150 MHz in DMSO-d6. ....... 75	  Figure 3.28 1H NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6 ........ 76	  Figure 3.29 13C NMR spectrum of dhilirolide G (3.15) recorded at 150 MHz in DMSO-d6 ....... 76	  Figure 3.30 tROESY NMR spectrum and expansions of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6 ......................................................................................................................... 77	  Figure 3.31 1H NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6 ........ 79	  Figure 3.32 13C NMR spectrum of dhilirolide H (3.16) recorded at 150 MHz in DMSO-d6 ....... 79	  Figure 3.33 tROESY NMR spectrum expansions of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6 ...................................................................................................................................... 80	  Figure 3.34 tROESY NMR spectrum expansions of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6 ...................................................................................................................................... 81	  Figure 3.35 1H NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 .......... 82	  Figure 3.36 13C NMR spectrum of dhilirolide I (3.17) recorded at 150 MHz in DMSO-d6 ......... 82	  Figure 3.37 gHMBC NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 83	  Figure 3.38 gHMBC NMR spectrum expansions of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 ...................................................................................................................................... 83	    xvii Figure 3.39 tROESY NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 84	  Figure 3.40 tROESY NMR spectrum expansions of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 ...................................................................................................................................... 84	  Figure 3.41 1H NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6. ........ 86	  Figure 3.42 13C NMR spectrum of dhilirolide J (3.18) recorded at 150 MHz in DMSO-d6. ....... 86	  Figure 3.43 1H NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6 ........ 88	  Figure 3.44 13C NMR spectrum of dhilirolide K (3.19) recorded at 150 MHz in DMSO-d6 ....... 88	  Figure 3.45 1H NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6 ......... 89	  Figure 3.46 ORTEP diagram for dhilirolide L (3.20). .................................................................. 90	  Figure 3.47 13C NMR spectrum of dhilirolide L (3.20) recorded at 150 MHz in DMSO-d6 ....... 90	  Figure 3.48 1H NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6 ....... 92	  Figure 3.49 13C NMR spectrum of dhilirolide M (3.21) recorded at 150 MHz in DMSO-d6 ...... 92	  Figure 3.50 1H NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6. ....... 94	  Figure 3.51 13C NMR spectrum of dhilirolide N (3.22) recorded at 150 MHz in DMSO-d6 ....... 94	  Figure 3.52 2D-INADEQUATE spectrum of dhilirolide A (3.9), obtained from a 13C doubly labeled acetate feeding experiment, recorded at 150 MHz in DMSO-d6. .................................... 96	  Figure 3.53 gHSQC NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6 102	  Figure 3.54 gCOSY60 NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 103	  Figure 3.55 gHSQC NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 103	  Figure 3.56 gHMBC NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 104	  Figure 3.57 tROESY NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 104	  Figure 3.58 gCOSY60 NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4..................................................................................................................................................... 105	  Figure 3.59 gHSQC NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4..................................................................................................................................................... 105	    xviii Figure 3.60 gHMBC NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4..................................................................................................................................................... 106	  Figure 3.61 tROESY NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4..................................................................................................................................................... 106	  Figure 3.62 gHSQC NMR spectrum of dhilirolide D (3.12) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 107	  Figure 3.63 gCOSY 60 NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 107	  Figure 3.64 gHSQC NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 108	  Figure 3.65 gHMBC NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 108	  Figure 3.66 tROESY NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 109	  Figure 3.67 gCOSY 60 NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 109	  Figure 3.68 gHSQC NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 110	  Figure 3.69 gHMBC NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 110	  Figure 3.70 tROESY NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 111	  Figure 3.71 gCOSY 60 NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 111	  Figure 3.72 gHSQC NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 112	  Figure 3.73 gHMBC NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 112	  Figure 3.74 gCOSY 60 NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 113	    xix Figure 3.75 gHSQC NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 113	  Figure 3.76 gHMBC NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 114	  Figure 3.77 gCOSY 60 NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 114	  Figure 3.78 gHSQC NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 115	  Figure 3.79 gCOSY 60 NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 115	  Figure 3.80 gHSQC NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6 116	  Figure 3.81 gHMBC NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 116	  Figure 3.82 gCOSY 60 NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 117	  Figure 3.83 gHSQC NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 117	  Figure 3.84 gHMBC NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 118	  Figure 3.85 tROESY NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 118	  Figure 3.86 gCOSY 60 NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 119	  Figure 3.87 gHSQC NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 119	  Figure 3.88 gHMBC NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 120	  Figure 3.89 tROESY NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 120	  Figure 3.90 gCOSY 60 NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6 ................................................................................................................................................. 121	    xx Figure 3.91 gHSQC NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 121	  Figure 3.92 gHMBC NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 122	  Figure 3.93 tROESY NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 122	  Figure 3.94 gCOSY 60 NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 123	  Figure 3.95 gHSQC NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 123	  Figure 3.96 gHMBC NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 124	  Figure 3.97 tROESY NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 124	  Figure 4.1 Two plausible mechanisms for indole cleavage by IDO.143, 144 ................................ 125	  Figure 4.2 Tryptophan metabolites and catabolic pathway 141, 146 .............................................. 126	  Figure 4.3 Marine natural product inhibitors of IDO and synthetic analogues .......................... 129	  Figure 4.4 All known carbon skeletons, represented by a known natural product, related to the xestoquinone family of compounds.159–163, 165–167 ....................................................................... 131	  Figure 4.5 Proposed biogenesis159, 163, 168–170 .............................................................................. 132	  Figure 4.6 Fungal produced related compounds.171, 172, 174, 175 .................................................... 133	  Figure 4.7 Xestoquinones isolated from X. vansoestii160, 162, 183, 184 ............................................ 135	  Figure 4.8 1H NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6. ... 136	  Figure 4.9 13C NMR spectrum of xestolactone A (4.19) recorded at 150 MHz in acetone-d6. .. 136	  Figure 4.10 Selected HMBC and COSY correlations for xestolactone A (4.19). ...................... 138	  Figure 4.11 Inset of gCOSY NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6. ................................................................................................................................... 140	  Figure 4.12 Insets of olefinic region of the gHMBC NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in DMSO-d6. ............................................................................................ 140	    xxi Figure 4.13 Insets of aliphatic region of the gHMBC NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in DMSO-d6. ............................................................................................ 141	  Figure 4.14 ORTEP diagram of 4.19. ......................................................................................... 142	  Figure 4.15 Structure of xestosaprol O (4.20) showing selected HMBC and COSY correlations..................................................................................................................................................... 143	  Figure 4.16 1H NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6. .... 143	  Figure 4.17 13C NMR spectrum of xestosaprol O (4.20) recorded at 150 MHz in DMSO-d6. .. 144	  Figure 4.18 gCOSY 60 spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6. 144	  Figure 4.19 gHMBC spectrum expansions of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6. ................................................................................................................................... 145	  Figure 4.20 Xestosaprol P (4.21) showing selected gHMBC and gCOSY correlations ............. 146	  Figure 4.21 The 1H NMR spectrum of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6...................................................................................................................................................... 146	  Figure 4.22 The 13C NMR spectrum of xestosaprol P (4.21) recorded at 150 MHz in DMSO-d6...................................................................................................................................................... 147	  Figure 4.23 gHMBC spectrum expansion of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. ................................................................................................................................... 147	  Figure 4.24 gHMBC spectrum expansion of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. ................................................................................................................................... 148	  Figure 4.25 gCOSY spectrum expansion of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. ................................................................................................................................................ 148	  Figure 4.26 ORTEP diagram of 3,13-dideoxo-1, 2,14,15-tetrahydro-3, 13-dihydrohalenaquinone (4.23)162 ....................................................................................................................................... 149	  Figure 4.27 Proposed biogenesis for xestolactone (4.19) carbon skeleton. ................................ 150	  Figure 4.28 1H NMR spectrum of adociaquinone A (4.11) recorded at 600 MHz in DMSO-d6...................................................................................................................................................... 151	  Figure 4.29 13C NMR spectrum of adociaquinone A (4.11) recorded at 150 MHz in DMSO-d6..................................................................................................................................................... 151	  Figure 4.30 Structure assignment of isolated 4.11 and the reported 13C NMR chemical shifts of 4.11 and 4.12.162 .......................................................................................................................... 152	    xxii Figure 4.31 gHMBC NMR spectrum of adociaquinone A (4.11) recorded at 600 MHz in DMSO-d6 ................................................................................................................................................. 153	  Figure 4.32 Nonlinear regression curves were based on a sigmoidal dose-response equation and used to calculate the corresponding IC50 values shown in the table. The data are shown as an average of quadruplicates with error bars representing SD. ....................................................... 154	  Figure 4.33 gHSQC NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6...................................................................................................................................................... 157	  Figure 4.34 tROESY NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6 ................................................................................................................................................. 158	  Figure 4.35 gHSQC NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 158	  Figure 4.36 gHMBC NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 159	  Figure 4.37 tROESY NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6..................................................................................................................................................... 159	  Figure 4.38 gHSQC spectrum of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6 ....... 160	  Figure 4.39 tROESY spectrum of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. ..... 160	  Figure 5.1 Several previously synthesized members of the xestoquinone family of compounds.162, 164, 189–195 ............................................................................................................. 161	  Figure 5.2 R. Caralini synthesis of (±)-xestoquinone.195 ............................................................ 162	  Figure 5.3 Xestosaprol O (4.20) and synthetic target (5.1) ........................................................ 162	  Figure 5.4 Silver and light promoted cyclization reported by Sato and Tamura.199, 200 .............. 163	  Figure 5.5 Retrosynthetic analysis .............................................................................................. 163	  Figure 5.6 Representative over oxidized side products .............................................................. 168	  Figure 5.7 Proposed abbreviated synthetic targets and tetrahydroxestoquinol (4.22) ................ 170	  Figure 5.8 Diosphenol and diketone abitane diterpenoids .......................................................... 170	  Figure 5.9 Comparison of the effect of 5.24 (filled circles) and 5.23 (hollow circles) on the enzymatic .................................................................................................................................... 173	  Figure 5.10 IC50 values for synthetic xestoquinone analogues with SAR diagram of xestosaprol O colored to reflect findings. Black bonds/ atoms represent the minimal pharmacophore; green   xxiii bonds/ atoms are neutral additions or untested; and blue bonds/ atoms appear deleterious for hIDO inhibition. .......................................................................................................................... 174	  Figure 5.11 Analogues of the xestoquinones tested. ................................................................... 175	  Figure 5.12 1H and 13C NMR spectra of 5.11 in acetone-d6 at 600 MHz and 150 MHz ............ 178	  Figure 5.13 1H and 13C NMR spectra of 5.12 in MeOD-d4 at 600 MHz and 150 MHz ............. 179	  Figure 5.14 1H and 13C NMR spectra of 5.15 in CDCl3 at 600 MHz and 150 MHz .................. 181	  Figure 5.15 1H and 13C NMR spectra of 5.16 in CDCl3 at 600 MHz and 150 MHz .................. 183	  Figure 5.16 1H and 13C NMR spectra of 5.17A in CDCl3 at 600 MHz and 150 MHz ............... 185	  Figure 5.17 1H and 13C NMR spectra of 5.17B in acetone-d6 at 600 MHz and 150 MHz ......... 187	  Figure 5.18 1H and 13C NMR spectra of 5.17C in CDCl3 at 600 MHz and 150 MHz ............... 189	   Figure 5.19 1H and 13C NMR spectra of 5.18 in CD2Cl2 at 600 MHz and 150 MHz ................ 191	  Figure 5.20 1H and 13C NMR spectra of 5.20 in CDCl3 at 600 MHz and 150 MHz ................... 193	  Figure 5.21 1H and 13C NMR spectra of 5.19 in CDCl3 at 600 MHz and 150 MHz .................. 195	  Figure 5.22 1H and 13C NMR spectra of 5.22 in DMSO-d6 at 600 MHz and 150 MHz ............. 197	  Figure 5.23 1H and 13C NMR spectra of 5.26 in CDCl3 at 600 MHz and 150 MHz .................. 199	  Figure 5.24 1H and 13C NMR spectra of 5.27 in DMSO-d6 at 600 MHz and 150 MHz ............. 201	  Figure 5.25 gHMBC and gHSQC NMR spectra of 5.27 in DMSO-d6 at 600 MHz ................... 202	  Figure 5.26 1H and 13C NMR spectra of 5.21 in DMSO-d6 at 600 MHz and 150 MHz ............. 204	  Figure 5.27 1H and 13C NMR spectra of 5.5 in DMSO-d6 at 600 MHz and 150 MHz ............... 206	  Figure 5.28 1H and 13C NMR spectra of 5.23 in DMSO-d6 at 600 MHz and 150 MHz ............. 208	  Figure 5.29 1H and 13C NMR spectra of 5.24 in DMSO-d6 at 600 MHz and 150 MHz ............. 210	  Figure 5.30 1H and 13C NMR spectra of 5.29 in CD2Cl2 at 600 MHz and 150 MHz ................. 213	  Figure 5.31 1H and 13C NMR spectra of 5.40 in acetone-d6 at 600 MHz and 150 MHz ............ 215	  Figure 5.32 gHMBC NMR spectrum of 5.40 in acetone-d6 at 600 MHz ................................... 216	  Figure 5.33 1H and 13C NMR spectra of 5.38 in CD2Cl2 at 600 MHz and 150 MHz ................. 217	  Figure 5.34 1H and 13C NMR spectra of 5.39 in CDCl3 at 600 MHz and 150 MHz .................. 219	  Figure 5.35 1H and 13C NMR spectra of 5.42 in acetone-d6 at 600 MHz and 150 MHz ............ 221	  Figure 5.36 1H and 13C NMR spectra of 5.43 in acetone-d6 at 600 MHz and 150 MHz ............ 223	  Figure 5.37 1H NMR spectrum in CDCl3 and 13C NMR spectrum in DMSO-d6 of 5.14 at 600 MHz and 150 MHz ..................................................................................................................... 226	    xxiv Figure 5.38 1H and gHSQC NMR spectra of 5.45 in DMSO-d6 at 600 MHz ............................ 228	  Figure 5.39 gHMBC NMR spectrum of 5.45 in DMSO-d6 at 600 MHz .................................... 229	  Figure 5.40 1H and 13C NMR spectra of 5.44 in DMSO-d6 at 600 MHz and 150 MHz ............. 230	  Figure 5.41 1H and 13C NMR spectra of 5.47 in DMSO-d6 at 600 MHz and 150 MHz ............. 232	  Figure 5.42 1H NMR spectrum of 5.32/ 5.33 mixture in DMSO-d6 at 600 MHz ....................... 234	  Figure 6.1 Trafficking of WT and mutant CFTR in the cell.217–223 ............................................ 237	  Figure 6.2 Various treatments of CF in the clinical or preclinical stage of development219, 223 . 240	  Figure 6.3 General conceptual design of an ABPP tool and “Pull Down” experiments to separate an enzyme linked product.229, 231 ................................................................................................. 242	  Figure 6.4 Latonduine scaffold SAR .......................................................................................... 243	  Figure 6.5 Assay for trafficking correction of latonduine photo-affinity probe (6.23) .............. 246	  Figure 6.6 ABT-88879 ................................................................................................................. 247	  Figure 6.7 Mechanism of ADP-ribose attachment to protein substrate ...................................... 248	  Figure 6.8 Latonduine bromopyrrole azepinone and benzaepinone for comparison78, 234 .......... 250	  Figure 6.9 Alcohol 6.27 and olefin 6.11 ..................................................................................... 251	  Figure 6.10 General reaction sequence for styrene derivatives tested .243 .................................. 254	  Figure 6.11 General formula for side product ............................................................................ 255	  Figure 6.12 Generalized formula for the Schmidt reaction ........................................................ 256	  Figure 6.13 Trafficking correction data for 6.56 and 6.57 .......................................................... 257	  Figure 6.14 PARP inhibition data for 6.56 and 6.57 .................................................................. 258	  Figure 6.15 1H and 13C NMR spectra of 6.18 in DMSO-d6 at 600 and 150 MHz ...................... 262	  Figure 6.16 1H and 13C NMR spectra of 6.19 in DMSO-d6 at 600 and 150 MHz. ..................... 264	  Figure 6.17 1H and 13C NMR spectra of 6.14 in acetone-d6 at 600 and 150 MHz. .................... 266	  Figure 6.18 1H and 13C NMR spectra of 6.15 in acetone-d6 at 600 and 150 MHz ..................... 268	  Figure 6.19 1H and 13C NMR spectra of 6.22 in DMSO-d6 at 600 and 150 MHz. ..................... 270	  Figure 6.20 1H and 13C NMR spectra of 6.20 in DMSO-d6 at 600 and 150 MHz. ..................... 272	  Figure 6.21 1H and 13C NMR spectra of 6.23 in DMSO-d6 at 600 and 150 MHz. ..................... 274	  Figure 6.22 1H and 13C NMR spectra of 6.27 in DMSO-d6 at 600 and 150 MHz. ..................... 276	  Figure 6.23 1H and 13C NMR spectra of 6.30 in DMSO-d6 at 600 and 150 MHz. ..................... 278	  Figure 6.24 1H and 13C NMR spectra of 6.29 in DMSO-d6 at 600 and 150 MHz. ..................... 280	    xxv Figure 6.25 1H and 13C NMR spectra of 6.31 in DMSO-d6 at 600 and 150 MHz. ..................... 282	  Figure 6.26 1H and 13C NMR spectra of 6.32 in DMSO-d6 at 600 and 150 MHz. ..................... 284	  Figure 6.27 1H and 13C NMR spectra of 6.35 in DMSO-d6 at 600 and 150 MHz. ..................... 287	  Figure 6.28 1H and 13C NMR spectra of 6.36 in CDCl3 at 600 and 150 MHz. .......................... 289	  Figure 6.29 1H and 13C NMR spectra of 6.37 in DMSO-d6 at 600 and 150 MHz. ..................... 291	  Figure 6.30 1H and 13C NMR spectra of 6.38 in DMSO-d6 at 600 and 150 MHz. ..................... 293	  Figure 6.31 1H and 13C NMR spectra of 6.40 in DMSO-d6 at 600 and 150 MHz. ..................... 296	  Figure 6.32 1H and 13C NMR spectra of 6.41 in DMSO-d6 at 600 and 150 MHz. ..................... 298	  Figure 6.33 1H and 13C NMR spectra of 6.42 in DMSO-d6 at 600 and 150 MHz. ..................... 300	  Figure 6.34 1H and 13C NMR spectra of 6.43 in DMSO-d6 at 600 and 150 MHz. ..................... 302	  Figure 6.35 1H and 13C NMR spectra of 6.44 in DMSO-d6 at 600 and 150 MHz. ..................... 304	  Figure 6.36 1H and 13C NMR spectra of 6.45 in MeOD-d4 at 600 and 150 MHz. ..................... 306	  Figure 6.37 1H and 13C NMR spectra of 6.46 in acetone-d6 at 600 and 150 MHz. .................... 308	  Figure 6.38 1H and 13C NMR spectra of 6.47 in DMSO-d6 at 600 and 150 MHz. ..................... 310	  Figure 6.39 1H and 13C NMR spectra of 6.48 t in DMSO-d6 at 600 and 150 MHz. ................... 312	  Figure 6.40 tROESY NMR spectrum of 6.48 t in DMSO-d6 at 600 MHz ................................. 313	  Figure 6.41 1H and 13C NMR spectra of 6.48 c in DMSO-d6 at 600 and 150 MHz. .................. 314	  Figure 6.42 tROESY NMR spectra of 6.48 c in DMSO-d6 at 600 MHz .................................... 315	  Figure 6.43 1H and 13C NMR spectra of 6.50A in acetone-d6 at 600 and 150 MHz. ................. 318	  Figure 6.44 1H and 13C NMR spectra of 6.50C in acetone-d6 at 600 and 150 MHz. ................. 320	  Figure 6.45 1H and 13C NMR spectra of 6.51B in DMSO-d6 at 600 and 150 MHz. .................. 322	  Figure 6.46 1H and 13C NMR spectra of 6.51C in DMSO-d6 at 600 and 150 MHz. .................. 324	  Figure 6.47 1H and 13C NMR spectra of 6.52A in DMSO-d6 at 600 and 150 MHz. .................. 326	  Figure 6.48 1H and 13C NMR spectra of 6.52B in DMSO-d6 at 600 and 150 MHz. .................. 328	  Figure 6.49 1H and 13C NMR spectra of 6.52C in DMSO-d6 at 600 and 150 MHz. .................. 330	  Figure 6.50 1H and 13C NMR spectra of 6.53A in acetone-d6 at 600 and 150 MHz. ................. 332	  Figure 6.51 1H and 13C NMR spectra of 6.53B in acetone-d6 at 600 and 150 MHz. ................. 334	  Figure 6.52 1H and 13C NMR spectra of 6.53C in acetone-d6 at 600 and 150 MHz. ................. 336	  Figure 6.53 1H and 13C NMR spectra of 6.54 in DMSO-d6 at 600 and 150 MHz. ..................... 338	  Figure 6.54 1H and 13C NMR spectra of 6.55 in DMSO-d6 at 600 and 150 MHz. ..................... 340	    xxvi Figure 6.55 1H and 13C NMR spectra of 6.56 in acetone-d6 at 600 and 150 MHz. .................... 342	  Figure 6.56 gHMBC NMR spectra of 6.56 in acetone-d6 at 600 MHz. ..................................... 343	  Figure 6.57 1H and 13C NMR spectra of 6.57 in acetone-d6 at 600 and 150 MHz. .................... 344	  Figure 7.1 Xestosaprol O derivatives 5.24 and 5.33 showing the preferred thiazine ring regiochemistry ............................................................................................................................. 346	  Figure 7.2 Chiral derivitization and resolution plan for 6.57. .................................................... 347	           xxvii List of Schemes  Scheme 3.1 Proposed biogenetic pathway for the dhilirolides showing representative examples of each of the new dhilirane, isodhilirane, 14, 15-dinordhilirane and 23, 24-dinorisodhilirane skeletons. The labeling pattern indicating intact acetate units shown for dhilirolide A (3.9) was determined from the [1,2-13C2]-acetate feeding study. ................................................................. 97	  Scheme 5.1 1, 4-dimethoxynaphthalene trials. ........................................................................... 164	  Scheme 5.2 Successful photocyclization reactions and non-productive naphthyl starting materials..................................................................................................................................................... 165	  Scheme 5.3 Halogen effects on photocyclization ....................................................................... 166	  Scheme 5.4 Oxidations to form quinone products ...................................................................... 167	  Scheme 5.5 Two proposed reaction pathways leading to 5.21 and 5.5 ...................................... 167	  Scheme 5.6 Thiazine formation. ................................................................................................. 168	  Scheme 5.7 Synthetic route overview ......................................................................................... 169	  Scheme 5.8 Synthetic efforts towards 5.28. ................................................................................ 171	  Scheme 5.9 Attempted synthesis of 5.30 .................................................................................... 172	  Scheme 5.10 Reaction sequence for proposed minimal pharmacophore analogues .................. 173	  Scheme 6.1 Latonduine A (6.9) and B (6.10) and the previous total synthetic route for latonduine A (6.9).78 ..................................................................................................................................... 241	  Scheme 6.2 Proposed Heck coupling to form latonduine A (6.9) and analogues. ..................... 244	  Scheme 6.3 Synthesis of 6.14 and 6.15 ...................................................................................... 245	  Scheme 6.4 Latonduine probe synthesis ..................................................................................... 246	  Scheme 6.5 Pyrrole azepinone regio-isomer synthesis ............................................................... 252	  Scheme 6.6 Phthalimide photochemistry243–245 .......................................................................... 253	  Scheme 6.7 Syntheses of 6.32–6.36 ............................................................................................ 253	  Scheme 6.8 Syntheses of 6.47, 6.48 c, and 6.48 t. ...................................................................... 255	  Scheme 6.9 Synthetic route to 6.54 and 6.55 .............................................................................. 257	  Scheme 6.10 Synthetic route to 6.56 and 6.57. ........................................................................... 257	     xxviii List of Abbreviations °  - degree °C      - degree Celsius 1D  - 1-dimensional 2D  - 2-dimensional λmax  - wavelength correlating to a local maximum in absorbance λ  - wavelength Δ  - unsaturation/ alkene [α]20D  - specific rotation at sodium D-line (598nm) recorded at 20 °C Å  - angstrom δ  - chemical shift in parts per million from tetramethyl silane µg  - micrograms µl  - microliter µM  - micromolar µg  - microgram µm  - micrometer ε  - extinction coefficient β  - beta (two positions away) α  - alpha (one position away) ABCN  - 1, 1’-azobis(cyclohexanecarbonitrile) ABPP  - Activity Based Protein Profiling Ac  - acetyl acetone-d6 - deuterated acetone ACN  - acetonitrile AcOH  - acetic acid Ala  - alanine AlCl3  - aluminum trichloride Arg  - L-argenine ARTD  - diphtheria toxin like mono or poly ADP-ribosyltransferase enzyme Asp  - aspartate ATP  - adenosine triphosphate ax  - axial b  - broad B.C.  - Before Christ Benzene-d6 - deuterated benzene BOC  - tertiary-butyl carbonate BRSM  - based on recovered starting material nBu  - normal butyl tBu  - tertiary butyl tBuOH  - tertiary butanol c  - concentration 13C  - carbon isotope with 7 neutrons calcd.  - calculated CCl4  - carbon tetrachloride CD  - circular dichroism   xxix CDCl3  - deuterated chloroform CD2Cl2 - deuterated methylene chloride CFTR  - cystic fibrosis transmembrane conductance regulator CHO  - aldehyde CF  - cystic fibrosis cm  - centimeter CoA  - coenzyme A gCOSY - Correlation Spectroscopy CrO3  - chromic acid Cys  - L-cysteine d  - doublet dba  - dibenzylidene acetone DC50  - half maximal deterrence concentration DCE  - dichloroethane DCM  - dichloromethane dd  - doublet of doublets DEPT  - Distortionless Enhancement by Polarization Transfer  DMF  - N, N-dimethyl formamide DMAc  - N, N-dimethyl acetamide DMAP  - 4-(dimethylamino)pyridine DMP  - Dess-Martin periodinane DMSO  - dimethyl sulfoxide DMSO-d6 - deuterated dimethyl sulfoxide DNA  - deoxyribonucleic acid dpent  - doublet of pentets dppf  - 1,1’-bis(diphenylphosphino)ferrocene dt  - doublet of triplets EC50  - half maximal effective concentration e.g.  - exempli gratia (Latin) eq  - equatorial ER  - endoplasmic reticulum ERAD  - endoplasmic-reticulum-associated protein degradation Et  - ethyl EtOAc  - ethyl acetate EtOH  - ethanol del F508 - deletion of a phenylalanine residue at position 508 FDA  - U.S. Food and Drug Administration fwt  - formula weight g  - gram(s) G155D - deletion of a glycine at position 155 GABAA - γ-amino butyric acid receptor type A Gly  - glycine 1H  - proton [H]  - reduction hν  - light   xxx HCl  - hydrochloric acid HDAC  - histone deacetylase Hex  - hexanes HIV  - Human Immunodeficiency Virus gHMBC - Heteronuclear Multiple-Bond Correlation spectroscopy HPLC  - High Pressure Liquid Chromatography hour(s)  - hour(s) HRESIMS - high-resolution electrospray ionization mass spectrometry HSP-70 - 70 kilodalton heat shock protein  gHSQC - Heteronuclear Single Quantum Coherence spectroscopy Hz  - hertz IBX  - 2-iodoxybenzoic acid IC50  - half maximal inhibitory concentration IDO  - Indolamine 2,3-Dioxygenase i.e.  - id est (Latin) INADEQUATE   - incredible natural-abundance double-quantum transfer experiment IFN-γ  - interferon-gamma IR  - infrared spectroscopy ITS  - internal transcribed spacer J  - coupling constant (in hertz) Ki  - inhibition constant KI  - potassium iodide KMnO4 - potassium permanganate KOAc  - potassium acetate KOtBu  - potassium tert-butoxide l  - liter LH20  - Sephadex LH20 LiBH4  - lithium borohydride LRESIMS - Low-resolution electrospray ionization mass spectrometry Lys  - L-lysine m  - multiplet (NMR) or meter depending on context M  - molar mCPBA - meta-chloroperoxybenzoic acid Me  - methyl MeOH  - methanol MeOD-d4 - deuterated methanol MeSO3H - methanesulfonic acid Met  - L-methionine mg  - milligram MgSO4 - magnesium sulfate MHz  - megahertz min  - minute mL  - milliliter mm  - millimeter mmol  - millimole   xxxi MnO2  - manganese dioxide mp  - melting point MtB  - Mycobacterium tuberculosis MTPA  - α-methoxy-α-(trifuoromethyl)phenylacetyl chloride mTOR  - mammalian target of rapamycin m/z  - mass-to-charge ratio NaBH4  - sodium borohydride NAD+  - nicotinamide adenine dinucleotide NaHCO3 - sodium bicarbonate NaOH  - sodium hydroxide NaN3  - sodium azide Na2S2O3 - sodium thiosulfate NBS  - N-bromosuccinimide NCI  - National Cancer Institute NEt3  - triethyl amine ng  - nanogram NH4Cl  - ammonium chloride nM  - nanomolar nm  - nanometer NMR  - Nuclear Magnetic Resonance NOE  - Nuclear Overhauser Effect NOESY - Nuclear Overhauser Effect Spectroscopy [O]  - oxidation 18O  - oxygen isotope containing 10 neutrons ORTEP - Oak Ridge Thermal-Ellipsoid Plot PARP  - poly [ADP-ribose] polymerase PCR  - Polymerase Chain Reaction pent  - pentet PKS  - polyketide synthase pM  - picomolar ppm  - parts per million PTK  - protein tyrosine kinase Pyr  - pyridine q  - quartet R  - rectus RNA  - ribonucleic acid r.t  - room temperature S  - sinister s  - singlet SAHA  - suberoylanilide hydroxamic acid SAM  - S-adenosyl methionine SAR  - structure activity relationship SD  - standard deviation SDS-Page - sodium dodecyl sulfate polyacrylamide gel electrophoresis Ser  - L-serine   xxxii siRNA  - short interfering RNA sp.  - species Staph.  - Staphylococcus t  - triplet TB  - Tuberculosis td  - triplet of doublets THF  - tetrahydrofuran Tf  - triflyl TFA  - trifluoroacetic acid Tf2O  - triflic anhydride TLC  - Thin Layer Chromatography TMS  - trimethyl silyl TNF-α  - tumor necrosis factor-alpha TOF  - Time of Flight tRNA  - transfer RNA tROESY - Rotating Frame Nuclear Overhauser Effect Spectroscopy Trp  - L-tryptophan Tyr  - L-tyrosine UPLC  - Ultra High Pressure Liquid Chromatography UPR  - unfolded protein response UV  - ultra violet WT  - wild type       xxxiii Acknowledgements The number of people to whom I owe a debt of gratitude for helping to make this degree and document possible is far to large to fit within the confines given here.  I would, however, like to thank a few individuals who were particularly instrumental throughout this process.  I would first and foremost like to thank my supervisor Professor Raymond J. Andersen for his patience, support and encouragement. None of the work presented here would have been possible without his positivity and guidance. I would like to thank Dr. David E. Williams and Mike LeBlanc for their help and guidance in the lab. I would also like to thank Professor E. Dilip de Silva for his mentorship.  A great deal of thanks goes to Dr. Doralyn Dalisay for her help in growing microorganisms for many of my projects.  I would also like to thank all members of the Andersen Lab both present and past for their help and support. Thanks also go to the collaborators that I worked with over the course of my graduate career. Professors Grant Mauk, David Thomas, and Murray Isman provided a tremendous learning environment where I felt free to ask questions and engage with the subject matter.  Many other collaborators have greatly helped me and provided invaluable insights and contributions to the projects presented. They include in no particular order: Dr. Vivian Miao, Dr. Santiago Ramon-Garcia, Dr. Anne Steino, Dr. Graeme Carlile, and Dr. Fred Rosell. Special thanks goes to my loved ones.  Thanks to my mother, Beryl, for being a constant source of support and inspiration. Thanks to my father, John, who has always encouraged me to be curious and work hard.  To my sisters Cara and Anya, and brothers David and Jonathon, for being immensely supportive.  Lastly, I would like to thank my fiancé Leana; I couldn’t hope to find a more loving and inspiring partner.  Simply put, she is the best.   xxxiv Dedication           For Ruth, Roy, John, and Mary.    1 Chapter 1: Introduction to Natural Products Chemistry 1.1  Natural Products Chemistry The discipline of natural products chemistry aims to describe and characterize purified chemical entities from natural sources and determine their potential utility. The field is so broad and diverse that its limits are often blurred with others such as synthetic organic chemistry, molecular biology, pharmacology, medicine, microbiology, ecology, and anthropology.  In the following introductory chapter an overview of natural products chemistry will be presented through the lens of drug discovery. Several examples where natural products have made an indelible mark on modern society will be integrated into discussions on the history, techniques used in the field, and an overview of the biosynthetic pathways employed by the producing organisms.  Hopefully, this brief introduction will provide the reader with a context onto which the subsequently presented research can build. 1.2  A Brief History of the Field  Natural products chemistry can trace its origins to the advent of modern chemical sciences in the early 1800’s. However, the roots of this research field harken back much further, to the first uses of plants and animals by early humans for the purposes of healing, dying fabrics, and ceremonial or religious practices. In many ways these traditional uses provided the foundation for the modern chemical study of natural products.1 The first documentations of natural product use can be traced to the dawn of early writing systems. The earliest surviving texts, written in cuneiform, come from ancient Mesopotamia ~2,600 B.C., where the uses of nearly 1,000 plants and plant products are described.2–4 The ancient Egyptians, famous for their written documentation, also produced one of the original medicinal natural products texts, the Ebers Papyrus.2, 5 This nearly 3,600-year-old document describes 800 prescriptions involving roughly 700 plant products for many different ailments.5 Later, the Greek “father of medicine” Hippocrates (460–377 B.C.), wrote a compendium of nearly 400 natural agents, many of them plant derived, to be used as medicines.6 These western texts were mirrored by contemporaneous recordings in Indian Ayruvedic texts Charaka Samhit (~900B.C.) and Sushruta Samhita (~600 B.C.), and the traditional Chinese medical text Wu Shi Er Bing Fan (~350 B.C.).5,7 All of these early recordings explicitly show the importance of plant and animal derived natural products in the maintenance of human health in ancient societies.   2  In 1804, a major leap forward in medicine and natural products chemistry transpired with the isolation of morphine by the German pharmacist Friedrich Serturner.8 This marked the first instance of a purified medicinally relevant chemical substance being isolated from its natural source, Papverum somniferum. By 1826, the German pharmacist Emanuel Merck, offered Serturner’s isolated morphine (1.1, Figure 1.1) as the first chemically pure plant derived medicine.8 Subsequently many plant natural products, most of them alkaloids, were isolated, purified, and characterized by chemists in the 1800s.8  Figure 1.1 Morphine (1.1) and salicylic acid (1.3) and their acetyl derivatives Another major breakthrough in natural products chemistry and medicine came in 1897, when Felix Hoffman, while working for the Bayer Co., synthesized diacetylmorphine (1.2, Figure 1.1).9 This compound was found to be nearly twice as potent as morphine (1.1) and was sold by Bayer under the trade name “Heroin” (1.2, Figure 1.1). This fortuitous discovery by Hoffman was repeated only two weeks later with the anti-arthritis compound; salicylic acid (1.3, Figure 1.1).9 Felix Hoffman used acetyl modification to ablate the stomach irritation this natural product caused. The product, acetosalicylic acid (1.4, Figure 1.1), named aspirin (1.4, Figure 1.1) (a-acetyl, spirin- short for Spiraea ulmaria “meadow sweet” a source for salicylic acid) was patented and marketed in 1900 by Bayer Co. as a “miracle” general pain reliever.9 Heroine and aspirin represent the first semi-synthetic natural products produced by mankind, and their discovery marked a major turning point in the development strategy of natural products as medicines.  The next leap forward in the field of medicinal natural products, and arguably the most important of the 20th century, was the discovery of penicillin (1.5, Figure 1.2). In 1928, Alexander Fleming observed a Penicillium mold which cleared bacterial growth.  He named the antibiotic substance penicillin, but was unable to purify or identify the compound.10 In 1940, Howard Florey and Ernst Chain were able to purify and mass-produce penicillin (1.5, Figure 1.2) for use as an antibiotic; a feat which saved countless lives during World War II and continues to OHO HH NOH O OHOH O OHO OOO HH NOO O1.1 1.2 1.3 1.4  3 do so to this day.11 All three scientists, Fleming, Florey and Chain, were awarded the Nobel prize in 1945 for their discovery.  However, it wasn’t until 1945 that Dorothy Crowfoot Hodgkin solved the chemical structure for penicillin (1.5, Figure 1.2).12 To this day penicillin (1.5, Figure 1.2) and many derivatives (e.g. amoxicillin 1.6, Figure 1.2) are still widely used as antibiotics for bacterial infections. This discovery became a tipping point for natural products research focusing on fungal and microbial sources, a trend that still endures today.     Figure 1.2 Penicillin G (1.4) and amoxicillin (1.6)   The use of terrestrial plant and microbial natural products in medicine was well documented by the 1940’s, but the marine realm remained unexplored. That changed in 1950 with the discovery of the sponge nucleosides spongothymidine (1.7, Figure 1.3) and spongouridine by Werner Bergman.13–15 These compounds became the inspiration for the synthetic compounds Ara-C (1.8, Figure 1.3) and Ara-A (1.9, Figure 1.3), two of the first antimetabolite cancer chemotherapeutics to be used in the clinic.16 Ara-C (1.8) and Ara-A (1.9) represent the first clinically approved marine natural product derived drugs, and their discovery established the marine environment as a viable source for drug discovery.   Figure 1.3 Spongothymidine (1.7) with Ara-C (1.8) and -A (1.9) The field of natural products has continued to produce a large number of medicinally relevant small molecule scaffolds.  In a review from 2012, natural product small molecules were O HN NO S OHO O HN NO S OHONH2HO amoxicilin (1.6)penicillin G (1.5)OOHHO N NHO O OOHHO N N ONH2 NNNN NH2OOHHO HOHOspongothymidine (1.7) Ara C (1.8) Ara A (1.9)HO  4 shown to account for roughly 28% of all drugs approved between 1981–2010.17 If we take into account the fact that a large number of pharmaceuticals are actually synthetic mimics of natural products, that percentage increases to a staggering ~64% of all approved small molecule drugs.17      For anticancer drugs the numbers are even more convincing, as more than 70% of all the small molecules approved for cancer treatment are in some way derived from a natural product scaffold.17 These figures argue that natural products are an unsurpassed resource for initial drug development and offer structures prescreened by nature for biological relevance and efficacy.17   1.3 Producers of Natural Products  The exploration of a wide variety of source organisms such as plants, microbes, and invertebrates has been a strategy of natural products chemists since the 19th century.  The search for new biomes to explore and novel organisms to study has driven natural products chemists to some of the most remote places on earth. Source organism collection sites vary from tropical Papua New Guinea,18 to frigid Antarctica,19 or even deep-ocean hydrothermal vents,20 each offering unique competitive environments for organisms to evolve both physically and chemically.  In recent years, a paradigm shift towards microbial and genomic research has raised questions about the identity of the actual producer of many natural products isolated from macro-organisms.  Recent findings have shown that camptothecin (1.10), vinblastine (1.11) and vincristine (1.12) (Figure 1.4), all plant derived anticancer agents, are likely produced by endophytic fungi.21, 22 Recently it was shown that a phylum of obligate gram-negative bacteria (Tectomicrobia => Entotheonella) is responsible for producing the majority of the complex natural products isolated from the sponges they inhabit (e.g., Theonella swinhoei).  Natural products like onamide A (1.13) or aurantoside A (1.14) (Figure 1.4) have been verified through genome mining and gene cluster isolation to arise from these microbes.23 Following these findings, little doubt remains that the true producers of many natural products are in fact the microbiota residing within the host macro-organisms.   5  Figure 1.4 Plant and sponge derived natural products likely produced by microorganisms21–23 1.4 Biosynthetic Route to Natural Products Regardless of the identity of the producers of natural products, these compounds are made biosynthetically through either a primary metabolic pathway or more often than not, a secondary metabolic pathway. There are five canonical biosynthetic pathways for secondary metabolites: polyketide, terpenoid, shikimate, non-ribosomal/ribosomal peptide, or alkaloid pathways.  It is important to note that although many natural products are constructed using any one of these five pathways, many arise from the combination of two or more biosynthetic routes.  O O HN OO O O O NH NHH2N NHCOOHHOOHO onamide A (1.13)N N OO OHOcamptothecin (1.10)NHNHO H NOO NHO OH OO O OO vincristine (1.12)NHNHO H NOO NH OH OO O OO H Hvinblastine (1.11)Cl Cl NOOOH O OHOHOOOH OHOOH2N OO OHaurantoside A (1.14)  6 Therefore, the following five categories are by no means the only methods for biosynthesis of natural products. 1.4.1 Polyketide Biosynthesis  Polyketides are constructed by the successive condensation of acetate with malonyl CoA (other units such as propionyl CoA have been observed) through an enzyme catalyzed Claissen condensation (Figure 1.6) to form an elongated carbon chain of 1,3 dicarbonyls.  The β-carbonyl in the chain is successively reduced after each condensation (reducing) or kept fully oxidized (non-reducing) until the full chain has been created.  There are three classes of polyketide synthetases: types II, III, and I. Type I-PKSs are large modular enzymes that are either iterative (each enzyme is reused multiple times to produce a growing chain), or non-iterative (each module is assigned a specific function).25  Type II-PKSs are a conglomeration of several polyfunctional enzymes that act in an iterative fashion.  The last class, type III-PKSs, are homodimeric enzymes that employ a single active site to produce products.  A non-reducing iterative polyketide synthetase (usually type II or type III) produces aromatic polyketides, by reusing condensation domains to elongate a chain of 1,3 dicarbonyls.  This chain is then cyclized into a polycyclic aromatic product, such as tetracycline (1.15, Figure 1.5), through the elimination of water and then further elaborated post-translationally.25 A reducing iterative type I-PKS, as the name suggests, reduces each newly formed ketone into a methylene unit in an iterative fashion.  As such, reducing type I-PKSs are analogous with fatty acid biosynthesis and the lipid products can be primary metabolites. Non-iterative type I-PKSs are large multi-domain enzymes that can produce highly functionalized products like the macrolide antibiotic monensin (1.16, Figure 1.6).24 An abbreviated biosynthetic scheme for monensin (1.16) is shown below in Figure 1.6.25  Figure 1.5 Type II PKS product tetracycline (1.15) O O NH2OHHOOOH NMe2OHOHtetracycline (1.15)H H  7  Figure 1.6 Type I PKS biosynthetic route for monensin (1.16).25 1.4.2 Terpene Biosynthesis  Terpene biosynthesis, like polyketide biosynthesis, can be involved in both primary (cholesterol, estrogen, vitamin D) and secondary metabolism.  Terpenes are created using isoprene C5 units, which are usually connected in a head-to-tail fashion and then cyclized and derivatized.25 The biosynthetic machinery creates dimethyl allyl pyrophosphate (DMAPP) and isopentenyl pyrophosphate (IPP) as building blocks from either mevalonic acid or methylerythritol phosphate.25 Terpene cyclase enzymes then generate a carbocation through epoxide ring opening or double bond protonation. A polyene cascade usually ensues and creates the parent carbon skeleton, while oxidative enzymes (e.g., cytochrome P450’s) supply the point oxidations and additions that create the final products.25 Terpenes are classified by the number of isoprene units present in the carbon backbone, C5-hemi, C10-mono (1.17, Figure 1.7), C15-sesqui, C20-di (1.19, Figure 1.7), C25-sester, C30-tri, carotenoids (between C35-45) and poly-terpenes O SEnz+ X 4 + CO2HOSCoAX 7 CO2HOCoASHO OHHOO OO SEnzyme OO OHOHCO2HHOOO OHOHCO2HHO O OOHOO OOHCO2HO O O O OHmonensin (1.16)monensin epoxidasemonensin PKS Tailoring enzymesPKS release +–CO2O SEnz BOSCoAO OH  8 (natural latex rubber).25 Any degraded compounds derived from terpene precursors, that contain a number of carbon atoms less than a multiple of C5, are called nor-terpenoids. An example of this is cholesterol, a C27 sterol derived from the C30 triterpene lanosterol (1.18, Figure 1.7).25 These compounds come from the oxidative degradation of a normal terpenoid starting material.    Figure 1.7 Terpene biosynthesis25 1.4.3 Shikimate Biosynthesis  The Shikimic acid pathway is an important primary and secondary biosynthetic pathway, which provides a route to all of the aromatic amino acids L-phenylalanine, L-tyrosine (1.20, Figure 1.8) and L-tryptophan and many aromatic natural products.25 This pathway is present in plants and microbes but is absent in animals. Therefore, all of the aromatic amino acids are said OPPOPP PP = P OO POOHOHHODMAPP IPP OPPH H B EnzOPPgeranyl diphosphate (GPP) OPPfarnesyl diphosphate (FPP) OPPgeranylgeranyl diphosphate (GGPP)DiterpenesMonoterpenesSesquiterpenes FPP squaleneTriterpenes and Steroidstaxadiene HO OAcO OOAcO OO OHOHONHO paclitaxel (1.19 )HO lanosterol (1.18 )H HS-(-)-limonene(citrus)(1.17 )OPP OPPGPP HB+  9 to be essential, i.e. obtained through dietary supplementation in animals.  The products of the shikimate pathway are incredibly varied, frequently aromatic in structure, and represent some of the most economically important natural products.25 Fragrant compounds such as vanillin (1.21, Figure 1.8) (vanilla scent) or cinnamaldehyde (cinnamon scent), structural compounds such as the wood polymer lignin, and medicinally relevant molecules like salicylic acid (1.3, Figure 1.1), are all made using the shikimate pathway.25 The biosynthetic route to shikimic acid (1.22), the namesake and branch point in the shikimate pathway, begins with D-erythrose 4-phosphate (1.23) and phosphoenol pyruvate (1.24).25 The basic pathway from these starting materials to shikimic acid (1.22) is shown below in Figure 1.8.    Figure 1.8 Shikimic acid formation and examples of economically important shikimate products25 1.4.4 Non-Ribosomal Peptide and Ribosomal Peptide Biosynthesis  There are two pathways by which peptide containing natural products are usually produced, the ribosomal and the non-ribosomal pathways.  The ribosomal pathway, as the name suggests, produces peptide scaffolds constructed with the ribosome using the 20-proteinogenic amino acids. These polypeptides are then altered and modified post-translationally.26 Ribosomally produced peptides (RP) are often constructed containing a leader peptide, which is proteolytically cleaved once the active peptide has been constructed and post translational modifications have been completed.26 The sequences for these peptides are also encoded next to or near the genes for modifying enzymes and the enzymes responsible for their producer’s resistance.26 The posttranslational modifications can include, but are not limited to, P OHO2C OOHHOPO PO O CO2HOHOHHOH O CO2HOHOHHO HO CO2HO OHOH CO2HHO OH OHshikimic Acid (1.22)CO2HOH O CO2HHO2CHO OCO2HNAD+PLPOH NH2CO2HL-Tyr (1.20) H chrosimic AcidD-erythrose 4-phosphate (1.23)phosphoenol pyruvate (1.24)OHCO2HOHO Ovanillin (1.21) 4-coumaric acid 3-dehydroquinic acid  10 epimerization, aromatization, methylation, disulfide formation, lanthionine formation and even macrolactonization.26 Ziconotide (Prialt®) (1.25, Figure 1.9), an analgesic cone shell snail toxin, is a clinically approved ribosomally derived peptide.27, 28   Non-ribosomal peptides (NRPs) are produced using large, modular enzymes called non-ribosomal peptide synthetases (NRPSs).25 These enzymes sequentially construct the core peptide scaffold of their products in much the same fashion as polyketide synthases.25 Each module or domain contains a discrete series of enzymes, which adds a monomeric unit to the growing product chain. Much like RPs, NRPs are known to contain post-translational modifying enzymes, which further manipulate the core structure once created in full.25 Unique to NRPs is the ability to incorporate non-proteinogenic amino acids as starting materials, and posttranslational N-methylation of the peptide backbone, which confers resilience to hydrolysis.25, 26 Some of the most important clinically approved medicinal natural products are NRPS derived, including the penicillins (1.5, Figure 1.2) and dolastatins (1.26, Figure 1.9).29    Figure 1.9 RP and NRP products Prialt® (1.25) and dolastatin 10 (1.26).27–29 1.4.5 Alkaloid Biosynthesis  Alkaloids by definition are compounds that contain at least one basic nitrogen atom; therefore, they are an extremely diverse and complex group of natural products.  Many biosynthetic pathways are used to construct alkaloids, including every previously discussed pathway. A large number of alkaloids are biosynthetically derived from the combination of two or more secondary metabolic pathways such as the fungal derived hybrid PKS-NRPS responsible for penicillin G (1.5).30 However, many classes have dedicated biosynthetic machinery that are unique to their production.  These enzymes often rely on amino acid starting materials, which are then dimerized, degraded, or rearranged.  Examples of this can be seen with the biosynthesis of alkaloids like lysergic acid (1.27), quinine (1.28), or staurosporine (1.29)(Figure 1.10).25  Cys Lys Gly Lys Gly Ala Lys Cys Ser Arg Leu Met Tyr Asp Cys Cys Thr Gly Ser Cys Arg Ser Gly Lys Cys NH2NS O O NHO O NO O NHO NPrialt! (1.25 )dolastatin 10 (1.26 )  11  Figure 1.10 Lysergic acid (1.27), quinine (1.28) and staurosporine (1.29)  Another class of alkaloids, the oroidins (1.30, Figure 1.11), a pyrrole imidazole-containing class of sponge alkaloids, offers an example of how simple starting materials (proline (1.31) and lysine (1.32), Figure 1.11) can be biosynthetically transformed into a diverse class of natural products.31 This particular pathway is known to produce some of the most interesting and diverse structures found in marine natural products chemistry, such as palau’amine (1.33, Figure 1.11).32 However, the biosynthetic enzymes, responsible for these putative transformations, have yet to be elucidated. The possible clinical uses of members of this class, such as sceptrin (1.34, Figure 1.11) for inhibition of cell motility, are currently being evaluated.33, 34 A discussion of latonduine, a member of this class of alkaloids and its biological activity are presented in Chapter 6. NN OHO HN NHHO2C H N NHN OOO NH Hlysergic acid (1.27) quinine (1.28) staurosporine (1.29)  12  Figure 1.11 Oroidin proposed biosynthetic pathway based on labeling studies33, 34 1.5 Methods Used for Natural Products Chemistry  Natural products chemists rely on a large number of techniques in order to isolate and characterize the constituents of an extract from an organism.  A brief overview of the techniques used for isolation of natural products, specifically those used in subsequent chapters, will be described.  An overview of characterization techniques, focusing on Nuclear Magnetic Resonance (NMR) spectroscopy experiments, will also be provided.  1.5.1 Isolation  Isolation techniques and analytics have evolved greatly over the past two centuries.  Early isolation chemists relied heavily on harsh extraction techniques using acids, bases, and high temperatures (e.g., soxhlet extraction). These primitive extraction mechanisms, though fruitful, allowed for the isolation of only the most robust, abundant, and crystalline organic scaffolds.13–15 Today, initial extraction is often done under mild conditions, with organic solvents such as MeOH or methylene chloride. The concentrate of this extract, obtained using reduced pressure evaporation, is then partitioned between a number of increasingly polar organic solvents and NH OOH [O] NH OOH Bromination NH OOHBrBrH2N CO2H H2N NH2OH2N CO2HHN H2N CO2HHN[O]NH2 NH2NH NH2NHO H2N CO2HHN NNH2+- CO2- H2O[O] NHHN O BrBrNNHH2N oroidin (1.30 )NHHN OBrNHNH2N NHNHO BrN NHH2N sceptrin (1.34 ) NN NHNHN NO NH2H2N OHNH2ClH H(-)-palau'amine (1.33 )lysine (1.32 ) proline (1.31 )  13 water.  Purification of a desired component relies heavily on multiple rounds of chromatography until a single purified substance is obtained.   Liquid chromatographic techniques are frequently employed after initial extraction and partitioning. These include: size exclusion (e.g., LH20), normal phase (silica gel), and reversed phase (C18 coated silica gel) chromatography.  The use of high-pressure (HPLC) or ultra high-pressure (UPLC) liquid chromatography is frequently employed for final purification. After purification, the compound can then be characterized with the confidence that all data collected will be a result of the single compound in question.   1.5.2 Characterization  Structure elucidation is a time consuming and challenging task, relying on the combination of several techniques to accurately determine the shape and connectivity of a molecule. Mass spectrometry and various nuclear magnetic resonance (NMR) experiments are the most frequent characterization techniques employed.  The molecular formula and the degrees of unsaturation in a given molecule are routinely determined using high-resolution mass spectrometry. In order to establish the connectivity, 1D and 2D NMR experiments are performed including COrrelation SpectroscopY (COSY) (2-, 3-, 4-, and 5-bond homonuclear correlations), Heteronuclear Single Quantum Coherence spectroscopy (HSQC)(indirect detection 1-bond heteronuclear), and Heteronuclear Multiple-Bond Correlation (HMBC) (indirect detection 2–4-bond heteronuclear) experiments.  Each of these experiments relies on scalar coupling between the irradiated nucleus and its coupling partner nuclei.  Signals from these experiments are plotted as a contour or intensity plot at the intersection between the chemical shifts of the two nuclei in a 2D plane.   Several other experiments, such as Nuclear Overhauser Effect Spectroscopy (NOESY) and Rotating frame nuclear Overhauser Effect SpectroscopY (ROESY) give through-space coupling information in a similar format as above, often allowing for assignment of relative configuration in chiral molecules.  When combined together, the high-resolution mass spectrum and various NMR experiments usually provide enough information for a structure to be elucidated and relative configuration assigned.  Various derivatization methods (e.g., Mosher’s or Marfey’s), X-ray crystallography, or CD spectroscopy can then be employed as a means for establishing the absolute configuration of a molecule. Numerous accompanying experiments   14 such as UV spectroscopy or IR spectroscopy can provide support for a challenging structural assignment. 1.6 Modern Natural Products in Drug Design and Discovery  Natural products, as discussed earlier, are unparalleled as a source for novel drug scaffolds and leads.  In the past decade (2000–2010), the number of new chemical entity small molecules to be approved as drugs averaged ~25 each year.17 This is a major decline from previous decades, which averaged 40 approvals per year.17 The contributing factors to this decline are a contentious subject, but many implicate the shift in research focus away from natural products and towards combinatorial methods for drug discovery.17 This decline in approvals, however, did not affect the direct contributions of natural products to the field of drug discovery, as nearly 50% of all molecules approved over the past decade were natural products or close derivatives thereof.17     Drug approval rates offer a myopic view of the overall impact made by natural products. Often, natural products are exquisite tools to probe our understanding of medicinally relevant biological processes.  For example, the compound rapamycin (1.35, Figure 1.12), a polyketide macrolide isolated rom Streptomyces hygroscopicus, was used as a molecular tool to describe a thereto unknown pathway for immunosuppression named mTOR (mammalian target of rapamycin).35 This pathway has become an active area of research, and has produced clinically relevant discoveries, which have led to the approval of rapamycin (1.35, Figure 1.12) and analogues as drugs for immunosuppression and cancer treatment.36, 37  Another example of a natural product being used as a molecular biology tool can be seen with trapoxin (1.36, Figure 1.12).37 This modified peptide natural product, through protein profiling studies, was found to inhibit certain histone deacetylases (HDAC).37  From this observation, a better understanding of histone acetylation in cell cycle and progression was gained, eventually leading to several HDAC inhibitor drugs (e.g., Romidepsin)38 for the treatment of cancer.37 These two examples offer a glimpse of the countless instances where a natural product has aided in the unraveling of a biological process, leading to drug development.        15   Figure 1.12 Rapamycin (1.35) and trapoxin (1.36)35, 37 1.7 Conclusion  The staggering number of clinically approved drugs derived from natural products offer reason enough to continue looking to natural sources for answers to pressing global health concerns. However, these numbers reflect only a small fraction of the contributions natural products have provided to the study of human health and disease.  Many more natural products, never approved for clinical use, have aided our understanding of the underlying biology and biochemistry in many diseases. In the next five chapters, several novel natural products as tools and technological leads will be presented. The majority of these studies aim to provide greater understanding of a medicinally relevant target, while simultaneously supplying a chemical tool towards disease treatment. N NHHNNHOOO OOOtrapotoxin (1.36)OHO OO N O O OHOO O OO OHrapamycin (1.35)  16 Chapter 2: Ramariolides A–D, Novel Antimycobacterial Butenolides From the Pacific Northwest Native Coral Mushroom Ramaria cystidiophora.  2.1  Tuberculosis Tuberculosis (TB), caused by Mycobacterium tuberculosis (Mtb), is and has been one of the world’s most deadly diseases. It was estimated that in 2012, the number of new TB cases was nearly 8.6 million, with approximately 1.3 million of those cases being terminal.39 Tuberculosis is widely known as a disease of the lungs (pulmonary TB), but it can also affect other areas of the body (extra pulmonary TB).  TB is most often spread through inhalation of sputum containing bacteria from someone with pulmonary TB.39 In general, the percentage of people who are infected by M. tuberculosis (latent TB) is much higher than those who develop the disease (active TB).  However, immune-compromised persons, such as HIV patients, have a higher incidence of infection and mortality.39   Tuberculosis is more common amongst men and is statistically more likely to affect adults of working age than children or the elderly.  The disease is deadly if left untreated, killing approximately 70% of people who contract it without treatment within 10 years.39, 40 Several treatments for TB have been developed in the last century, including a preventative vaccine (Bacille Calmette-Guerin; BCG vaccine), which unfortunately has been shown to be relatively ineffective in preventing pulmonary TB.39, 41 Starting in the 1960’s, several drug regimes were developed which aimed to cure the disease once contracted.39, 42  Early attempts at clinical treatment of TB revealed the infections’ ability to relapse after monotherapy, i.e. treatment with a single drug like streptomycin (2.1, Figure 2.1).42 This has led to the development of combination therapy for TB treatment.39 The major first-line treatment for active TB today is a six-month combination therapy of four drugs: rifampicin (2.2, Figure 2.1), isoniazid (INH) (2.3, Figure 2.1), ethambutol (EMB) (2.4, Figure 2.1), and pyrazinamide (PZA) (2.5, Figure 2.1).39, 42 This treatment has lead to outcomes of 85% clearance rates or higher for non-immunocompromised patients.39  In populations where TB infection rates are high, treatment to remove the latent infections is begun before an active infection can occur.  In this case, the use of INH (2.3, Figure 2.1) for 9 months is the standard of treatment.39 Despite the   17 introduction of these treatment regimes, the development of resistance is still a major problem in combating TB.39   Currently, three broad terms define the drug resistant strains of TB: multidrug-resistant TB (MDR-TB), extensively drug-resistant TB (XDR-TB), and totally drug-resistant TB (TDR-TB). MDR-TB is an infection by a strain of M. tuberculosis resistant to two of the first line TB drugs, rifampicin (2.2, Figure 2.1) and isoniazide (2.3, Figure 2.1).39 XDR-TB is defined as a subset of MDR–TB, where the infection has resistance to most of the treatments available including the second line course of injectable drugs. Totally drug resistant TB (TDR-TB) is characterized as being resistant to all known TB drugs. Though a few cases have been reported, exact figures are difficult to obtain.39 In 2012, there were an estimated 450,000 cases of MDR-TB reported, with approximately 9.6%, up from 9% in previous years, of those being XDR-TB.39 In general, the treatment regimes for these cases are prolonged to at least 20 months, with stronger antibiotics, and much lower success rates of clearance.39 Therefore, new treatment options and new drugs are needed to treat the increasing amount of resistance found to current drug regimes.  Figure 2.1 Streptomycin (2.1) and the current first-line antibiotics 39, 42 2.1.1  Mycobacterium tuberculosis (Mtb): Mycobacterium tuberculosis (Mtb) is a member of the Mycobacteriacea family of Actinobacteria, a unique group of prokaryotes given the moniker “myco” due their fungal-like NO HN NH2 NH HNOH HO OMeOAcOHHO O NHOHOH OHO OH N N NOO ONHMeHO OHHO OHO OO H HO N NOHOHH2N NH2 NH2NH2 NN NH2O2.1 2.22.3 2.4 2.5  18 appearance during cultured growth.43 The cellular envelopes of the Mycobacterium genus are their defining feature and are unique among prokaryotes.  The cell envelope of Mtb, like all other mycobacteria, consists primarily of three components: a MAPc matrix, a plasma membrane, and cell capsule rich in polysaccharides.44–46 This unique waxy rampart imparts a strong resistance to host immune attack and antibiotics.45, 46  The MAPc component of the cell wall is unique to mycobacteria, and consists of three building blocks, mycolic acid, arabinogalactan, and a peptidoglycan complex.45, 46 The most notable component of the cell wall are the mycolic acids (example 2.6, Figure 2.2).  These fatty acids are unique to mycolata, and are the name-sake fatty acid family of Mycobacterium.47 Mycolates are characterized by a long chain C54 to C63 beta-hydroxy subunit and a shorter C22 to C24 alpha alkyl side chain and come in three different substituted forms alpha, methoxy and keto-mycolates.45–47 The substitution differences in these side chains are hallmarks of the different species in this genus.   All mycobacterial mycolic acids (example 2.6, Figure 2.2) contain cyclopropyl moieties, which help provide the structural integrity of the cell wall, making it more rigid in comparison to analogues lacking cyclopropanation.47 The cyclopropyl groups also account for resistance to oxidative stress by providing resistance to reactive oxygen species.47 In the cell wall the mycolic acids are usually linked to either of two carbohydrate moieties, arabinose or trehalose.45–47 The arabinose-linked mycolate is called arabinogalactan-mycolate; a branched polymer of arabinose and galactose linked through a phosphodiester bond to the peptidoglycan substructure of the cell membrane. Mycolic acids and trehalose form a free glycolipid called trehalose dimycolate (TDM). TDM collects on the outer surface of the cell wall as a thick lipid layer, hindering host immune cell attack and antibiotic diffusion.47 These TDM units, which sit atop arabinogalactan-mycolate polymers, templated onto a peptidoglycan layer, provide the basic backbone for the three dimensional structure of the cell wall in Mtb (Figure 2.2).46, 47   19  Figure 2.2 A diagram of the cell capsule’s underlying structure in Mtb and an example of a mycolic acid. 46, 47  Figure 2.3 Several drugs used to inhibit Mtb cell wall biosynthesis or mycolic acid production42 2.1.2 Tuberculosis Drugs: Drugs Involving the Cell Capsule of Mtb The architecture of the mycobacterial cell wall, specifically that of Mtb, has been the subject of a great deal of research, with several reviews available.45, 46 This attention is largely due to the cell envelope as a first point of contact for any treatment that one might attempt.  Any drug that is to be developed for Mtb must either be sufficiently lipophilic to pass through the cell wall passively like rifampicin (2.2, Figure 2.1), or actively transported through the cell HOOHO 30mycolic acid ( 2.6 ) Plasma MembraneGrannular layerPeptidoglycan layerMycolic acid membraneArabinogalactan layerouter layerHNO O NH2 N SO HHN COOHON NH2S N N O O N O O CF3N+-OO OCF3OONNN+O-ON NHO OHHH CO2H2.7 2.8 2.92.10 2.112.12  20 membrane through a protein channel like pyrazinamide (2.5, Figure 2.1).46–48  This makes drug interaction with the cell wall a major hurdle for development of new therapies in TB.   An attractive area for the development of targeted TB antibiotics is the biosynthetic machinery of the cellular envelope.  In fact, two of the four first-line antibiotics used against Mtb, EMB (2.4, Figure 2.1), and INH (2.3, Figure 2.1) are inhibitors of a portion of the cell envelope’s biosynthesis.39, 48 EMB (2.4) is an inhibitor of arabinogalactan biosynthesis,49 and appears also to inhibit transport of mycolipids to the cell wall.48, 50 INH (2.3, Figure 2.1) has been shown to inhibit mycolic acid biosynthesis, likely inhibiting the enzyme responsible for cyclopropanation, causing disruption of the cellular envelope and eventual cell death. 48, 50–53 Cycloserine (2.7, Figure 2.3) and penicillins (2.8, Figure 2.3) are drug classes gaining use against Mtb due to their inhibition of peptidoglycan biosynthesis.48, 54 Historically, use of penicillins (2.8) has been precluded due to resistance of Mtb conferred by the presence of a native β-lactamase (BlaC).54 β-Lactamase inhibitors like clavulanate (2.9, Figure 2.3), in combination with penicillins (2.8, Figure 2.3), are now being proposed as possible alternative therapies for Mtb.55 Other drugs, approved or in clinical trials, which have mechanisms of action related to the cell wall include ethoniamide (2.10, Figure 2.3), delamanid (2.11, Figure 2.3), and PA-824 (2.12, Figure 2.3), all which inhibit the biosynthesis of mycolic acid.48, 56 Many other compounds, derived from natural products and medicinal chemistry, are currently in preclinical development.48, 56, 57 2.1.3 Tuberculosis Drugs: Alternative Mechanisms of Action The cell capsule of Mtb provides a proven and unique target for many Mtb related drugs. However, when sufficiently lipophilic to passively enter the cell capsule, several other antibiotics, with varying mechanism of action, have proven themselves as effective treatment options.48, 56, 57 Two of the first-line drugs used in clearance of active and latent Mtb infections, rifampicin (2.2, Figure 2.1) and PZA (2.5, Figure 2.1), work through mechanisms other than cell wall biosynthesis.48 Rifampicin (2.2, Figure 2.1), a member of the natural product family rifamycins isolated from an Amycolatopsis sp. in 1957,58 is one of the most potent sterilizing agents available for TB. Its mechanism of action involves inhibition of transcription through binding with high affinity to DNA dependent RNA-polymerase in bacteria.48 Pyrazinamide’s (2.5, Figure 2.1) mechanism of action is poorly understood, but it appears to act on the proton   21 motive force needed for cellular membrane transport leading to acidification of the cell and eventual death.48    Another well-known class of anti-mycobacterials are the aminoglycosides.  These potent natural product broad-spectrum antibiotics, typified by streptomycin (2.1, Figure 2.1), show a relatively poor absorption profile orally, needing to be administered by injection. They also display significant toxicity profiles.39, 48 As a result, they have been reserved largely for second line treatment for MDR-TB. These compounds bind to the 30s ribosomal subunit, ultimately inhibiting protein synthesis and translation.48 Macrolides, such as clarithromycin (2.13, Figure 2.4), are also natural product antibiotics gaining attention for the treatment of TB. Macrolides commonly work by inhibiting RNA dependent protein synthesis, binding to the bacterial 50s ribosomal subunit. In both macrolides and aminoglycosides low levels of inherent resistance by Mtb is likely attributable to low cell wall permeability.48, 57 Another unique class of antibiotics in the clinic for treatment of TB is the fluoroquinolones.  These compounds, exemplified by moxifloxacin (2.14, Figure 2.4), are powerful antibiotics which have the potential to shorten the duration of treatment for TB and are currently being used as second-line antibiotics.48 Fluoroquinolones act by trapping DNA gyrase as ternary complexes, effectively blocking transcription complexes and replication forks.48, 56, 59  The development of new and diverse drug classes to combat bacterial resistance is a never-ending arms race. One success story culminated in 2012 with FDA approval of bedaquiline (2.15, Figure 2.4), the first novel mechanism of action antimycobacterial introduced into the clinic in 40 years.  This drug, which acts on ATP synthesis in mycobacteria, is the latest link in a long chain of drug discovery efforts related to Mtb.39, 60 With resistance to bedaquiline (2.15, Figure 2.4) and all other approved antibiotics an ever present concern, the search is on for new tuberculosis drugs with novel mechanisms of action.39, 60   22  Figure 2.4 Mtb drugs with mechanisms of action other than inhibition of cell wall biosynthesis.39, 60 2.2 Mushrooms as a Source for Drug Leads Mushrooms, the macroscopic fruiting bodies of Basidiomycetes and some Ascomycetes, are the larger relatives of the microscopic molds and fungi commonly employed in drug discovery.  Mushrooms have been used in traditional medicine for hundreds of years and are an active area of natural products drug discovery.61 Some notable mushroom-derived drugs are the antimicrobial retapamulin (2.16, Figure 2.5), an antibiotic derivative of pleuromutilin, first isolated from Clitopilus passeckerianus.61–63 Irofullven (2.17, Figure 2.5), a derivative of illudin S, is a notable drug candidate that exhibits potent antimitotic activity.  It was isolated from Omphalotus illudins, the “Jack O’ Lantern” mushroom, and has progressed to phase III clinical trials.64–66 The scarcity of examples of mushroom-derived drugs in the clinic is possibly due to the inherently difficult nature of drug discovery research on such unpredictable and complex organisms. The differences in growth, chemo-geography, and size for many mushrooms often preclude collection of sufficient sample quantity and uniformity.61 Added to this is the fact that many wild mushrooms have so far proven uncultivable, making resupply nearly impossible. However, with the proper tools and adequate supply of material, mushrooms can be an important source of medicinally relevant natural products.61 O OH OMeO OHO OO OMeOHO NMe2HOO NNHNH HF OMe O O OHN HOOMe N2.13 2.142.15  23  Figure 2.5 Clinically relevant mushroom natural products61–66 2.2.1 Ramaria: Description, Distribution and Known Chemistry Ramaria is a genus of primarily ectomycorrhizal fungi, belonging to the phylum Basidiomycota, which are widely distributed throughout the world’s forests.  They are commonly called coral fungi due to their highly branched appearance, and some species are known to be edible (Figure 2.6).  At least sixty recognized species are represented in the Pacific Northwest region of North America, where species richness has been strongly associated with old growth forests.67, 68 Descriptions of natural products from Ramaria are rare.  To date, only three papers, the author’s work included, describe purified natural products from Ramaria species.  These include a novel ceramide isolated from R. botrytis, along with several previously reported ergosterol derivatives, and the known siderophore, pistillarin, from R. subaurantiaca.69, 71 No previous reports on the chemistry of R. cystidiophora have been published prior to our article, identifying it as a novel source organism for natural products isolation.  Figure 2.6 Ramaria sp. in situ. O HOS OHON OHOHO2.16 2.17  24 2.3 Isolation of Ramariolides A–D Given the hurdles associated with studies of the natural products chemistry of mushrooms, our collaborators Dr. Vivian Miao working in the Davies laboratory at UBC, and Terry Taylor, an amateur mycologist, in conjunction with the Vancouver Mycological Society, amassed a collection of uncultivated native western British Columbian mushroom extracts.  Dr. Santiago Ramon-Garcia, from the Thompson lab, then screened extracts of these samples for antimycobacterial activity against the Mtb model organism M. smegmatis. A MeOH extract of the coral mushroom R. cystidiophora showed promising antibacterial effects against M. smegmatis. The isolation and detailed description of the structure elucidation are presented below for the antimycobacterial constituents of the crude extract.72 Approximately 110 g fresh weight of mushroom was extracted three times with MeOH. The combined extracts were concentrated under reduced pressure at room temperature to yield an orange solid, which was partitioned between 200 mL of H2O and EtOAc. The EtOAc was concentrated in vacuo to give 0.27 g of residue, which was then chromatographed on Sephadex LH20 (3 cm x 95 cm) (eluent: 1:1 CH2Cl2/MeOH), yielding four fractions. The fourth fraction, containing the compounds of interest (monitored by TLC and anti-M. smegmatis activity) was subjected to C18 and C8 reversed-phase HPLC yielding 2.18 (10 mg), 2.19 (0.5 mg), 2.20 (0.3 mg), and 2.21 (0.1 mg).  Figure 2.7 Structures of ramariolides A–D. OO OHOOO OHOOO OHOO OH2.182.192.202.21  25 2.3.1 Structure Elucidation of Ramariolide A Ramariolide A (2.18, Figure 2.7) was isolated as an optically active white crystalline solid that gave a [M-H]– ion in the HRESIMS at m/z 281.1769, appropriate for a molecular formula of C16H26O4 requiring four sites of unsaturation. The 13C NMR spectrum contained 16 resolved carbon resonances in agreement with the HRMS measurement, and the DEPT 135 and gHSQC spectra identified 25 hydrogens attached to carbon (2 x C, 4 x CH, 9 x CH2, and 1 x CH3), indicating that there was one exchangeable hydrogen atom. Resonances in the 13C NMR spectrum could be assigned to a disubstituted alkene [δ 126.7 (C-2) and δ 149.6 (C-3)] and an unsaturated ester or carboxylic acid carbonyl [δ 168.9 (C-1)]. The absence of 13C NMR evidence for additional unsaturated functionalities indicated that 2.18 was bicyclic (Figure 2.7).    Figure 2.8 1H NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in methylene chloride-d2.   1.52.02.53.03.54.04.55.05.56.06.57.07.5 ppm  26  Figure 2.9 13C NMR spectrum of ramariolide A (2.18) recorded at 150 MHz in methylene chloride-d2.   Figure 2.10 Selected 2D NMR correlations for ramariolide A (2.18). Fragments A, B and C were constructed using gHMBC and gCOSY data. A pair of olefinic methine doublets at δ 6.34 (H-2, J = 5.6 Hz) and δ 7.55 (H-3, J = 5.6 Hz) in the 1H NMR spectrum showed gCOSY correlations to each other and gHMBC correlations to the carbonyl resonance at δ 168.9 (C-1) and a ketal resonance at δ 90.3 (C-4), consistent with Fragment A shown in Figure 2.10. 1H NMR resonances at δ 3.75 (H-5) and δ 4.05 (H-6), that showed gHSQC correlations to carbon resonances at δ 64.3 (C-5) and δ 67.9 (C-6), respectively, were assigned to a pair of oxymethine carbons. The H-6 resonance at δ 4.05 showed gCOSY correlations to the H-5 resonance at δ 3.75, to an exchangeable resonance at δ 1.99 (OH-6), and to a pair of geminal methylene resonances at δ 1.63 (H-7) and δ 1.47 (H-7’) as indicated in Fragment B in Figure 2.10. gHMBC correlations observed between H-5 (δ 3.75) and 1520253035404550556065707580859095100105110115120125130135140145150155160165170175 ppmOOHH • OO • ••14A • O •H •O HH5 6 7 8B •OOO HH OHHH 1 4 53 6C COSYHMBC2 3  27 C-6 (δ 67.9), between OH-6 (δ 1.99) and both C-5 (δ 64.3) and C-6 (δ 67.9), and between H-6 (δ 4.05) and C- 7 (δ 34.0) and C-8 (δ 32.0) supported the structure assigned to Fragment B. gHMBC correlations observed between both H-5 (δ 3.75) and H-6 (δ 4.05) and the ketal carbon C-4 (δ 90.3) showed that Fragment A was linked to Fragment B via a C-4/C-5 bond to give expanded Fragment C (Figure 2.10). A C-1 to C-4 butenolide and an oxirane linkage between the oxymethine C-5 and the C-4 ketal provided the two rings required by the molecular formula. The remaining fragment of ramariolide A (2.18, Figure 2.7) had to be saturated and account for C8H17. Only one methyl resonance, a triplet at δ 0.88 (Me-16), was observed in the 1H NMR spectrum (Figure 2.8) of 2.18, indicating that C-8 had to be one terminus of a nine carbon linear alkyl chain attached to C-7 to complete the structure. In agreement with this assignment, the H-7/H-7’ methylene resonances (δ 1.47 and 1.63) showed gHMBC correlations to C-8 (δ 25.6) and to one member (C-9) of a complex of resonances between δ 29.9 and 30.1, all having chemical shifts typical of the methylene carbons of a saturated linear alkyl chain.  Ramariolide A (2.18, Figure 2.7) gave crystals from CH2Cl2 that were subjected to single crystal x-ray diffraction analysis. An ORTEP diagram generated from the analysis is shown in Figure 2.11. Due to the weakly scattering crystal, an unrestrained anisotropic refinement of the structure was not possible; however the atomic connectivity, particularly with respect to the rigid spiro oxirane butenolide fragment of the molecule, is not in doubt. This confirmed the constitution determined by the NMR analysis outlined above and also provided the 4R*, 5R*, 6R* relative configurations shown in the ORTEP diagram (Figure 2.11). A Mosher ester analysis of 2.18 revealed that C-6 had the S absolute configuration and, therefore, ramariolide A (2.18, Figure 2.7) has the 4S, 5S, 6S absolute configuration.9  Figure 2.11 ORTEP diagram for ramariolide A (2.18).   28 Table 2.1 NMR data for ramariolide A (2.18) and ramariolide B (2.19).  ramariolide A (2.18) a ramariolide B (2.19) b position δH (J in Hz) δc, type δH (J in Hz) δc 1  168.9, C  168.0 2 6.34, d (5.6) 126.7, CH 5.37, d (5.6) 124.5 3 7.55, d (5.6) 149.7, CH 6.06, d (5.6) 148.8 4  90.4, C  112.7 5 3.77, d (4.1) 65.5, CH 3.81, dd  (11.0, 6.1) 74.5 6 4.05, m 67.9, CH 4.33, q (6.7) 90.7 7 1.62,1.49, m 34.1, CH2 1.47, 1.38, bm 34.9 8 1.37, bm 25.6, CH2 1.18, bm 24.9 9 to 13 1.27, bm 30.1, CH2 30.1, CH2 30.0, CH2 30.0, CH2 29.9, CH2 1.50, bm  29.9, 29.8, 29.8, 29.6, 29.5 14 1.25, m 32.4, CH2 1.29, bm 23.0 15 1.27, m 23.2, CH2 1.28, bm 32.2 16 0.88, t (7.2) 14.4, CH3 0.93, t (7.2) 14.9 OH-6 1.26, bm    OH-5   2.12, d (12.3)  a Recorded in methylene chloride-d2 (1H 600 MHz, 13C 150 MHz) b Recorded in benzene-d6 (1H 600 MHz, 13C 150 MHz) 2.3.2 Structure Elucidation of Ramariolide B  Ramariolide B (2.19, Figure 2.7) was isolated as an optically active oil that gave a [M+Na]+ ion in the HRESIMS at m/z 305.1721 appropriate for a molecular formula of C16H26O4 (calcd 305.1729), indicating it was an isomer of 2.18 (Figure 2.7). The 13C NMR spectrum of 2.19 contained 16 resolved resonances (2 x C, 4 x CH, 9 x CH2, 1 x CH3) in agreement with the HRMS measurement, and the gHSQC spectrum identified 25 hydrogen atoms attached to carbon, requiring one exchangeable hydrogen atom as in 2.18. The 1H (Figure 2.13) and 13C (Figure 2.14) NMR spectra obtained for ramariolide B (2.19) showed a strong resemblance to the spectra obtained for ramariolide A. gCOSY and gHMBC correlations observed for 2.19 confirmed the presence of the same butenolide and linear C-8 to C-16 alkyl chain fragments present in A (2.18). The gCOSY data also identified a partial linear spin system consisting of vicinal oxymethine protons [δ 3.81 (H-5), δ 4.32 (H-6)] and an aliphatic methylene [δ 1.38 (H-7), 1.47 (H-7’)] similar to the H-5 to H-7/H-7’ spin system in A (2.18) (Figure 2.11). However, the ketal resonance at δ 112.0 (C-4) and oxymethine carbon resonances at δ 74.4 (C-5) and δ 90.8 (C-6),   29 and their attached proton resonances at δ 3.81 (H-5) and δ 4.32 (H-6), were shifted downfield compared with the corresponding resonances in the NMR spectra of 2.18. In addition, the exchangeable proton resonance at δ 2.11 (OH-5) showed gCOSY correlations to H-5, suggesting a hydroxy substituent at C-5 in ramariolide B instead of at C-6 as in 2.18 (Figure 2.12).  The presence of the C-5 hydroxy substituent was confirmed by the observation of gHMBC correlations between OH-5 (δ 2.11) and C-4 (δ 112.0), C-5 (δ 74.4), and C-6 (δ 90.8). An ether linkage between C-6 and C-4 (Figure 2.12), which formed an oxetane in 2.19, in place of the oxirane in A (2.18, Figure 2.7), provided the second ring required by the molecular formula.  The change in ring size accounts for the downfield trend in chemical shifts of the 1H and 13C resonances assigned to the C-4 to C-6 region of 2.19 (Figure 2.7) compared with the corresponding assignments for ramariolide A (2.18).   Figure 2.12 Relevant COSY and HMBC correlations for ramariolide B (2.19).  Figure 2.131H NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. OOOHH •HHO H12 3 4 56 7 COSYHMBC0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm  30  Figure 2.14 13C NMR spectrum of ramariolide B (2.19) recorded at 150 MHz in benzene-d6  A strong 1D NOESY correlation observed between H-3 (δ 5.37) and H-5 (δ 3.81) showed that C-3 and H-5 were on the same face of the oxetane ring in 2.19 (Figure 2.15). A weak 1D NOESY signal was observed between H-5 (δ 3.81) and H-6 (δ 4.32), but there was no NOE between H-3 (δ 5.37) and H-6 (Figure 2.15). Examination of molecular models revealed the H-5 and H-6 can show an NOE regardless of whether they are on the same or opposite sides of the oxetane ring, but the absence of a H-3/H-6 NOE indicated that C-3 and H-6 must be on opposite faces as shown in the 4S*, 5S*, 6S* relative configuration assigned in ramariolide B (2.19, Figure 2.7).  We have assumed that the absolute configurations at C-6 and C-5 are identical in ramariolides A (2.18) and B (2.19) (Figure 2.7). This assumption, which is consistent with the syn C-3/H-5 and anti C-3/H-6 relative configurations assigned above, gives the 4S, 5S, 6S absolute configuration for ramariolide B (2.19) as shown in Figure 2.15.    101520253035404550556065707580859095100105110115120125130135140145150155160165170175 ppm  31    Figure 2.15 1D NOESY NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. 2.3.3 Structure Elucidation of Ramariolide C Ramariolide C (2.20, Figure 2.7) was isolated as an optically active white solid that gave a [M-H]– ion in the HRESIMS at m/z 265.1802 appropriate for a molecular formula of C16H26O3 (calcd 265.1804), requiring four sites of unsaturation.  Extensive analysis of the 1D and 2D NMR data revealed many of the same structural features seen in ramariolide A (2.18, Figure 2.7), with some intriguing differences.  The lactone (C-1, δ 168.9) with α, β-unsaturation (C-2, δ 121.3 and C-3, δ 140.4) was present, but the UV λmax (273 nm) was higher than observed for A (2.18) and B (2.19).  gHMBC correlations observed between both H-2 (δ 5.54) and H-3 (δ 6.88) and an olefinic carbon resonance at δ 150.6, assigned to C-4, and between H-3 and a resonance at δ 117.0, assigned to C-5, identified a Δ4,5 alkene.  The extended conjugation agreed with the bathochromic shift in the UV data. gCOSY correlations were observed between an oxymethine resonance at δ 3.81 (H-6) and H-5 (δ 5.29), a doublet at δ 0.69 (with no carbon correlation in the gHSQC), and a pair of diastereotopic methylene proton resonances at δ 1.30 (H-7) and 1.23 (H-7’). These gCOSY correlations identified a secondary alcohol at C-6 and an aliphatic methylene at C-7 as in ramariolide A (2.18) and B (2.19) (Figure 2.7). The remainder of the NMR data was consistent with a saturated linear 10-carbon alkyl chain extending from C-7 to C-OO OHO 2.19H HH NOEIrradiated H-3 H-2 H-5   32 16. A tROESY correlation between H-6 (δ 3.81) and H-3 (δ 6.88) showed that the Δ4,5 alkene had the E configuration. We have assumed that C-6 in ramariolide C (2.20, Figure 2.16) has the S configuration as in ramariolide A (2.18, Figure 2.7).    Figure 2.16 1H NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6.  Figure 2.17 13C NMR spectrum of ramariolide C (2.20) recorded at 150 MHz in benzene-d6. OO OH 2.201520253035404550556065707580859095100105110115120125130135140145 ppm  33 2.3.4 Structure Elucidation of Ramariolide D Ramariolide D (2.21, Figure 2.18) was isolated as an optically active white solid that gave a [M+H]+ ion in the HRESIMS at m/z 287.1633 appropriate for a molecular formula of C16H24O3 (calcd 287.1623) requiring five sites of unsaturation. Analysis of the NMR data obtained for D (2.21) showed that it differed from ramariolide C (2.20) only in the alkyl side chain. Two olefinic resonances in the 13C NMR spectrum at δ 140.4 (C-15) and δ 115.0 (C-16), that showed gHSQC correlations to an olefinic methine at δ 5.81 (H-15) and two geminal olefinic doublets at δ 5.07 and δ 5.01 (H-16/H-16’), respectively, identified a terminal Δ15,16 alkene on the 10-carbon side chain in ramariolide D (2.21). Again, we have assumed that C-6 has the S configuration in ramariolide D (2.21) as in A (2.18)(Figure 2.7). Table 2.2 NMR Data (C6D6, 1H 600 MHz, 13C 150 MHz) for ramariolides C (2.20) and D (2.21).  ramariolide C (2.20) ramariolide D (2.21) position δH (J in Hz) δc, type δH (J in Hz) δc, type 1  168.9, C  167.9, C 2 5.54 dd (4.0, 1.6) 121.3, CH 5.54 dd (5.6, 2.1) 120.2, CH 3 6.88 d (5.6) 140.4, CH 6.88 d (5.6) 139.3, CH 4  150.6, C  149.8, C 5 5.29 d (7.2, 0.8) 117.0, CH 5.29 dd (8.0, 1.5) 115.7, CH 6 3.81 bm 68.2, CH2 3.81 q (5.1) 67.2, CH2 7 1.30, 1.23 bm 38.2, CH2 1.28, 1.14 bm 37.4, CH2 8 1.03 bm 25.7, CH2 1.01 bm 24.8, CH2 9 to 13 1.55, bm  30.4, CH2 30.4, CH2 30.3, CH2 30.2, CH2 30.1, CH2 1.45, bm  29.2, CH2  29.1, CH2  29.1, CH2  28.8, CH2 13   1.35, bm 28.6, CH2 14 1.29, bm 32.5, CH2 2.02, q (7.2) 33.3, CH2 15 1.38, bm 23.5, CH2 5.81, m 138.4, CH 16 0.93, t (6.7) 14.7, CH3 5.02, d (10.3) 5.08, d (17.0) 113.7, CH2 OH-6 0.69, d (4.0)  0.68, bm     34  Figure 2.18 1H NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6.   Figure 2.19 13C NMR spectrum of ramariolide D (2.21) recorded at 150 MHz in benzene-d6.  0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppmOO OH 2.21253035404550556065707580859095100105110115120125130135140145150155160165170 ppm  35 2.4 Biological Activity of Ramariolide A  The minimum inhibitory concentration (MIC) of ramariolide A (2.18, Figure 2.7) determined by broth dilution against M. smegmatis was 8 µg/mL, and the minimum bactericidal concentration (MBC) to MIC ratio was 1, indicating that the effect was bactericidal (Figure 2.19 A). For comparison, isoniazid, currently a first-line anti-tuberculosis bactericidal antibiotic, had an MIC of 8 µg/mL and a MBC/MIC ratio of 1; spectinomycin, a bacteriostatic agent not used therapeutically for treatment of tuberculosis, had an MIC of 32 µg/mL and a MBC/MIC ratio of 2-4. When ramariolide A (2.18, Figure 2.7) was tested against several strains of M. tuberculosis (Figure 2.19 B), a small inhibition of growth (calculated IC50 = 53 µg/mL and an MIC = 64–128 µg/mL) was observed.  In addition to its anti-mycobacterial effect, ramariolide A (2.18, Figure 2.7) also inhibited growth of Micrococcus luteus and Staphylococcus aureus (inhibition zones of 18 mm and 15 mm diameter, respectively, on disk diffusion assays), but there was no effect on Gram-negative tester strains. Figure 2.20A shows the resazurin assay for activity against M. smegmatis. The pink/purple wells indicate live cells while the blue wells indicate dead cells. Two-fold serial dilutions were performed from top to bottom. The highest concentration tested for 2.18 was 128 µg/mL. Isoniazid (INH) and spectinomycin (SPT) were included for comparison at maximum concentrations of 64 µg/mL. Control wells contained untreated cells (CT+) or media (CT-) only.  Figure 2.20B shows the results of the MTT assay for activity against M. tuberculosis. A log2(dose)-response non-linear (four parameters) fitting curve was used to calculate the IC50. The 95% confidence interval of the IC50 was 5.161 to 6.329 (35.77 to 80.39 µg/mL). Experiments were performed in triplicate.     36  Figure 2.20 Anti-mycobacterial activity of ramariolide A (2.18).  2.5 Conclusions  Ramariolides A–D (2.18–2.21, Figure 2.7) represent a novel carbon chain length for unbranched butenolides.  The most similar compounds to the ramariolides in the literature are the hygrophorones F (2.22) and G (2.23)(Figure 2.21),73 which resemble ramariolide C (2.20) in structure, but contain two additional methylenes in the alkyl chain. The spiro oxirane-butenolide and spiro oxetane-butenolide moieties in ramariolide A (2.18) and B (2.19) (Figure 2.7) represent novel functionalities for polyketide-derived natural products. The spiro-oxetane-butenolide in ramariolide B (2.19) has only been observed in nature once before in the sesquiterpenoid parthexetine (2.24, Figure 2.21).74 Given their unique structures, proposed mechanism of action studies on the ramariolides may provide information on a novel target for Mtb inhibition. The results shown above suggest the Ramaria genus merits further investigation as a source of novel compounds with antimycobacterial activity. !"# $!% &'( )*" !"+"#$#% ! & ' ( ) * +%!%&%'%(%)%*%+%,%-%!%%!!%&!,,,-,./01,$.2/32,!4567%.89,:;<=>?7<4,,@,$!%,A,$!4567%!  37  Figure 2.21 Hygrophorone F (2.22) and G (2.23) and parthexetine (2.24).72–74 2.6 Experimental section 2.6.1 General Experimental Procedures Optical rotations were measured using a Jasco P-1010 Polarimeter with sodium light (589 nm). UV spectra were recorded with a Waters 2487 Dual λ Absorbance Detector. The 1H and 13C NMR spectra were recorded on a Bruker AV-600 spectrometer with a 5 mm CPTCI cryoprobe. 1H NMR chemical shifts are referenced to the residual benzene-d6, dichloromethane-d2, or chloroform-d3 signal (δ 7.15, 5.32, 7.26, respectively) and 13C NMR chemical shifts are referenced to the benzene-d6 or dichloromethane-d2 solvent peak (δ 128.0 or 53.8, respectively). Low and high resolution ESI-QIT-MS were recorded on a Bruker-Hewlett Packard 1100 Esquire–LC system mass spectrometer. Merck Type 5554 silica gel plates and Whatman MKC18F plates were used for analytical thin layer chromatography. Reversed-phase HPLC purifications were performed on a Waters 1525 Binary HPLC Pump attached to a Waters 2487 Dual λ Absorbance Detector.  All solvents used for HPLC were Fisher HPLC grade. 2.6.2 Mushroom Sample The source specimen, W179, was a display sample used in the 2009 Vancouver Mycological Society Fall Mushroom show in which members collected examples of local (southwestern British Columbia) fungi for exhibition to the public. The sample was donated for the purpose of this study, and identified specifically as Ramaria cystidiophora using morphological67 and genetic criteria. For the latter, DNA was extracted using standard OO HHO C 11 H22 OO HO C 11 H22HOO OHOOAcOAc2.22 2.232.24  38 procedures, and the ITS75 region between the 18S and 28S ribosomal RNA genes was amplified by PCR and sequenced.  When compared to publicly available accessions in Genbank, the 511 nucleotides obtained were observed to be identical to accessions EU597077 (nt 34-544) and DQ384590 (nt 79-589), both derived from samples originating in southwestern British Columbia, and previously identified as R. cystidiophora. A voucher sample of W179 has been deposited in the herbarium (UBC) at the University of British Columbia. 2.6.3 Extraction and Isolation   Approximately 110 g fresh weight of mushroom was cut into pieces and extracted three times with MeOH (50 mL). The combined extracts were concentrated in vacuo at room temperature to yield an orange solid, which was partitioned between 200 mL each of H2O and EtOAc.  The EtOAc was concentrated in vacuo to give 0.27 g of residue, which was then chromatographed on Sephadex LH20 (3 cm x 95 cm) yielding four fractions. The fourth fraction, containing the compounds of interest (monitored by TLC), was subjected to C18 reversed-phase HPLC using a CSC-Inertsil 150Å/ODS2, 5 µm 25 x 0.94 cm column, with 4:5 ACN/H2O as eluent to yield all 4 compounds in a nearly pure state (retention times of 2.18-19.5 min, 2.19-17 min, 2.20-18.5 min, and 2.21-14 min). Final purification was done using C8 reversed-phase HPLC using Phenomenex Luna C8100 Å, 5 µm 250 X 10 mm column with 7:10 ACN/H2O as eluent (retention times: 2.18-27.5 min, 2.19-26 min, 2.20-25.5 min, and 2.21-19 min), yielding 2.18 (10 mg), 2.19 (0.5 mg), 2.20 (0.3 mg), and 2.21 (0.1 mg).  Ramariolide A (2.18): irregular colorless crystals (CH2Cl2); mp 92–94° C; [α]25D  -42 (c 5.4, MeOH); UV (MeOH) λmax (log ε) 213 (2008) nm; 1H and 13C NMR see Table 1; negative ion HRESIMS [M-H]- m/z 281.1769 (calcd. for C16H25O4, 281.1753). Ramariolide B (2.19): clear oil; [α]25D -50 (c 0.3, MeOH); UV (MeOH) λmax (log ε) 203 (1476) nm; 1H and 13C NMR see Table 1; positive ion HRESIMS [M+Na]+ m/z 305.1721 (calcd. for C16H26O4Na, 305.1729). Ramariolide C (2.20): clear amorphous oil; [α]25D +85 (c 0.2, MeOH); UV (MeOH) λmax (log ε) 273 (1332) nm; 1H and 13C NMR see Table 2; negative ion HRESIMS [M-H]- m/z 265.1802 (calcd. for C16H25O3, 265.1804).   39 Ramariolide D (2.21): clear amorphous solid; [α]25D +42 (c 0.05, MeOH); UV (MeOH) λmax (log ε) 273 (2332) nm; 1H and 13C NMR see Table 2; positive ion HRESIMS [M+H]+ m/z 287.1633 (calcd. for C16H24O3Na, 287.1623 ). 2.6.4 Mosher’s Analysis of Ramariolide A76  Figure 2.22 Mosher’s Analysis: Showing Δδ(δS-δR) values from Mosher’s ester analysis of ramariolide A76 (2.18). 2.6.4.1 Synthesis of MTPA Esters of Ramariolide A Synthesis of (S)-(S)-1-((2S, 3S)-5-oxo-1, 4-dioxaspiro[2.4]hept-6-en-2-yl)undecyl 3, 3, 3-trifluoro-2-methoxy-2-phenylpropanoate (2.25)  To a stirred solution of ramariolide A (2.18, 0.5 mg, 0.002 mmol) in 1 mL of dried dichloromethane was added a catalytic amount of DMAP and pyridine.  Then approximately 10 µL (0.05 mmol) of (R)-MTPACl were added and the reaction was stirred for 5 hours.  The reaction mixture was subsequently concentrated and purified using C8 reversed-phase HPLC.  The product eluted in ACN (retention time: 10 min) giving 0.2 mg (20%) of the desired product as a clear solid. 1H NMR (600 MHz, CDCl3) δ 7.56 (m, 2H), 7.45 (m, 3H), 7.35 (d, J = 5.6 Hz, 1H), 6.08 (d, J = 5.1 Hz, 1H), 4.22 (d, J = 4.1 Hz, 1H), 5.41 (m, 1H), 3.53 (s, 3H), 1.77 (m, 1H), 1.71 (m, 1H), 1.26 (m, 16H), 0.89 (t, J = 6.7 Hz, 3H); negative ion HRESIMS [M-H]– m/z 497.2148 (calcd. for C26H32O6F3, 497.2151). 357 +0.41-0.12-0.11 +0.112.25  R = (R)-MTPA2.26 R = (S)-MTPAO ORO OHOO OO O OMePhCF32.25  40  Figure 2.23 1H NMR spectrum of ramariolide A MTPA-R ester (2.25) recorded at 600 MHz in CDCl3 (R)-(S)-1-((2S, 3S)-5-oxo-1,4-dioxaspiro[2.4]hept-6-en-2-yl)undecyl 3, 3, 3-trifluoro-2-methoxy-2-phenylpropanoate (2.26).    Procedure same as above substituting (S)-MTPACl.  Yield 0.4 mg (40%). 1H NMR (600 MHz, CDCl3) δ 7.57 (m, 2H), 7.45 (m, 3H), 6.94 (d, J = 5.6 Hz, 1H), 6.01 (d, J = 5.6 Hz, 1H), 5.39 (pent, J = 4.1 Hz, 1H), 4.11 (d, J = 5.1 Hz, 1H), 3.58 (s, 3H), 1.85 (m, 1H), 1.79 (m, 1H), 1.28 (bm, 16H), 0.89 (t, J = 6.7 Hz, 3H); negative ion HRESIMS [M-H]– m/z 497.2146 (calcd. for C26H32O6F3, 497.2151). 1.52.02.53.03.54.04.55.05.56.06.57.07.5 ppmOO OO O OMeCF 3Ph2.26  41  Figure 2.24 1H NMR spectrum of ramariolide A MTPA-S (2.26) ester recorded at 600 MHz in CDCl3. 2.6.5 2D NMR Spectra of Ramariolide A–D  Figure 2.25 gCOSY NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in CD2Cl2. 1.52.02.53.03.54.04.55.05.56.06.57.07.5 ppmppm1.01.52.02.53.03.54.04.55.05.56.06.57.07.5 ppm1.01.52.02.53.03.54.04.55.05.56.06.57.07.5  42   Figure 2.26 gHSQC NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in CD2Cl2.   Figure 2.27 gHMBC NMR spectrum of ramariolide A (2.18) recorded at 600 MHz in CD2Cl2. ppm8.0 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 0.5 0.0 ppm−101601501401301201101009080706050403020100ppm8.0 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 0.5 0.0 ppm2030405060708090100110120130140150160170  43  Figure 2.28 DEPT 135 NMR spectrum of ramariolide A (2.18) recorded at 150 MHz in CD2Cl2.   Figure 2.29 gCOSY NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0  44  Figure 2.30 gHSQC NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6.  Figure 2.31 gHMBC NMR spectrum of ramariolide B (2.19) recorded at 600 MHz in benzene-d6. ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.07.5 ppm1601501401301201101009080706050403020100ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm2030405060708090100110120130140150160170  45  Figure 2.32 gCOSY NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6.  Figure 2.33 gHSQC NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6. ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm2030405060708090100110120130140  46  Figure 2.34 gHMBC NMR spectrum of ramariolide C (2.20) recorded at 600 MHz in benzene-d6.  Figure 2.35 gCOSY NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6. ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm2030405060708090100110120130140150160170ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.07.5 ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.07.5  47  Figure 2.36 gHSQC NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6.  Figure 2.37 gHMBC NMR spectrum of ramariolide D (2.21) recorded at 600 MHz in benzene-d6. ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.07.5 ppm102030405060708090100110120130140150ppm0.51.01.52.02.53.03.54.04.55.05.56.06.57.0 ppm2030405060708090100110120130140150160170  48 Chapter 3: Dhilirolides A–N, Meroterpenoids Produced in Culture by the Fruit-Infecting Fungus Penicillium purpurogenum Collected in Sri Lanka 3.1  Chemical Prospecting: Structure Guided Isolation of Natural Products   The following project is an example of chemical prospecting, which is the term given to isolation of novel natural products without the guidance of biological activity. This method of discovery requires NMR-guided isolation techniques, combining experience and rapid de-replication methods,77 to discover new natural products.78–80  Subsequently, these new compounds can be screened in the hopes of finding a use, medicinal or otherwise.81 Isolation of unprecedented structures can be a matter of finding novel and diverse source organisms,82, 83 and as will be shown below, this makes fungi ideal as a starting point for chemical prospecting efforts.84–86 3.2 Global Fungal Diversity Global fungal diversity is a contentious subject, with estimates ranging from 1.5 million species to recent estimates near 5 million.85, 87 These figures are based largely on extrapolation of numbers of individual fungal species isolated from plants as endophytes, saprophytes or pathogens, as well as environmental DNA sequencing. The current number of identified and recognized fungal species is approximately 100,000, with many of these being Ascomycota and Basidiomycetes.85 From the above estimates, it is apparent that there is a disconnect between the number suggested for uncollected or un-cultivatable fungi, and those already identified. The fact remains, however, that the number of species is clearly very large.85  High fungal diversity is not encountered equally throughout every biome.85 In fact, significant variations in fungal diversity have been found depending on a number of factors including annual rainfall and climate.88 By most estimates, the tropics hold the highest diversity of fungi.  In one study of fungal speciation in the tropics, 83 leaves were collected from two species of Panamanian understory plants Heisteria concinna (Olacaceae) and Ouratea lucens (Ochnaceae), and 347 genetically distinct endophytic fungal species were isolated.88 This study underscores the hyper-diversity of fungi in tropical regions, and adds weight to the high estimation for the number of global fungal species.    49 3.2.1 Fungal Diversity in Sri Lanka Endemic organisms are frequently encountered on islands such as Sri Lanka, which are geographically isolated, and have a large landmass.89, 90 Fungi are subject to the same specialization and selection pressures that both plants and animals undergo on an island. Adding to these pressures is the fact that Sri Lanka enjoys a tropical climate, offering a near ideal habitat for a diverse endemic fungal flora.90 3.3 The Relationship Between Fungi and Plants  Many fungi are known to live within plant cells.91–94 The relationships between plants and fungi run the gamut from transient saprophyte, to mutualistic, or even parasitic.95, 96 Fungal endophytes are classified as species that live within the tissue of a plant, but appear not to harm the host. This relationship has been described as a “teetering scale”, where if the plants’ defenses are compromised, or if fungal virulence is increased, the fungus can quickly become harmful and parasitic.95–98 This has led to a redefining of the term endophyte in some cases, to mean a “balanced pathogen-host antagonism.”86, 97, 98  Some plant-endophyte relationships have been shown to be purely mutualistic.86, 99 Endophytes often receive nutrients and protection while the host plant benefits by increased ecological adaptability, giving the host greater resilience against herbivores or resistance to pathogens.  Compounds such as peramine (3.1, Figure 3.1) or the ergot alkaloids (e.g., 3.2, Figure 3.1), produced by several species of endophytic fungi (Neotyphodium/Epichloë complex) associated with rye grasses (Lolium spp.), are examples where endophyte-produced compounds, deter insect and mammalian predators of the plant host.99–101 The unique relationships that have developed between plants and fungi have likely driven selection for production of structurally diverse, biologically active metabolites by the fungus.  In a study comparing the biological activity of extracts from both soil fungi (~2,800) and endophytes (~6,500), it was found that 80% of extracts coming from endophytes had activity in at least one of the antibacterial, fungicidal, algicidal or herbicidal assays employed, compared with only 64% of those from soil isolates.97 Isolation of biologically active metabolites and chemical prospecting from endophytes are active areas of research.86, 97 In a recent review of the natural products produced by endophytes, every major structural class of secondary metabolites was represented. The high level of structural diversity and biological activity has cemented endophytic fungi as promising source organisms   50 for natural products discovery.84, 86 3.4 Diversity of Fungal Secondary Metabolites: The C25 Meroterpenoids Meroterpenes produced by fungi are a large group of compounds with medicinally relevant members such as Xenovulene A® (3.3, Figure 3.1), an inhibitor of benzodiazepine binding to the GABAA receptor, isolated from Acremonium strictum.102, 103 Pyripyropene A (3.4, Figure 3.1), isolated from A. fumigatus, is a potent and selective inhibitor of acyl CoA cholesterol acyltransferase (ACAT). 3.4 is now in preclinical development for the treatment of atherosclerosis.104, 105 Meroterpenoids come in several classes, differentiated by their non-terpene portion, and have a vast array of biological activities.103 One such class, is a family of C25 meroterpenoids derived from the alkylation of 3,5-dimethylorsellinic acid by farnesyl diphosphate.  The diversity, biogenesis, and biological activity of these compounds will be discussed.   Figure 3.1 Peramine and ergotamine alkaloids and meroterpenoid clinical candidates. 3.4.1 Diversity and Biosynthetic Origins of C25 Meroterpenoids From 3,5-Dimethylorsellinic acid The metabolites arising from farnesyl diphosphate (FPP) alkylation of 3,5-dimethylorsellinic acid (3.5, Figure 3.3) have been found in over 20 species of fungi from OO OHO HH HXenovulene A! (3.3)OO NOAcO HO OAcOHHOAcpyripyropene A (3.4)HN NHO NH NO NOHO OHergotamine (3.2)N NO NH NH2NHperamine (3.1)  51 primarily two genera, Aspergillus and Penicillium.103 The compounds that arise are similar in their initial starting materials, however, their final products are often oxidized or rearranged to an almost untraceable state. Examples of this can be seen in the anditomins and andibenins (Figure 3.2) isolated from A. variecolor. In these molecules, the 3,5-dimethylorsellinic acid (3.5, Figure 3.3) portion of the molecule has been degraded to a near unrecognizable state, with only carbon number giving some hint as to its origin.106, 107 Another interesting variation on the theme is novo-fumigatonin (Figure 3.2), isolated from A. fumigatus, which has had several bonds cleaved and rearranged to arrive at the final carbon skeleton.108 In Figure 3.2, representatives from each of the previously reported novel carbon skeletons produced by this pathway are shown. In 1976, the first member of this class of compounds, andibenin, was isolated from Aspergillus variecolor.107 At that time the natural product was thought to be derived from a sesterterpenoid biosynthetic origin. In 1978, Simpson et al. performed the first of many isotopic labeling studies using 13C doubly labeled acetate and 18O gas for incorporation into this diverse group of compounds, confirming a meroterpenoid biosynthetic origin.103, 104, 113, 120, 134  In 2011, the biochemical techniques were in place to finally sequence and annotate the biosynthetic genes in Aspergillus nidulans responsible for the production of austinol.28 The study by Hsien-Chun Lo et al. showed that the polyketide, 3,5-dimethylorsellinic acid (3.5), was derived from a non-reducing polyketide synthase (NR-PKS) named ausA, on a separate chromosome from a prenyltransferase, named ausN, responsible for farnesyl diphosphate alkylation.104 This was followed by a methyltransferase (ausD), a flavin-dependent epoxidase (ausM), a polyene cyclase (ausL) and a monooxygenase (ausB) to arrive at berkeleyone A (3.6, Figure 3.3), a proposed intermediate common to several families of compounds. 103, 104, 121 In a second paper by Matsuda et al., the oxidation, Baeyer-Villiger lactonization and spiro-ring formation seen in the final product, austinol (3.7, Figure 3.3), were determined as enzymatic products of ausB, ausE and ausC.121 The final rearrangements seen in austinol (3.7) were attributed to the genes ausJ, ausK, ausH, ausI, ausG and ausL. However, the authors conclude that other genes may be necessary and certain transformations may happen without the need for enzyme catalysis.104 Shortly after, the biosynthesis of andrastin (3.8, Figure 3.3) was described from nine biosynthetic genes in P. chrysogenum, as well as the biosynthesis of terretonin from A. terreus.122, 123   52  Figure 3.2 Representatives of each novel fungal C25 meroterpenoid carbon skeleton.103, 104, 106–119 tropolactone Aberkeleydione preaustinoid A1citreohybridone A andilesen CberkeleytrioneO OAcO OO OO OMecitreohybridone Canditominnovofumigatoninandibenin BOOO O OO OOHberkeleyacetal A miniolutelide Aandrastin C insuetolide Aaustin OO OOH OO OOMeOH HterretoninOO OOH OOHterretonin FOO OOH OOOH Hterretonin C COOMetropolactone DOO AcO O OO OHisoaustinoneOO O OHO OHOneoaustinoneOO AcO O OHO OOpreaustinoid A5emervaridione OOOO H OHO OOO HO OOOOOOOO OO OH O OO O OO OH OO OOOOHOO H OOOOO HO O O OO OO OOOOOO H OO OHAcO CO2MeH O OAcAcO CO2MeH OOOO O OHOH CO2Me OHO OOOO OCO2MeHO OHHO OH CO2Me OHOO O OH CO2Me OHO OHH H  53  Figure 3.3 Biosynthetic route overview to C25 meroterpenoidsAdapted from 103, 123–126 3.4.2 Biological Activity  The energy devoted by the producing fungi to the biosynthesis of these metabolites likely makes them important molecules in their respective niches.97 However, the exact utility to the fungi remains a mystery, except in the case of dihydroaustinol.  Recently, it was shown that dihydroaustinol, in conjunction with diorcinol, acted as a signaling substance that prompts sporulation in non-sporulating colonies of A. nidulans.127 This finding suggests that the C25 meroterpenoids may act as endogenous signaling molecules within or between fungi.   Many members of this family of compounds have shown biological activity either discovered by chemical prospecting, or bioassay guided fractionation. 109, 113, 117 Austin, for instance, presents potent insecticidal activity, shown by Kataoka et al. to be due to selective antagonism of nicotinic acetylcholine receptors in the insect neurons.128 Dehydroaustin was also shown to be toxic to insects, controlling Aedes aegypti, a common mosquito species implicated as a disease vector.129 Another insecticidal compound in this family, citreohybridonol from P. OH OHCOOHHO OCOOHHO COOHO O HOOO COOHOH HO O OHCOOHO OHO CO2Me austinol ( 3.7 )andrastinscitreohybridonesatlantinoneterretoninpenisimplicinemervaridioneanditominsandilesinsandibeninsinseutolidetropolactonesfumigatonins3,5-dimethylorsellinic acid ( 3.5 )AdrI HO COOHO O ausEausCausLausB OHberkeleyone A ( 3.6 ) berkeleyonesberkeleyacetalsparaherquonindhilirolides ausH-LausG O OOO HO OOaustinsHHandrastin E ( 3.8 ) HHFPP  54 citreo-viride, was shown to be a potent anti-feedant against the moth Plutella xylostella, a known European agricultural pest.130   The biological activities found have also included modulation of medicinally relevant targets. For instance, berkeleydione and berkeleytrione isolated from P. rubrum, are inhibitors of metalloproteinase-3 and caspase-1. The structurally related berkeleyacetals have been shown to be active against non-small-cell lung carcinoma NCI H460.114 The tropolactones A–C (Figure 3.2), isolated by the Fenical group from a marine Aspergillus sp., have shown anticancer activity against human colon carcinoma (HCT-116) with IC50 values of 13.2, 10.9 and 13.9 µg/mL respectively.118 Also, the andrastins (Figure 3.2) were found to be inhibitors of protein farnesyl transferase.119, 131 Several members of this class, however, were originally published without any indication of biological activity.109, 113, 117 Given the versatility of the biosynthetic machinery and complexity of structures generated, this family is likely to garner continued research interest. 3.5 Isolation of Dhilirolides A–N  An isolate of P. purpurogenum was obtained from infected A. bilimbi fruit collected in Nugegoda, Sri Lanka by our Sri Lankan collaborators E. Dilip DeSilva, Ravi L. C. Wijesundera and Dinith R. Jayanetti. Laboratory cultures of P. purpurogenum were grown on the surface of potato dextrose agar in Petri dishes for 5 days at room temperature. A deep red pigment produced by the fungus was excreted into the agar during the culture period, mimicking the coloration of the fruit infection. The cultures were harvested by scraping the fungal mycelium off the agar surface before cutting the agar into small pieces that were extracted three times with fresh EtOAc. The EtOAc extract was concentrated under reduced pressure and chromatographed on Sephadex LH20 (3 cm x 95 cm) using 4:1 MeOH/CH2Cl2. The fractions containing the compounds of interest were combined and subjected to C18 RPHPLC chromatography to give pure samples of dhilirolides A (3.9) (28.9 mg), B (3.10) (7.6 mg), C (3.11) (17.7 mg), D (3.12) (8.7 mg), F (3.14) (0.4 mg), J (3.18) (2.5 mg), L (3.20) (14.8 mg), M (3.21) (1.0 mg) and N (3.22) (0.5 mg). Additional fractions containing impure samples of dhilirolides E (3.13), G (3.15), H (3.16), I (3.17) and K (3.19) were purified using C8 reversed-phase HPLC to give 3.13 (0.9 mg), 3.15 (0.5 mg), 3.16 (1.0 mg), 3.17 (0.9 mg) and 3.19 (1.4 mg).   The dhilirolides were originally isolated using 1H NMR as a guide for isolation. Anti-feedant and insecticidal activity   55 against Trichoplusia ni (Lepidoptera: Noctuidae) was detected after isolation and characterization had occurred.132, 133  Figure 3.4 The dhilirolides A–N (3.9–3.22) and the dhilirane (I), isodhilirane (II), 14,15-dinordhilirane (III), and 23,24-dinorisodhilirane (IV) meroterpenoid carbon skeletons. 3.103.9 OO OOO OHOO 3.113.12 1 3 91012151617 18 2223 2425O OO OOO OHOOOO OOO OHO OHO OOO OOHO3.19 3.203.18 3.163.15 3.21 OOO OO3.17OO HO O OMeOOO OHO OAcOMeOOO OOO OHOHO OOO OOHO HOOO OOMeO OAcO OOO OOHOOOOOMeO3.13 3.144 719 81 73 85 917 10121816 15 1913 23 24252220 24OHOO OOO OOAcA B C O O13.22 OOO OOO O17 14 9O 5 917212020 21HOOHI II III IV1 35 7161718 19 82021 229 2324101213 15 25 1 35 7171618 819 20 9 23 241021 222512131514 14  56 3.6 Structure Elucidation of Dhilirolides A–N 3.6.1 Structure Elucidation of Dhilirolide A In collaboration with Dr. David E. Williams of the Andersen lab and Professor E. Dilip DeSilva, dhilirolide A (3.9, Figure 3.4) was obtained as optically active colorless crystals (mp 267–269 °C) that gave an [M+H]+ ion in the HRESIMS at m/z 473.1796 appropriate for a molecular formula of C25H28O9, requiring 12 sites of unsaturation. The 13C NMR spectrum obtained for dhilirolide A (3.9) contained 25-resolved resonances, in agreement with the HRESIMS data. A detailed analysis of the 1H/13C/gCOSY/gHSQC/gHMBC NMR data identified five methyl singlets [δ 0.74 (Me-25), 1.18 (Me- 17), 1.25 (Me-19), 1.50 (Me-18), 1.51 (Me-15)], one methyl doublet [δ 1.42 J = 7.1 Hz (Me-24)], a trisubstituted olefin [δ 6.42 (H-2), 125.0 (C-2); 152.2 (C-3)], two ester/lactone carbonyls [δ 162.0 (C-1); δ 170.3 (C-20)], a trisubstituted epoxide [δ 55.6 (C-4); 3.68 (H-5), δ 55.9 (C-5)]; a 1,1 disubstituted epoxide [δ 64.0 (C-21); δ 2.36/2.90 (H-22α/H- 22β), δ 45.6 (C-22)], two oxygenated tertiary carbons [δ 91.0 (C-9); δ 81.5 (C-16)], an oxygenated methine carbon [δ 4.73 (H-23), δ 80.9 (C-23)], two ketals [δ 105.7 (C-10); δ 107.7 (C-14)], four other quaternary carbons [δ 40.5 (C-7); δ 54.2 (C-8); δ 44.9 (C-11); δ 47.9 (C-13)], and two methylenes [δ 1.77/2.46 (H-6ax/H-6eq), δ 31.5 (C-6); δ 1.74/2.11 (H-12ax/H-12eq), δ 36.0 (C-12)]. The alkene, carbonyl, and epoxide functionalities described above accounted for five of the 12 sites of unsaturation indicated by the molecular formula, requiring that 3.9 contain seven additional rings.          57 Table 3.1 1H and 13C NMR chemical shifts for dhilirolides A–D.  A (3.9)a B (3.10)a C (3.11)b D (3.12)a Atom # δH (J in Hz) δC δH (J in Hz) δC δH (J in Hz) δC δH (J in Hz) δC 1  162.0  162.8  164.9  163.9 2 6.42 s 125.0 5.83 s 114.7 6.32 s 126.1 6.06 s 113.7 3  152.2  150.0  155.3  163.1 4  55.6  133.4  57.1  131.7 5 3.68 d (6.4) 55.9 6.15 m 129.4 3.56 d (6.3) 58.1 6.02 m 130.4 6ax/b 6eq/a 2.46 bd  (14.8) 1.77 dd  (14.8, 6.4) 31.5 2.92 bd  (19.3) 1.94 dd  (19.3, 5.8) 35.3 2.69 bd (14.7) 1.92 dd  (14.7, 6.3) 33.1 3.45 dd  (19.9, 3.4) 1.94 dd  (19.9, 6.1) 38.4 7  40.5  40.4  41.6  42.8 8  54.2  54.3  61.3  65.5 9  91.0  91.0  93.9  89.5 10  105.7  106.3  109.65  214.2 11  44.9  45.1  50.4  55.2 12ax 12eq 1.74 d (13.1) 2.11 d (13.1) 36.0 1.69 d (13.2) 1.98 d (13.2) 35.0 1.58 d (12.3) 2.21 d (12.3) 41.2 1.96 t (13.2) 1.87 dd  (13.2, 5.4) 41.6 13  47.9  47.5  49.8 2.55 dd  (13.2, 5.4) 48.8 14  107.7  107.4  109.71  74.6 15 1.51 s 21.091 1.48 s 20.3 1.57 s 21.9 1.18 s 32.9 16  81.5  82.0  83.3  82.7 17 1.18 s 21.131 1.50 s 29.4 1.26 s 21.8 1.38 s 26.5 18 1.50 s 26.4 1.54 s 27.8 1.54 s 27.0 1.53 s 26.1 19 1.25 bs 21.071 1.16 s 20.3 1.15 d (1.3) 21.7 1.08 s 15.9 20  170.3  170.3  173.9  172.6 21  64.0  64.0  151.7  148.1 22a 22b 2.90 d (3.7) 2.36 d (3.7) 45.6 2.92 d (3.7) 2.40 d (3.7) 45.8       5.11 s 5.04 s 107.7      5.15 s      4.92 s 107.3 23 4.73 q (7.1) 80.9 4.72 q (7.0) 80.8 4.69 q (7.0) 83.2 4.89 q (7.1) 83.1 24 1.42 d (7.1) 18.1 1.43 d (7.0) 18.1 1.42 d (7.0) 19.2 1.02 d (7.1) 17.8 25 0.74 s 13.2 0.76 s 13.3 1.17 s 16.1 1.15 s 15.4 9-OH /  /  /  7.22 s  10-OH 7.39 s  7.43 s  not observed  /  14-OH /  /  /  5.22 s   aRecorded in DMSO-d6  bRecorded in MeOH-d4  1These assignments are interchangeable     58  Figure 3.5 1H NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6 showing numbering.  Figure 3.6 13C NMR spectrum of dhilirolide A (3.9) recorded at 150 MHz in DMSO-d6.  8.5 8.0 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 0.5Chemical Shift (ppm)10-OH 2 23 5 22 a 22 b6ax 12 ax6 eq12 ax15 182419 17 25!"1 3 20 2 14 10 9 16 23 5 21 8 4 13 22 11 7 12 6 18 19 15 24 25   59 Only one proton resonance (δ 7.39 s, OH-10) did not correlate to a carbon resonance in the gHSQC spectrum of 3.9. This OH-10 resonance correlated to the ketal carbon at δ 105.7 (C-10) in the gHMBC experiment, consistent with the presence of a hemiketal. The two esters, two epoxides, and one OH accounted for seven of the nine oxygen atoms in the molecular formula. Since there were no further exchangeable protons or carbonyls, the remaining two oxygen atoms had to be present as ether linkages. Analysis of the gCOSY and gHMBC data as illustrated in Figure 3.7 readily established most of the constitution of dhilirolide A (3.9). Of particular note, were three weak 4-bond gHMBC correlations observed between Me-17 (δ 1.18) and C-1 (δ 162.0), and between Me-15 (δ 1.51) and both C-9 (δ 91.0) and C-10 (δ 105.7), that established the ester linkage between C-1 and C-16 and the ether linkages between the C-14 (δ 107.7) ketal carbon and both the C-9 oxygenated tertiary carbon and the C-10 hemiketal carbon. The gHMBC and gCOSY data failed to identify the final substituents on either C-8 or C-20 and one site of unsaturation. Simply linking the unsatisfied valences on C-8 and C-20 generated the lactone that provided the final ring required by the molecular formula, completing the constitution of dhilirolide A (3.9, Figure 3.4). The rigidity of 3.9, resulting from its fused polycyclic structure, facilitated the assignment of the complete relative configuration via analysis of the observed tROESY correlations as shown in Figure 3.10.     Figure 3.7 Selected gHMBC and gCOSY 60 correlations for dhilirolide A (3.9).   (! 1.50)(! 1.18)1817 OO O H2H6ax H6eq(! 2.46) (! 1.77)H5(! 3.68) (! 6.42)4(! 55.6)1(! 162.0) 3(! 152.2)16 (! 91)(! 105.7)(! 107.7)15 14 1091213 11 25242320197 8 21O O OOHO O 22 HMBCCOSY  60  Figure 3.8 gHMBC spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6.  Figure 3.9 gCOSY 60 NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6.   61    Figure 3.10 Selected tROESY correlations of dhilirolide A (3.9)    Figure 3.11 tROESY NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6. The structure proposed for dhilirolide A (3.9, Figure 3.4) from analysis of its NMR data has a complex, unprecedented carbon skeleton. A single crystal of dhilirolide A (3.9) was subjected by Brian O. Patrick to single crystal X-ray diffraction analysis in order to verify the proposed structure. The material crystallizes with four crystallographically independent molecules of dhilirolide A (3.9) in the asymmetric unit. The absolute configuration was established on the basis of the refined Flack x-parameter [x = -0.06(11)].134, 135 Additionally, the Me15 OO OH10O Me24OMe19 Me25O O O H6eqH6axH5Me Me18 O H22!H22"H12axH23ROESY  62 material crystallizes as a two component twin, with twin domains related by a 180° rotation about the [-1 1 0] reciprocal lattice axis. The ratio of major-to-minor twin components is 0.84:0.16. The ORTEP diagram shown in Figure 3.12 shows the proposed constitution of dhilirolide A (3.9) with its absolute configuration as 4S, 5R, 7R, 8R, 9R, 10R, 11R, 13S, 14S, 21R, and 23S.   Figure 3.12 ORTEP diagram of dhilirolide A (3.9) 3.6.2  Structure Elucidation of Dhilirolide B In collaboration with Dr. David E. Williams of the Andersen lab and Professor E. Dilip DeSilva, dhilirolide B (3.10, Figure 3.4) was isolated as an optically active amorphous solid that gave an [M+Na]+ ion at m/z 479.1725 in the HRESIMS appropriate for the molecular formula of C25H28O8, that differs from that of 3.9 simply by the loss of an oxygen atom. Although the 1H and 13C NMR spectra of dhilirolide B (3.10) were similar to those of dhilirolide A, the UV spectrum was markedly different. In dhilirolide A (3.9, Figure 3.4), a λmax typical of a trisubstituted α,β-unsaturated lactone was observed at 235 nm, while in 3.10, the λmax was shifted to 280 nm. Replacing the C-4/C-5 epoxide in 3.9 with a double bond in 3.10 would account for the loss of an oxygen and extend the conjugation from the enone in 3.9 to a dienone in 3.10, satisfying the λmax observed for 3.10. Examination of the 1D and 2D NMR data obtained for 3.10 revealed that the epoxide H-5 resonance (δ 3.68) in dhilirolide A (3.9) had been replaced by an olefinic resonance at δ 6.15 that showed gCOSY correlations to the methylene resonances at δ   63 1.94 and 2.92 assigned to H-6eq/H-6ax. gHMBC correlations between C-4 (δ 133.4) and H-2 (δ 5.83), H-6eq/H-6ax (δ 1.94/2.92), Me-17 (δ 1.50), and Me-18 (δ 1.54), and between H-5 (δ 6.15) and both C-3 (δ 150.0) and C-16 (δ 82.0), were consistent with the proposed structure.  Figure 3.13 1H NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6.  Figure 3.14 13C NMR spectrum of dhilirolide B (3.10) recorded at 150 MHz in DMSO-d6. 3.6.3 Structure Elucidation of Dhilirolide C In collaboration with Dr. David E. Williams of the Andersen lab and Professor E. Dilip DeSilva, dhilirolide C (3.11, Figure 3.4) was isolated as an optically active amorphous solid that   64 gave an [M+Na]+ ion at m/z 479.1746 in the HRESIMS consistent with the molecular formula C25H28O8, identical to the molecular formula of dhilirolide B (3.10). The UV spectrum observed for 3 (λmax 239 nm) was similar to that of dhilirolide A (3.9) (λmax 235 nm) indicating that it did not have the dienone moiety found in 3.10. Comparison of the 1H and 13C NMR spectrum of dhilirolide C (3.11) with the corresponding spectra recorded for dhilirolide A (3.9, Figure 3.4) revealed that the resonances assigned to the 1,1-disubstituted C-21/C-22 epoxide in 3.9 were absent. Instead, the 1H NMR spectrum of 3.11 contained two new singlet resonances at δ 5.04 (H-22α) and δ 5.11 (H-22β), which both correlated to an olefinic carbon at δ 107.7 (C-22) in the gHSQC spectrum and to a second olefinic carbon at δ 151.7 (C-21) in the HMBC spectrum. These observations showed that 3.9 and 3.11 differed simply by the replacement of the C-21/C-22 epoxide in 3.9 with a Δ21,22 exocyclic alkene in 3.11. gHMBC correlations observed between the H-22α/ H-22β olefinic methylene resonances at δ 5.04 and δ 5.11 and the methine carbons at δ 61.3 and δ 50.4, assigned to C-8 and C-11, respectively, supported this assignment. Interestingly, although all the other structural features of 3.9 and 3.11 were identical, it was found that when the 1H NMR spectrum of 3.11 was recorded in DMSO-d6 many resonances were doubled or broadened and significantly shifted. This phenomenon was attributed to a slow conformational equilibrium. A single set of well-resolved resonances could be obtained when the NMR spectra for 3.11 (Figure 3.4) were recorded in MeOH-d4.    65 Figure 3.15 1H NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4. Figure 3.16 13C NMR spectrum of dhilirolide C (3.11) recorded at 150 MHz in MeOD-d4   66 3.6.4 Structure Elucidation of Dhilirolide D In collaboration with Dr. David E. Williams of the Andersen lab and Professor E. Dilip DeSilva, dhilirolide D (3.12, Figure 3.4) was isolated as an optically active amorphous solid that gave an [M-H]- ion at m/z 441.1967 in the HRESIMS appropriate for a molecular formula of C25H30O7. The molecular formula of 3.12 differed from the molecular formula of 3.9 by the addition of two hydrogen atoms and the loss of two oxygen atoms, and it required only 11 sites of unsaturation. Examination of the 1H and 13C NMR spectra recorded for dhilirolide D (3.12, Figure 3.4) revealed a close relationship to dhilirolides A–C (3.9–3.11, Figure 3.4), but also several significant structural and functional group differences. The UV (λmax 276 nm) and 1H/13C/COSY/HSQC/HMBC NMR data obtained for 3.12 identified the C-1 to C-5 dienone substructure present in 3.10 and the Δ21,22 exocyclic alkene present in 3.11.  Figure 3.17 1H NMR spectrum of dhilirolide D (3.12) recorded at 600 MHz in DMSO-d6. 8.0 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 0.5Chemical Shift (ppm)9-OH 25 14-OH22 a 22 b 23 6 ax 13 6 eq 12 ax12 eq1817 1525 1924  67  Figure 3.18 13C NMR spectrum of dhilirolide D (3.12) recorded at 150 MHz in DMSO-d6 Significant differences in the 13C NMR data obtained for 3.12 were that the resonances assigned to the ketal carbons C-10 and C-14 and the quaternary carbon C-13 in 3.9, 3.10, and 3.11 had been replaced by a ketone resonance at δ 214.2 (C-10), a tertiary carbinol resonance at δ 74.6 (C-14), and a methine resonance at δ 48.8 (C-13) in 3.12, and the carbon resonance assigned to Me-15 had been shifted significantly downfield to δ 32.9 in 3.12 from ~δ 21 in 3.9, 3.10, and 3.11. The ketone resonance (δ 214.2) was assigned to C-10 because it showed a gHMBC correlation to the methyl singlet at δ 1.15, assigned to Me-25 (Figure 3.19). A proton resonance at δ 7.22 (s), which did not show an HSQC correlation to a carbon atom, was assigned to a tertiary alcohol at C-9. The OH-9 showed gHMBC correlations to the ketone resonance at δ 214.2, assigned to C-10, a quaternary resonance at δ 65.5, assigned to C-8, an oxygenated methine resonance at δ 83.1, assigned to C-23, and a tertiary carbinol resonance at δ 89.5, assigned to C-9. A second proton resonance at δ 5.22 (s), which did not show a HSQC correlation to a carbon atom, was assigned to a tertiary alcohol at C-14. The OH-14 resonance showed gHMBC correlations to a tertiary carbinol resonance at δ 74.6, assigned to C-14, an olefinic resonance at δ 163.1, assigned to C-3, a methyl resonance at δ 32.9, assigned to Me-15, and a methine resonance at δ 48.8, assigned to C-13 (Figure 3.19). This set of OH-14 gHMBC correlations required that there were carbon-carbon bonds between C-3 and C-14, C-14 and Me-15, and C-13 and C-14. 1 3 20 2 14 10 9 16 23 5 21 8 4 13 22 11 7 12 6 18 19 15 24 25 17   68 The proton resonance (δ 2.55 dd, J = 13.2, 5.4 Hz) assigned to the C-13 methine (δ 48.8) in the gHSQC spectrum was shown in the gCOSY spectrum to be coupled to the methylene resonances assigned to H-12eq/H-12ax (δ 1.87 dd, J = 13.2, 5.4 Hz/ δ 1.96 bt, J = 13.2 Hz), indicating a C-12/C-13 bond. Finally, a gHMBC correlation observed between the methyl singlet at δ 1.08, assigned to Me-19, and the C-13 methine resonance at δ 48.8 required a carbon-carbon bond between C-7 and C-13 giving the seven membered ring drawn in structure 3.12.   Figure 3.19 Selected gHMBC and gCOSY correlations of dhilirolide D (3.12)   Figure 3.20 gCOSY spectrum of dhilirolide D (3.12) in DMSO-d6. Me25HO OOMe18Me17 O O OHO Me19Me15 Me241(! 163.9) 3(! 163.1)4(! 131.7) 8(! 65.5)(! 214.3)1011(! 55.2) 21(!148.0)713(! 48.8) (! 89.5)96 HMBCCOSY  69  Figure 3.21 gHMBC spectrum of dhilirolide D (3.12) in DMSO-d6. Based on the assumption that the isodhilirane skeleton is a biosynthetic precursor to the dhilirane carbon skeleton, the configurations at the common stereogenic centers in 3.9 and 3.12 are the same. A detailed analysis of the 2D ROESY data obtained for 3.12 (Figure 3.22) showed that the absolute configuration of 3.12 is 7S, 8R, 9R, 11R, 13R, 14R, and 23S.  Figure 3.22 Selected tROESY correlations for dhilirolide D (3.12) Me25Me19 OH23O OHOH12Me15OHO Me17Me18O HH H6axH6eqH13 Me24H H  70  Figure 3.23 tROESY spectrum of dhilirolide D (3.12) recorded in DMSO-d6 3.6.5 Structure Elucidation of Dhilirolide E Dhilirolide E (3.13, Figure 3.4) was isolated as an optically active amorphous white powder that gave a [M+Na]+ ion in the HRESITOFMS at m/z 465.1887 appropriate for a molecular formula of C25H30O7, which was identical to that of dhilirolide D (3.13). Although the 1H and 13C NMR spectra of 3.13 were similar to those of 3.12, their UV spectra were markedly different. In dhilirolide D (3.12, Figure 3.4), a λmax typical of a dienone was observed at 276 nm, while in 3.13 the λmax was reduced to 248 nm indicating a break in conjugation with the lactone, but still indicative of a diene. Simply shifting the 2,4-diene present in 3.12 to a 3,5-diene in 3.13 would explain the hypsochromic shift in the λmax. Examination of the 1H/13C/gCOSY/gHSQC/gHMBC NMR data obtained for 3.13 revealed that the olefinic H-2 resonance (δ 6.06) in 3.12 had been replaced by diastereotopic methylene proton resonances at δ 3.29 and 3.45, that showed gHMBC correlations to the lactone carbonyl resonance assigned to C-1 (δ 170.6). The NMR data of 3.13 identified the Δ21,22  exocyclic alkene seen in 3.12 and also revealed the presence of additional tetrasubstituted (C-3: δ 136.4; C-4: δ 129.4) and disubstituted   71 [C-5: δC 118.1, H-5: δH 5.62 (d, J = 12.3 Hz); C-6: δ 143.3, H-6: δ 5.98 (d, J = 12.3 Hz)] olefins. gHMBC correlations observed between H-2α/β (δ 3.45/3.29) and C-3 (δ 136.4) and C-4 (δ 129.4), between H-5 (δ 5.62) and C-3 (δ 136.4), between H-6 (δ 5.98) and C-4 (δ 129.4), between Me-15 (δ 1.08) and C-3 (δ 136.4), between Me-17/Me-18 (δ 1.47/1.38) and C-4 (δ 129.4), and between Me-19 (δ 1.29) and C-6 (δ 143.3) identified a 3,5-diene substructure in 3.13. tROESY correlations confirmed that the relative configuration of 3.13 was the same as 3.12, and, therefore, the absolute configuration of dhilirolide E (3.13, Figure 3.4) has been assigned as 7S, 8R, 9R, 11R, 13R, 14R, and 23S. Table 3.2 1H NMR data for dhilirolides E–I (3.13–3.17) recorded in DMSO-d6.  E (3.13) F (3.14) G (3.15) H (3.16) I (3.17) Atom # δH,  (J in Hz) δH,  (J in Hz) δH,  (J in Hz) δH,  (J in Hz) δH,  (J in Hz) 2α 2β 3.45 d, (19.9) 3.29 d, (19.9) 3.28 d, (21.5) 3.49 d, (21.5) 6.05 s 4.79 d, (5.1) 6.11 s 5 5.62 d, (12.3) 6.15 dd, (8.7, 4.1) 6.04 bm 6.18 m 6.03 dd, (7.2, 3.9) 6α 6β 5.98 d, (12.3) 2.20 dd, (14.3, 8.7) 1.62 dd, (14.3, 4.1) 2.21 dd, (19.5, 7.2) 3.03 dd, (19.5, 2.6) 1.83 dd, (14.9, 4.6) 2.07 dd, (14.9, 8.2) 2.63 dd, (15.9, 7.2) 2.94 dd, (15.9, 3.9) 9   5.78 s 5.81 s  10     3.62 d, (5.1) 12ax 12eq 2.24 t, (13.1) 1.71 dd, (13.1, 5.2) 1.85 t, (12.8) 1.56 dd, (12.8, 4.9) 1.71 t, (11.9) 1.93 dd, (11.9, 4.4) 1.93 dd, (12.3, 12.3) 1.60 dd, (12.3, 4.6) 1.78 t, (12.8) 1.92 dd, (12.8, 5.1) 13 2.51 dd, (13.1, 5.2) 2.75 dd, (12.8, 4.9) 2.27 dd, (11.9, 4.4) 2.49 nda 1.86 dd, (12.8, 5.1) 15 1.08 s 1.60 s 1.23 s 1.80 s 1.18 s 17 1.47 s 1.31 s 1.40 s 1.30 s 1.53 s 18 1.38 s 1.50 s 1.54 s 1.57 s 1.38 s 19 1.29 s 1.10 s 1.11 s 1.26 s 1.11 s 22α 22β 5.19 s 4.97 s 5.05 s 5.61 s 5.13 s 5.25 s 5.14 s 5.23 s 5.13 s 4.70 s 23 4.50 q, (7.0)     24 1.04 d, (7.0) 2.17 s    25 1.14 s 1.06 s 1.13 s 1.12 s 1.21 s 2-OH    5.90 d, (5.1)  9-OH 6.88 s 6.78 s    9-OAc   2.18 s 2.26 s  10-OH     6.19 d, (5.1) 14-OH 4.78 s  5.32 s  5.26 bs 20-OMe  3.60 s 3.71 s 3.71 s 3.58 s a Multiplicity not determined due to overlapping signals/chemical shifts determined from 2D data.       72 Table 3.3 13C NMR data for dhilirolides E–G (3.13–3.22) recorded in DMSO-d6.  E (3.13) F (3.14) G (3.15) H (3.16)a I (3.17) J (3.18) K (3.19)a L (3.20) M (3.21) N (3.22)a atom # δC δC δC δC δC δC δC δC δC δC 1 170.6 169.6 163.7 170.8 164.0 162.8 169.5 167.6 163.3 161.1 2 32.7 34.1 113.9 65.5 130.2 114.8 30.6 30.8 114.3 128.6 3 136.4 125.6 161.7 130.5 163.7 150.3 122.0 128.7 147.1 151.4 4 129.4 136.8 133.5 138.9 130.0 133.3 139.3 135.7 130.6 77.5 5 118.1 129.1 129.6 127.5 130.1 130.1 63.5 64.8 129.4 196.5 6 143.3 35.9 38.9 35.9 37.0 35.1 40.5 33.3 37.3 43.3 7 44.9 61.0 44.3 60.3 46.8 40.0 42.4 37.2 41.1 40.7 8 65.8 67.9 64.7 65.7 70.5 59.4 63.2 60.3 67.6 65.2 9 89.0 92.4 78.4 78.6 212.6 91.8 94.0 87.0 88.1 91.3 10 213.9 212.9 210.3 211.7 80.4 108.6 197.1 212.6 211.9 211.2 11 54.9 53.1 52.4 52.5 45.4 49.1 55.6 53.9 54.9 55.4 12 42.5 41.7 40.4 39.5 33.8 38.3 122.1 44.6 39.5 48.8 13 52.9 41.4 52.4 42.3 48.8 47.4 143.1 59.0 59.9 57.4 14 74.6 135.5 74.1 141.2 75.4 107.3  204.5 202.7 209.3 15 29.3 15.0 33.2 15.5 32.5 20.3  28.2 26.3 30.7 16 83.7 81.9 82.8 83.0 82.5 82.0 84.0 83.7 81.2 81.9 17 26.5 28.4 26.9 30.1 25.9 29.4 29.0 25.6 26.4 22.5 18 26.7 26.0 26.2 28.0 26.3 27.7 25.4 26.2 28.2 25.4 19 11.5 22.4 16.8 22.2 14.8 19.7 21.4 17.2 23.9 19.5 20 173.2 168.9 169.9 169.6 168.7 171.5 170.7 171.2 172.7 172.0 21 148.0 147.2 145.6 146.4 149.7 150.1 144.0 147.4 148.6 144.9 22 107.1 106.7 108.4 107.6 106.4 106.4 110.0 106.9 107.8 109.8 23 82.8 207.0    80.9 77.9 80.8 83.6 82.0 24 17.9 26.8    18.5 18.6 16.5 17.6 17.4 25 15.4 16.8 15.7 15.8 20.7 15.7 13.2 15.4 15.8 15.2 20-OMe  51.2 51.7 52.3 52.0      9-OAc   168.4/ 20.4 169.2/ 20.7   170.2/ 21.7    aMany of the chemical shift assignments are based on interpretation of the 2D NMR data due to limited material.  Figure 3.24 1H NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6   73  Figure 3.25 13C NMR spectrum of dhilirolide E (3.13) recorded at 150 MHz in DMSO-d6 3.6.6 Structure Elucidation of Dhilirolide F Dhilirolide F (3.14, Figure 3.4) was isolated as an optically active solid that gave a [M+Na]+ ion in the HRESITOFMS at m/z 479.2050 appropriate for a molecular formula of C26H32O7, that differs from the molecular formula of dhilirolides D (3.12, Figure 3.4) and E (3.13, Figure 3.4) by the addition of a carbon and two protons, but requires the same eleven sites of unsaturation. A UV λmax at 240 nm suggested a diene in the structure of 3.14 as is found in 3.13. However, significant differences in the 1H and 13C NMR spectra of dhilirolides F (3.14, Figure 3.4) and E (3.13, Figure 3.4) showed that the diene in 3.14 was in a different location than the diene in 3.13. The NMR data for 3.14 identified a trisubstituted Δ4,5 alkene (C-4: δ 136.8; C-5: δ 129.1, H-5: δ 6.15, dd, J = 8.7, 4.1 Hz), which is also found in dhilirolide D (3.12), but the resonance assigned to the OH-14 proton in dhilirolide D (3.12) and E (3.13) was missing and Me-15 had a chemical shift at δ 1.60 (s) (C-15: δ 15.0) appropriate for an olefinic methyl. A Δ3, 14 tetrasubstituted alkene (C-3: δ 125.6; C-14: δ 135.5) satisfied these spectroscopic observations and gHMBC correlations between H-2α/β (δ 3.28/3.49) and C-3 (δ 125.6), C-4 (δ 136.8) and C-14 (δ 135.5), between H-5 (δ 6.15) and C-3 (δ 125.6), between H-13 (δ 2.75) and C-3 (δ 125.6), C-14 (δ 135.5) and C-15 (δ 15.0), and between Me-15 (δ 1.60) and C-3 (δ 125.6), C-13 (δ 41.4) and C-14 (δ 135.5) confirmed the Δ3, 14 structural feature. Although the characteristic oxymethine quartet assigned to 2030405060708090100110120130140150160170180190200210 ppm  74 H-23 (δ 4.51- 4.89) in dhilirolides D (3.12) and E (3.13) was missing in the 1H NMR spectrum of 3.14, a singlet resonance at δ 6.78, that did not show a gHSQC correlation to a carbon atom, could still be assigned to a tertiary alcohol at C-9. The OH-9 resonance showed gHMBC correlations not only to the C-10 (δ 212.9) ketone, C-8 (δ 67.9), and C-9 (δ 92.4), but also to a second ketone carbon resonance at δ 207.0 (C-23). A methyl singlet with a chemical shift (δ 2.17, Me-24) appropriate for a α-keto methyl also showed HMBC correlations to the ketone resonance at δ 207.0 (C-23) and the C-9 resonance at δ 92.4, establishing the presence of a methyl ketone substituent at C-9. The remaining degree of unsaturation and C2H3O2 fragment required by the molecular formula was attributed to a methoxy carbonyl substituent at C-8 in 3.14 in place of the lactone carbonyl found at C-8 in 3.12 and 3.13. A strong gHMBC correlation observed between a methyl singlet at δ 3.60 and an ester carbonyl at δ 168.9, assigned to C-20, supported a methoxy carbonyl substituent at C-8. tROESY correlations observed between the OH-9 (δ 6.78) resonance and the H-5 (δ 6.15) and H-13 (δ 2.75) resonances showed that the configuration at C-9 was identical to the C-9 configuration in dhilirolide D (3.12, Figure 3.4) and additional tROESY correlations revealed that the relative configurations at the C-7, C-8, C-11, and C-13 stereogenic centres shared by 3.12 and 3.14 were identical, leading to the absolute configuration assignment of 7S, 8R, 9R, 11R, and 13R for dhilirolide F (3.14, Figure 3.4).   Figure 3.26 1H NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6   75  Figure 3.27 13C NMR spectrum of dhilirolide F (3.14) recorded at 150 MHz in DMSO-d6. 3.6.7 Structure Elucidation of Dhilirolide G Dhilirolide G (3.15, Figure 3.4) was isolated as an optically active solid that gave a [M+Na]+ ion in the HRESITOFMS at m/z 495.1982 appropriate for a molecular formula of C26H32O8, that differs from the molecular formula of dhilirolide F (3.14, Figure 3.4) by the addition of an oxygen. Detailed analysis of the 1D and 2D NMR data and UV spectra (λmax 269 nm) obtained for dhilirolide G (3.15, Figure 3.4) revealed that the constitution of rings A, B and C were the same as in dhilirolide D (3.12, Figure 3.4) and that the methyl ester seen in dhilirolide F (3.14) at C-20 [C20: δ 169.9; 20-OMe: δC 51.7, δH 3.71, s] was intact. However, compared with 3.14, the resonances assigned to the C-23/C-24 methyl ketone substituent attached at C-9 and the OH-9 at C-9 had been replaced by resonances appropriate for an acetylated oxymethine [C-9: δC 78.4, H-9: δH 5.78 (s); OAc: δC 168.4, 20.4, δH 2.18]. gHMBC correlations between the H-9 oxymethine singlet resonating at δ 5.78 and the acetate carbonyl at δ 168.4, and between the H-9 resonance and the C-7 (δ 44.3), C-8 (δ 64.7), C-10 (δ 210.3), C-11 (δ 52.4: weak) and C-20 (δ 169.9) resonances confirmed the structural fragment and accounted for the molecular formula. tROESY correlations between the H-9 (δ 5.78) and both of the H-6β (δ 3.03) and H-13 (δ 2.27) 2030405060708090100110120130140150160170180190200210 ppm  76 resonances showed that H-9 was cis to C-7 and, therefore, the absolute configuration for 3.15 was assigned as 7S, 8R, 9R, 11R, 13R, and 14R.    Figure 3.28 1H NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6  Figure 3.29 13C NMR spectrum of dhilirolide G (3.15) recorded at 150 MHz in DMSO-d6 102030405060708090100110120130140150160170180190200210 ppm  77       Figure 3.30 tROESY NMR spectrum and expansions of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6  !"#$!"%&'!"#$!"%&'  78 3.6.8 Structure Elucidation of Dhilirolide H Dhilirolide H (3.16, Figure 3.4) was isolated as an optically active solid that gave a [M-H]– ion in the HRESITOFMS at m/z 471.2014 appropriate for a molecular formula of C26H32O8, the same as dhilirolide G (3.15). Extensive analysis of the 1D and 2D NMR spectra of dhilirolide H (3.16) (Figures 3.31–3.34) and comparison with the NMR data for dhilirolides D–G (3.12–3.15) revealed that the constitution of ring B was the same as that seen in dhilirolide F (3.14) and rings C and D were the same as seen in dhilirolide G (3.15, Figure 3.4), while structural novelty was present in ring A. An oxymethine doublet resonating at δ 4.79 (J = 5.1 Hz) (H-2) was coupled in the gCOSY 60 experiment to a hydroxyl doublet at δ 5.90 (J = 5.1 Hz). gHMBC correlations between H-2 (δ 4.79) and C-1 (δ 170.8), C-3 (δ 130.5), C-4 (δ 138.9) and C-14 (δ 141.2) placed a secondary alcohol at C-2. The observation of tROESY correlations between Me-17 (δ 1.30) and both of the H-2 (δ 4.79) and H-13 (δ 2.49) resonances required the alcohol at C-2 to be in the α orientation as shown. Additional tROESY correlations observed between H-9 (δ 5.81) and both H-6β (δ 2.07) and H-13 (δ 2.49) showed that the relative configuration at C-9 in 3.16 was the same as in 3.15, in agreement with the nearly identical 13C chemical shifts observed for C-8 to C-12 in 3.15 and 3.16 (Table 3.3).  Two quite different conformations, which must be equilibrating rapidly on the NMR time scale, are required to generate the observed Me-17 to H-2 and the Me-17 to H-13 tROESY correlations in 3.16. By analogy with dhilirolide D (3.12), the absolute configuration of dhilirolide H (3.16, Figure 3.4) was assigned as 2R, 7S, 8R, 9R, 11R, and 13R.      79  Figure 3.31 1H NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6   Figure 3.32 13C NMR spectrum of dhilirolide H (3.16) recorded at 150 MHz in DMSO-d6   102030405060708090100110120130140150160170180190200210 ppm  80    Figure 3.33 tROESY NMR spectrum expansions of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6  !"#$%!"#&'!"#$%!"#&'  81  Figure 3.34 tROESY NMR spectrum expansions of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6 3.6.9 Structure Elucidation of Dhilirolide I Dhilirolide I (3.17, Figure 3.4) gave a [M+Na]+ ion in the HRESITOFMS at m/z 453.1920 appropriate for the molecular formula C24H30O7, that differs from that of dhilirolide D (3.12, Figure 3.4) by the loss of one carbon atom, and required only ten sites of unsaturation instead of eleven. Examination of the 1H and 13C NMR spectra recorded for dhilirolide I (3.17, Figure 3.4) revealed a close relationship with dhilirolides D (3.12) and G (3.15), but also several significant differences in ring D. Resonances attributable to the methyl esters seen in 3.14, 3.15, and 3.16 were observed in the NMR data obtained for 3.17 (C20: δ 168.7; 20-OMe: δC 52.0, δH 3.58). However the methyl singlet assigned to Me-25 (δ 1.21) in 3.17 no longer showed a gHMBC correlation to a ketone resonance that could be assigned to C-10. Instead a correlation was seen to a carbinol methine resonating at δ 80.4, that in the gHSQC correlated to a doublet at δ 3.62 (J = 5.1 Hz) (H-10), which was in turn coupled in the gCOSY60 to an exchangeable resonance at δ 6.19 (d, J = 5.1 Hz) (OH-10). These observations suggested a switch in the locations of the C-9 carbinol methine and C-10 ketone seen in dhilirolides G (3.15) and H (3.16) to a C-9 ketone and C-10 carbinol methine in dhilirolide I (3.16). gHMBC correlations between the H-10 resonance (δ 3.62) and a ketone resonance at δ 212.6, that could only be assigned to C-9 since H-10 also correlated with C-11 (δ 45.4), C-12 (δ 33.8) and C-25 (δ 20.7), confirmed the switch. The change in functionality at C-9 and C-10 in 3.17 was further supported by a tROESY correlation between H-10 (δ 3.62) and Me-25 (δ 1.21) and a strong gHMBC correlation between Me-25 (δ 1.21) and C-10 (δ 80.4). tROESY correlations observed between OH-10 (δ 6.19) and both H-12eq (δ 1.92), and H-13 (δ 1.86), and between H-10 (δ 3.62) and both of the H-22 resonances (δ 4.70, !"#$!"%&'!"#$!"%&'!"#$!"()'  82 5.13) showed that OH-10 was cis to C-12. Therefore, the absolute configuration of dhilirolide I (3.17, Figure 3.4) was assigned as 7S, 8R, 10R, 11R, 13R, and 14R.  Figure 3.35 1H NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6  Figure 3.36 13C NMR spectrum of dhilirolide I (3.17) recorded at 150 MHz in DMSO-d6 2030405060708090100110120130140150160170180190200210 ppm  83  Figure 3.37 gHMBC NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6  Figure 3.38 gHMBC NMR spectrum expansions of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 !"#$%&"'()!"'(%&"#$)  84  Figure 3.39 tROESY NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6   Figure 3.40 tROESY NMR spectrum expansions of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6 !"#$%!"&&'( !"#$%!"&&)(  85  Table 3.4 1H NMR data for dhilirolides J–N (3.18–3.22) recorded in DMSO-d6.  J (3.18) K (3.19) L (3.20) M (3.21) N (3.22) position # δH,  (J in Hz) δH,  (J in Hz) δH,  (J in Hz) δH,  (J in Hz) δH,  (J in Hz) 2α 2β 5.77 s 3.06 bd, (19.9) 3.43 dd, (19.9, 2.3) 2.58 d, (19.8) 2.90 d, (19.8) 5.94 s 6.80 s 5 6.21 m 4.39 bm 4.77 bs 5.97 bm  6α 6β 1.98 dd, (19.2, 6.1) 3.00 d, (19.2) 2.04 dd, (11.5, 5.5) 1.70 bt, (11.5) 2.19 bd, (13.6)a 1.80 bd, (13.6)a 2.48 dd, (20.6, 4.1) 2.61 dd, (20.6, 3.8) 2.81 d, (20.7)a 2.79 d, (20.7)a 12ax 12eq 1.45 d, (12.3) 2.09 d, (12.3) 5.35 s 2.55 d, (14.1) 1.87 d, (14.1) 2.38 d, (14.3) 3.00 d, (14.3) 3.06 d, (13.6) 2.91 d, (13.6) 15 1.48 s  2.06 s 2.05 s 2.36 s 17 1.50 s 1.52 s 1.55 s 1.49 s 1.55 s 18 1.54 s 1.63 s 1.49 s 1.12 s 1.36 s 19 0.95 s 1.34 s 1.27 s 1.41 s 1.40 s 22α 22β 5.07 s 4.93 s 5.38 s 5.08 s 5.25 bs 5.03 bs 5.19 s 4.98 bs 5.36 s 5.16 s 23 4.68 q, (7.0) 4.92 q, (7.0) 4.45 q, (7.2) 4.32 q, (7.1) 4.69 q, (7.7) 24 1.37 d, (7.0) 1.10 d, (7.0) 1.00 d, (7.2) 0.95 d, (7.1) 0.97 d, (7.7) 25 1.10 s 1.35 s 1.14 s 1.24 s 1.21 s 5-OH  5.24 d, (6.2)    9-OH    6.57 s  9-OAc  2.07 s    10-OH 7.27 s     aAssignments within a column maybe interchanged 3.6.10  Structure Elucidation of Dhilirolide J Dhilirolide J (3.18, Figure 3.4) was isolated as an optically active crystalline solid that gave a [M+H]+ ion in the HRESITOFMS at m/z 441.1918 appropriate for a molecular formula of C25H28O7, that differed from dhilirolides B (3.10) and C (3.11) by the loss of one oxygen atom. Comparison of the 1H and 13C NMR spectrum of dhilirolide J (3.18) with the corresponding spectra recorded for dhilirolides B (3.10) and C (3.11) revealed the absence of resonances that could be attributed to epoxide fragments. Analysis of the NMR data and UV spectra (λmax 279 nm) suggested that dhilirolide J (3.18) contained the C-1 to C-5 dienone (C-1: δC 162.8; C-2: δC 114.8, H-2: δH 5.77; C-3: δC 150.3; C-4: δC 133.3; C-5: δC 130.1, δH 6.21) seen in dhilirolide B (3.10, Figure 3.4). As with dhilirolide C (3.11, Figure 3.4), the C-21/C-22 epoxide found in dhilirolide A (3.9) had been replaced with ∆21, 22 exocyclic alkene (C-21: δC 150.1; C-22: δC 106.4, H-22a/b: δH 5.07/4.93) in 3.18. The stereogenic centers in 3.18 are all shared with dhilirolide A (3.9) and, therefore, the absolute configuration of dhilirolide J (3.18, Figure 3.4) was assigned as 7R, 8R, 9R, 10R, 11R, 13S, 14S and 23S.   86  Figure 3.41 1H NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6.  Figure 3.42 13C NMR spectrum of dhilirolide J (3.18) recorded at 150 MHz in DMSO-d6. 3.6.11 Structure Elucidation of Dhilirolide K Dhilirolide K (3.19, Figure 3.4) gave a [M+Na]+ ion in the HRESITOFMS at m/z 479.1679 appropriate for a molecular formula of C25H28O8. Although this was the same molecular formula as observed for dhilirolide C (3.11), inspection of the NMR data and UV 176 168 160 152 144 136 128 120 112 104 96 88 80 72 64 56 48 40 32 24 16 8Chemical Shift (ppm)  87 spectra for 3.19 revealed significant structural and functional group differences compared with 3.11. Reminiscent of dhilirolides D (3.12, Figure 3.4) and E (3.13), NMR resonances observed for 3.19 could be assigned to the g-lactone and ring D substructures seen in 3.12 and 3.13 with the one difference being that an acetate functionality (δC 170.2/21.7, dH 2.07) was at C-9 in 3.19 in place of an exchangeable OH-9 proton. The UV λmax at 249 nm for 3.19 was suggestive of a diene and not the enone that is sometimes encountered in ring A of the dhilirolides. A pair of diastereotopic methylene resonances at δ 3.06 and 3.43 (H-2a/b) in the 1H NMR spectrum of 3.19, that showed gHMBC correlations to the lactone carbonyl assigned to C-1 (δ 169.5), supported the absence of a Δ2, 3 alkene in 3.19. However, the NMR data for 3.19 did confirm the presence of tetra- and tri-substituted conjugated Δ3, 4 and Δ12, 13 olefins (C-3: δC 122.0; C-4: 139.3; C-12:  122.1, H-12: δH 5.35; C-13: δC 143.1). A proton resonance at δ 5.24 (d, J = 6.2 Hz), that did not show a gHSQC correlation to a carbon atom but coupled to a multiplet at δ 4.39 (H-5) in the gCOSY60, was assigned to a secondary alcohol at C-5. The OH-5 showed gHMBC correlations to one of the olefinic carbons of the diene at δ 139.3. Since Me-17/18 (δ 1.52/1.63) and H-2a/b (δ 3.06/3.43) also correlated to the carbon at δ 139.3 in the gHMBC, this resonance was assigned to C-4. These observations supported tetrasubstituted ∆3,4 and trisubstituted ∆12,13 conjugated alkenes in the structure of 3.19 and the absence of the C-14-C-15 carbon fragment that is common to dhilirolides A–C (3.9–3.11) and J (3.18). The addition of the acetate at C-9 balances the loss of the C2H3O C-14/C-15 fragment and satisfies the constraints imposed by the molecular formula. gHMBC correlations observed between Me-19 (δ 1.34) and C-13 (δ 143.1), between Me-25 (δ 1.35) and C-12 (δ 122.1), between H-12 (δ 5.35) and both C-3 (δ 122.0) and C-13 (δ 143.1), and between H-2a/b (δ 3.06/3.43) and C-3 (δ 122.0), C-4 (δ 139.3) and C-13 (δ 143.1) were consistent with the proposed structure 3.19. The tROESY data for dhilirolide K (3.19, Figure 3.4) showed correlations between H-5 (δ 4.39) and both Me-18 (δ 1.63) and Me-19 (δ 1.34) suggesting that the C-5 alcohol is in the β orientation. Based on the assumption that the configurations at the common stereogenic centres in dhilirolides A (3.9) and 3.19 are the same, a detailed analysis of the 2D tROESY data obtained for K (3.19, Figure 3.4) confirmed that the absolute configuration of as 5R, 7S, 8R, 9R, 11S, and 23S.   88  Figure 3.43 1H NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6   Figure 3.44 13C NMR spectrum of dhilirolide K (3.19) recorded at 150 MHz in DMSO-d6 −10220 210 200 190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 ppm  89 3.6.12 Structure Elucidation of Dhilirolide L Dhilirolide L (3.20, Figure 3.4) was isolated as optically active clear prism like crystals that gave a [M+Na]+ ion in the HRESITOFMS at m/z 463.1745 appropriate for a molecular formula of C25H28O7. This was the same molecular formula as observed for dhilirolide J (3.18), but there were significant differences in the NMR data obtained for 3.18 and 3.20. As with dhilirolide K (3.19), the g-lactone and ring D substructures seen in 3.12 and 3.13 were present in the structure of 3.20. However, instead of an acetate substituent at C-9 an ether linkage between C-5 and C-9 was revealed by a gHMBC correlation observed between H-5 (δ 4.77) and C-9 (δ 87.0). The lack of any significant UV absorption for 3.20 precluded any of the conjugation seen in dhilirolides A–K (3.9–3.19). gHMBC correlations between the diastereotopic methylene proton resonances assigned to H-2a/b (δ 2.58/2.90) and C-3 (δ 128.7) and C-4 (δ 135.7), between H-5 (δ 4.77) and C-3 (δ 128.7) and C-4 (δ 135.7), and between Me-17/18 (δ 1.55/1.49) and C-4 (δ 135.7) established the presence of a tetrasubstituted Δ3, 4 alkene. gHMBC correlations observed between a methyl singlet at δ 2.06, assigned to Me-15 and carbon resonances at δ 204.5 (C-14), assigned to a ketone, and a quaternary resonance at δ 59.0, assigned to C-13, were consistent with a methyl ketone substituent attached to C-13.   Figure 3.45 1H NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6   90  Figure 3.46 ORTEP diagram for dhilirolide L (3.20).   Figure 3.47 13C NMR spectrum of dhilirolide L (3.20) recorded at 150 MHz in DMSO-d6 The rigidity of dhilirolide L (3.20, Figure 3.4), resulting from its fused polycyclic structure, facilitated the assignment of the relative configuration after analysis of all of the observed tROESY data. However, there was no unambiguous NMR evidence for the configuration at C-5, even though the proposed caged structure required that C-5 had the same configuration as in 3.19. In order to verify the proposed structure, dhilirolide L (3.20) was subjected by Brian O. Patrick to single crystal X-ray diffraction analysis, using Cu-Kα radiation. 2030405060708090100110120130140150160170180190200210 ppm  91 The absolute configuration was established on the basis of the refined Flack x-parameter [x = 0.00(4)].134, 135 The ORTEP diagram in Figure 3.46 shows the constitution and absolute configuration (5R, 7S, 8R, 9R, 11R, 13R and 23S) of dhilirolide L (3.20, Figure 3.4). At the common stereogenic centers, this is identical to the absolute configuration determined by single crystal X-ray diffraction analysis for dhilirolide A (3.9, Figure 3.4). At C-13, the configuration in 3.20 has been inverted compared with 3.9. 3.6.13 Structure Elucidation of Dhilirolide M Dhilirolide M (3.21, Figure 3.4) gave a [M+Na]+ ion in the HRESITOFMS at m/z 463.1740 appropriate for a molecular formula of C25H28O7, the same as dhilirolide L (3.20). Although the 1H and 13C NMR spectrum of 3.21 were similar to that of 3.20, the UV spectrum was markedly different. A λmax at 273 nm was again indicative of the dienone substructure seen in dhilirolides B (3.10), D (3.12), G (3.15), I (3.17) and J (3.19, Figure 3.4). Examination of the NMR data for 3.21 confirmed the presence of a C-1 to C-5 dienone. Since all other structural features of 3.21 appeared to be the same as in dhilirolide L (3.20), it was apparent that a tertiary alcohol should reside at C-9 to satisfy the requirements of the molecular formula. A proton resonance at δ 6.57 (s), that did not show a gHSQC correlation to a carbon, was assigned to the alcohol at C-9. The OH-9 showed gHMBC correlations to the ketone resonance at δ 211.9, assigned to C-10, a quaternary resonance at δ 67.6, assigned to C-8, an oxygenated methine resonance at δ 83.6, assigned to C-23, and a tertiary carbinol resonance at δ 88.1, assigned to C-9 which were consistent with the proposed structure. The tROESY data was consistent with the assumption that the configurations at all the common stereogenic centres in 3.20 and 3.21 were the same giving the absolute configuration of dhilirolide M (3.21, Figure 3.4) as 7S, 8R, 9R, 11R, 13R and 23S.         92  Figure 3.48 1H NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6  Figure 3.49 13C NMR spectrum of dhilirolide M (3.21) recorded at 150 MHz in DMSO-d6   93 Dhilirolide M (3.21, Figure 3.4) was isolated as a single peak from reversed phase HPLC that was well resolved from the HPLC peak containing dhilirolide L (3.20, Figure 3.4), as described in the experimental section.  However, when the 1D 1H NMR spectrum of dhilirolide M (3.21) was recorded in DMSO-d6, it showed evidence for small amounts of dhilirolide L (3.20). During the course of collecting a 2D NMR data set on dhilirolide M (3.21), the amount of dhilirolide L (3.20) in the sample increased until it was present at a concentration of roughly 10% of the dhilirolide M (3.21) concentration. Longer exposure in the NMR tube in DMSO-d6 resulted in almost complete conversion of dhilirolide M (3.21) to dhilirolide L (3.20). It was also observed that NMR samples of dhilirolide L (3.20) that sat for several days in DMSO-d6 started to show the presence of minor amounts of dhilirolide M (3.21). Thus, it appears that dhilirolides L (3.20) and M (3.21) are able to slowly interconvert in DMSO-d6.  3.6.14 Structure Elucidation of Dhilirolide N Dhilirolide N (3.22, Figure 3.4) gave a [M+Na]+ peak in the HRESITOFMS at m/z 477.1521 appropriate for a molecular formula of C25H26O8, that differs from the molecular formula for dhilirolide L (3.20) by the loss of two protons, the addition of one oxygen, and requires one additional site of unsaturation. Comparison of the NMR data obtained for 3.20 and 3.22, revealed the presence of an additional ketone resonance at δ 196.5 (C-5) that accounted for the additional site of unsaturation, the extra oxygen, and the loss of two hydrogen atoms. The third ketone in 3.22 was assigned to C-5 since gHMBC correlations were observed between C-5 (δ 196.5) and both H-6a/b (δ 2.81/2.79) and Me-19 (weak, δ 1.40). A singlet olefinic methine resonance at δ 6.80 (H-2) was suggestive of the trisubstituted α, β-unsaturated lactone seen in dhilirolides A (3.9) and C (3.11). gHMBC correlations from H-2 (δ 6.80) to the lactone C-1 carbonyl (δ 161.1) and C-13 (δ 57.4), and between H-12ax (δ 3.06) and C-3 (δ 151.4) confirmed the presence of an enone in ring A. An oxygenated tertiary carbon resonating at δ 77.5 was assigned to C-4 since it showed gHMBC correlations to both H-2 (δ 6.80) and Me-17/18 (δ 1.55/1.36). The g-lactone and ring D substructures and the methyl ketone residue at C-13 in 3.22 were all identical to those seen in dhilirolide L (3.20, Figure 3.4). With no other apparent structural or functional differences between 3.20 and 3.22 simply satisfying the valences on the two oxygenated quaternary carbons C-4 and C-9 (δC 77.5 and 91.3, respectively), through an ether linkage provided the final ring required by the molecular formula and completed the   94 constitution of 3.22. A tROESY correlation observed between Me-17 (δ 1.55) and H-23 (δ 4.69) provided additional support for the C-4/C-9 ether linkage and established the relative configuration at C-4. By analogy with dhilirolide L (3.20), the absolute configuration of dhilirolide N (3.22, Figure 3.4) was assigned as 4R, 7S, 8R, 9R, 11R, 13R and 23S.   Figure 3.50 1H NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6.  Figure 3.51 13C NMR spectrum of dhilirolide N (3.22) recorded at 150 MHz in DMSO-d6 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0Chemical Shift (ppm)2030405060708090100110120130140150160170180190200210 ppm  95 3.7 Dhilirolide Feeding Studies and Biosynthetic Proposal The dhilirolides are presumed to be highly rearranged meroterpenoids. Dhilirolides D–I (3.12–3.17, Figure 3.4), possessing the isodhilirane (III, Figure 3.4) or dinorisodhilirane (IV, Figure 3.4) carbon skeleton, share many structural features with berkeleydione115 and the miniolutelides (Figure 3.2).116 We undertook a stable isotope feeding study with [1,2-13C2]-acetate aimed at providing experimental support for the proposed biogenesis of the new dhilirane (I, Figure 3.4), isodhilirane (II, Figure 3.4), 14, 15-dinordhilarane (III, Figure 3.4), and 23, 24-dinorisodhilarane (IV) (Figure 3.4) carbon skeletons. Dilip E. DeSilva performed the feeding study of the organisms in Sri Lanka. In the [1,2-13C2]-acetate feeding study, no statistically significant 13C-enrichment was observed at any site, just as was found in the terretonin136 study when labeled [1-13C]-acetate was fed. However, 13C-13C coupling satellites were observed for all the carbon resonances except C-3, C-6, C-17, C-24, and C-25 in the 13C spectra of the dhilirolides obtained from P. purpurogenum cultures fed [1,2-13C2]-acetate. The average level of enrichment was estimated at 0.14% from the relative heights of the coupling satellites and the singlet natural abundance signals. Dhilirolide A (3.9) was isolated in sufficient quantities to acquire a 2D-INADEQUATE spectrum (Figure 3.52), which allowed the unambiguous assignment of intact acetate units as summarized in Scheme 3.1. David E. Williams isolated all 13C labeled compounds and ran all spectra relating to the feeding studies. A proposed biogenetic pathway to the dhilirolides that accounts for the [1,2-13C2]-acetate incorporation data is shown in Scheme 3.1. The pathway provides additional support for the earlier labeling work of the Simpson and Vederas groups.103, 106, 120 As observed, no incorporation would be expected in the dhilirolides at Me-24 and Me-25 since the 3- and 5-methyl substituents of dimethylorsellinate (3.24) are presumably derived from SAM.103, 106, 136    96  Figure 3.52 2D-INADEQUATE spectrum of dhilirolide A (3.9), obtained from a 13C doubly labeled acetate feeding experiment, recorded at 150 MHz in DMSO-d6.    97   Scheme 3.1 Proposed biogenetic pathway for the dhilirolides showing representative examples of each of the new dhilirane, isodhilirane, 14, 15-dinordhilirane and 23, 24-dinorisodhilirane skeletons. The labeling pattern indicating intact acetate units shown for dhilirolide A (3.9) was determined from the [1,2-13C2]-acetate feeding study.    OPP•O OH OPP• • • OHCO2HOH! !+ •• •O OHCO2HO !!H O OCO2HHO • • •! !H2O[O] HOCO2HO • •!OH OHOH• !OCO2H! OHOH !OOCO2H OHOHO OOCH2O2H! OHOH !O O OCH2O2H! OHOH !OO OCO2H OHOHO[O] OCO2H OHOHOOOCO2H OHOHO[O] OCO2H OHOHOOO Dhilirolide K (3.19)'14,15-Dinordhilirane'O OAcOO OH OO OCO2Me OAcOOHODhilirolide H (3.16)'23,24-Dinorisodhilirane'OCO2H OOHOOOO OO O OHOO O [O]Baeyer-Villiger,hydrolysis,reduction,acetyaltion!O OH + SAM• ••! !• ••! !• ••HH •••• • •! !• • •Dhilirolide A (3.9)'Dhilirane'[O] Baeyer-Villiger, hydrolysis,dehydration O OHOO OOHODhilirolide D (3.12)"Isodhilirane" [O]24253.24 173.23[1,2-13C2]acetate [1,2-13C2]acetate  98 3.8 Dhilirolide Biological Activity After isolation of the dhilirolides was complete several biological assays were attempted initially with no success.  The dhilirolides were also tested on the fruits of A. bilimbi, with no damage being seen from either surface application or application to a lesion.  As a result, the crude fungal extract as well as the major constituents dhilirolides A (3.9), D (3.12), and L (3.20, Figure 3.4) were evaluated for insecticidal and anti-feedant bioactivity against the cabbage looper Trichoplusia ni (Lepidoptera: Noctuidae), an economically important agricultural pest. Dr. Yasmin Ahktar performed these experiments in Professor Murray Isman’s laboratory. The crude extract was a modest feeding inhibitor to third instar T. ni larvae, but an LH20 fraction enriched in dhilirolides A–N (3.9–3.22, Figure 3.4) was considerably more active in a two-choice feeding bioassay. The DC50 (concentration reducing feeding by 50%) for the LH20 fraction was 25 µg/cm2; comparable to a number of natural and synthetic anti-feedants previously tested against this pest.137–140 Dhilirolide L (3.20, Figure 3.4) showed the best activity of the pure compounds, giving a DC50 of 5.9 µg/cm2.   When sprayed directly on second instar larvae as a 1% aqueous emulsion, the crude extract caused 47% mortality, while the previously mentioned LH20 fraction caused 63% mortality.  Addition of either the crude extract or LH20 fraction to the insect’s diet at 1000-ppm fwt resulted in significant sub-lethal effects.  The crude extract reduced larval growth by 20% and adult emergence by 56%, while the dhilirolide-enriched LH20 fraction reduced larval growth by 70% and adult emergence by 87.5%.  3.9 Conclusion  The isolation of 14 new compounds named dhilirolides A–N (3.9–3.22, Figure 3.4), from P. purpurogenum, has resulted in the identification of the novel dhilirane (I), isodhilirane (II), 14, 15-dinordhilarane (III) and 23, 24-dinorisodhilarane (IV) carbon skeletons. Feeding studies of P. purpurogenum have confirmed the proposed mixed terpene-polyketide biogenesis of the dhilirolides.  Studies that show the potent anti-feedant activity of the dhilirolides, especially dhilirolide L (3.20), have been accomplished.  The role of the dhilirolides in the fungus remains uncertain.  However, the clear insecticidal activity of the dhilirolides, coupled with P. purpurogenum being found to infect and decay only fruit could possibly point to an endophytic relationship with Averrhoa bilimbi.  The dhilirolides likely do not play a clear role in fungal   99 infection, not causing any apparent damage to the fruit when spread on the surface or inside an opened lesion (experiments not shown).  The dhilirolides, therefore, could provide a protective role in the plant against herbivores. In return the fungus is then able to grow in a mutualistic fashion as an endophyte, with this balance perhaps being tipped towards fungal parasitism once the fruit nears maturation. Confirmation of an endophytic relationship, like the one described, would require additional experiments in the natural habitat of Averrhoa bilimbi and P. purpurogenum. 3.10 General Experimental Procedures  Optical rotations were measured using a polarimeter with sodium light (589 nm). UV spectra were recorded with a HPLC dual λ absorbance detector. The 1H and 13C NMR spectra were recorded on a 600 MHz spectrometer with a 5 mm cryoprobe. 1H chemical shifts are referenced to the residual DMSO-d6 (δ 2.50) and 13C chemical shifts are referenced to the DMSO-d6 (δ 39.5) solvent peak. Reversed-phase HPLC purifications were performed using a binary HPLC pump attached to a dual λ absorbance detector or a photodiode array detector. All solvents used for HPLC were HPLC grade and were filtered through a 0.45 µm filter prior to use.  3.10.1 Fungal Material The fungus Penicillium purpurogenum was isolated from infected fruits of Averrhoa bilimbi (Averrhoa bilimbi, L., (Oxalidaceae) in Sri Lanka. 3.10.2 Extraction of P. purpurogenum and Isolation of Dhilirolides A–N P. purpurogenum was cultured on potato dextrose agar (50 Petri dishes) and the culture medium cut into small pieces and extracted with EtOAc. The EtOAc extract was concentrated in vacuo and chromatographed on Sephadex LH20 (3 cm x 95 cm) using 4:1 MeOH/CH2Cl2. The fractions containing the compounds of interest were combined and subjected to reversed-phase C18 HPLC chromatography using a CSC-Inertsil 150Å/ODS2, 5µm 25 x 0.94 cm column, with a linear gradient of 40–45% ACN/H2O over 60 min at a flow rate of 2 mL/min. Pure samples of dhilirolides A (3.9) (28.9 mg), B (3.10) (7.6 mg), C (3.11) (17.7 mg), D (3.12) (8.7 mg), F (3.14) (0.4 mg), J (3.18) (2.5 mg), L (3.20) (14.8 mg), M (3.21) (1.0 mg) and N (3.22) (0.5 mg) were obtained. Additional fractions containing impure samples of dhilirolides E (3.13), G (3.15), H (3.16), I (3.17) and K (3.19) were fractionated via isocratic C8 reversed-phase HPLC using a Phenomenex Luna 100 Å, 5µm 25 x 1.0 cm column. Compounds are listed with eluent systems   100 and yields: 3.13 (29:71 ACN/H2O, 0.9 mg), 3.15 (9:16 ACN/H2O, 0.5 mg), 3.16 (7:13 ACN/H2O, 1.0 mg), 3.17 (9:16 ACN/H2O, 0.9 mg) and 3.19 (33:67 ACN/H2O, 1.4 mg).  Dhilirolide A (3.9): Isolated as irregular colourless crystals; mp 267–269 °C; [α]25D -128° (c 0.24, MeOH); UV (9:11 ACN/H2O) λmax (ε) 235 (3.8) nm; 1H NMR, see Table 3.1; 13C NMR, see Table 3.1; positive ion HRESITOFMS [M+H]+ m/z 473.1796 (calcd for C25H29O9, 473.1811). Dhilirolide B (3.10): Isolated as a clear amorphous solid; [α]25D +65° (c 0.40, MeOH); UV (9:11 ACN/H2O) λmax (ε) 220 (3.2), 280 (3.5) nm; 1H NMR, see Table 3.1; 13C NMR, see Table 3.1; positive ion HRESITOFMS [M+Na]+ m/z 479.1725 (calcd for C25H28O8Na, 479.1682).  Dhilirolide C (3.11): Isolated as a clear amorphous solid; [α]25D -53° (c 0.76, MeOH); UV (9:11 ACN/H2O) λmax (ε) 239 (3.2) nm; 1H NMR, see Table 3.1; 13C NMR, see Table 3.1; positive ion HRESIMS [M+Na]+ m/z 479.1746 (calcd for C25H28O8Na, 479.1682). Dhilirolide D (3.12): Isolated as a clear amorphous solid; [α]25D +115° (c 0.80, MeOH); UV (9:11 ACN/H2O) λmax (ε) 201 (3600), 276 (3500) nm; 1H NMR, see Table 3.1; 13C NMR, see Table 3.1; negative ion HRESITOFMS [M-H]- m/z 441.1967 (calcd for C25H29O7, 441.1913). Dhilirolide E (3.13): Isolated as an amorphous white powder; [α]20D +3.23 (c 0.09, MeOH); UV (ACN) λmax (ε) 206 (3103), 248 (2348) nm; 1H NMR, see Table 3.2; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 465.1887 (calcd for C25H31O7Na, 465.1889). Dhilirolide F (3.14): Isolated as an amorphous white solid; [α]20D  +18.1 (c 0.03, MeOH); UV (ACN) λmax (ε) 204 (7859), 240 (4037) nm; 1H NMR, see Table 3.2; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 479.2050 (calcd for C26H32O7Na, 479.2046). Dhilirolide G (3.15): Isolated as an amorphous white solid; [α]20D  +8.9 (c 0.05, MeOH); UV (ACN) λmax (ε) 204 (5575), 269 (4786) nm; 1H NMR, see Table 3.2; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 495.1982 (calcd for C26H32O8Na, 495.1995). Dhilirolide H (3.16): Isolated as an amorphous white solid; [α]20D  +2.92 (c 0.10, MeOH); UV (ACN) λmax (ε) 201 (1847), 240 (941) nm; 1H NMR, see Table 3.2; 13C NMR, see Table 3.3; negative ion HRESITOFMS [M-H]- m/z 471.2014 (calcd for C26H31O8, 471.2019).   101 Dhilirolide I (3.17): Isolated as an amorphous white solid; [α]20D +2.89 (c 0.09, MeOH); UV (ACN) λmax (ε) 203 (2700), 272 (2067) nm; 1H NMR, see Table 3.2; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 453.1920 (calcd for C24H30O7Na, 453.1889). Dhilirolide J (3.18): Isolated as a clear crystalline solid; decomposed at 223–225°C; [α]20D  +5.94 (c 0.16, MeOH); UV (ACN) λmax (ε) 206 (4349), 279 (5346) nm; 1H NMR, see Table 3.4; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+H]+ m/z 441.1918 (calcd for C25H29O7, 441.1913). Dhilirolide K (3.19): Isolated as an amorphous white solid; [α]20D -2.26 (c 0.09, MeOH); UV (ACN) λmax (ε) 206 (883), 249 (978) nm; 1H NMR, see Table 3.4; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 479.1679 (calcd for C25H28O8Na, 479.1682). Dhilirolide L (3.20): Isolated as clear prism-like crystals; mp 271–273 °C [α]20D  +182 (c 0.07, ACN) UV (ACN) λmax (ε) 218 (2315) nm; 1H NMR, see Table 3.4; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 463.1745 (calcd for C25H28O7Na, 463.1733). Dhilirolide M (3.21): Isolated as a amorphous white solid; [α]20D  +59 (c 0.02, MeOH); UV (ACN) λmax (ε) 197 (4526), 273 (132) nm; 1H NMR, see Table 3.4; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 463.1740 (calcd for C25H28O7Na, 463.1733). Dhilirolide N (3.22): Isolated as an amorphous white solid; [α]20D -1.4 (c 0.05, MeOH); UV (ACN) λmax (ε) 215 (2285) nm; 1H NMR, see Table 3.4; 13C NMR, see Table 3.3; positive ion HRESITOFMS [M+Na]+ m/z 477.1521 (calcd for C25H26O8Na, 477.1525). 3.10.3 Stable Isotope Feeding Study  For the labeling study 1 g of [1,2-13C2]-acetate was dissolved in 20 mL of sterile distilled H2O and filtered through a 0.25-µm filter under sterile conditions. This solution was then added to 250 mL of autoclaved potato dextrose agar media, which was then divided evenly between 16 Petri dishes and the P. purpurogenum cultured as described above. The resulting culture medium was extracted with EtOAc and dhilirolides A (3.9) (6.2 mg), B (3.10) (1.9 mg), C (3.11) (1.8 mg), D (3.12) (1.7 mg), L (3.20) (3.3 mg) and M (3.21) (2.3 mg) were isolated as described above.   102 3.11 2D NMR Spectra for the Dhilirolides A–N  Figure 3.53 gHSQC NMR spectrum of dhilirolide A (3.9) recorded at 600 MHz in DMSO-d6       103  Figure 3.54 gCOSY60 NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6  Figure 3.55 gHSQC NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6   104  Figure 3.56 gHMBC NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6  Figure 3.57 tROESY NMR spectrum of dhilirolide B (3.10) recorded at 600 MHz in DMSO-d6   105   Figure 3.58 gCOSY60 NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4  Figure 3.59 gHSQC NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4   106  Figure 3.60 gHMBC NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4  Figure 3.61 tROESY NMR spectrum of dhilirolide C (3.11) recorded at 600 MHz in MeOD-d4    107  Figure 3.62 gHSQC NMR spectrum of dhilirolide D (3.12) recorded at 600 MHz in DMSO-d6  Figure 3.63 gCOSY 60 NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6   108  Figure 3.64 gHSQC NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6  Figure 3.65 gHMBC NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6    109  Figure 3.66 tROESY NMR spectrum of dhilirolide E (3.13) recorded at 600 MHz in DMSO-d6  Figure 3.67 gCOSY 60 NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6   110  Figure 3.68 gHSQC NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6  Figure 3.69 gHMBC NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6   111  Figure 3.70 tROESY NMR spectrum of dhilirolide F (3.14) recorded at 600 MHz in DMSO-d6  Figure 3.71 gCOSY 60 NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6    112  Figure 3.72 gHSQC NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6  Figure 3.73 gHMBC NMR spectrum of dhilirolide G (3.15) recorded at 600 MHz in DMSO-d6   113  Figure 3.74 gCOSY 60 NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6  Figure 3.75 gHSQC NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6   114  Figure 3.76 gHMBC NMR spectrum of dhilirolide H (3.16) recorded at 600 MHz in DMSO-d6  Figure 3.77 gCOSY 60 NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6   115  Figure 3.78 gHSQC NMR spectrum of dhilirolide I (3.17) recorded at 600 MHz in DMSO-d6  Figure 3.79 gCOSY 60 NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6   116  Figure 3.80 gHSQC NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6  Figure 3.81 gHMBC NMR spectrum of dhilirolide J (3.18) recorded at 600 MHz in DMSO-d6   117  Figure 3.82 gCOSY 60 NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6  Figure 3.83 gHSQC NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6   118  Figure 3.84 gHMBC NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6  Figure 3.85 tROESY NMR spectrum of dhilirolide K (3.19) recorded at 600 MHz in DMSO-d6   119  Figure 3.86 gCOSY 60 NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6  Figure 3.87 gHSQC NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6   120  Figure 3.88 gHMBC NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6  Figure 3.89 tROESY NMR spectrum of dhilirolide L (3.20) recorded at 600 MHz in DMSO-d6   121  Figure 3.90 gCOSY 60 NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6  Figure 3.91 gHSQC NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6    122  Figure 3.92 gHMBC NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6  Figure 3.93 tROESY NMR spectrum of dhilirolide M (3.21) recorded at 600 MHz in DMSO-d6   123  Figure 3.94 gCOSY 60 NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6  Figure 3.95 gHSQC NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6   124  Figure 3.96 gHMBC NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6  Figure 3.97 tROESY NMR spectrum of dhilirolide N (3.22) recorded at 600 MHz in DMSO-d6   125 Chapter 4: Xestolactone, Xestosaprol O and Known Xestoquinones Identified as Potent Indolamine 2, 3-Dioxygenase (IDO) Inhibitors 4.1 IDO Indolamine 2, 3-dioxygenase (IDO) is a heme-containing enzyme responsible for the catalysis of the rate-limiting step in oxidative cleavage of the indole ring present in L-tryptophan (4.1).141 IDO was originally identified, subsequently purified, and characterized in 1978 by Higuchi, Yamamoto, and Hayashi.141 In 1985, the distribution of IDO in the human body was described. Even though the enzyme is a heme containing protein, it shows little to no sequence homology with other human heme containing proteins such as hemoglobin, myoglobin, or cytochrome p-450s.141 The enzyme active site is largely hydrophobic and contains the heme iron in its ferrous (Fe(II)) activated state.141, 142 Though this enzyme has been studied extensively, the precise mechanism for indole ring cleavage is a contentious subject in the literature.143, 144  Presented in Figure 4.1 are the two most widely supported mechanisms for IDO-catalyzed indole cleavage.143, 144 Figure 4.1 Two plausible mechanisms for indole cleavage by IDO.143, 144 Human IDO (hIDO) plays a role in the de-novo production of N-formylkynurenine (4.2, Figure 4.2) and subsequently nicotinamide adenine dinucleotide (NAD) by the body (Figure 4.2).141, 142 This enzyme has been implicated in immunoregulation within the body, and hIDO is found in most tissue types.141, 145 The highest expression of IDO can be found in the lungs, cecum, colon, epididymis and placenta.  IDO production by the placenta has been shown to be essential for embryonic development and implantation.  Fetal production of IDO acts as a NH R OOB NR O O NH OOR NHR OOOONHR NH O OR NH O NH3COOH NH NH2O OCOOHFeN NN N FeN NN NFeN NN NFeN NN N FeN NN N FeN NN NO H  126 pathway for maternal immunotolerance towards the developing fetus and as such, protects the growing embryo from immune expulsion by the mother.141, 145   Figure 4.2 Tryptophan metabolites and catabolic pathway 141, 146  hIDO has also been implicated in the containment of several pathogenic microbes within the body.  The expression of hIDO near invading cells depletes L-tryptophan (Trp, 4.1, Figure 4.2), an essential amino acid not made by eukaryotes and several prokaryotic pathogens.147 This then prevents the proliferation of the pathogens by slowing protein synthesis.  Coupled to this is the fact that several of the Trp (4.1) catabolic products are proposed to be toxic to certain H2N COOHNHIDOTDO H2NNHHO AcHNNHONH2CO2HNH2O NOH CO2HNH2CO2HNHCHOO NH2CO2HNH2OOH OH NH2CO2H CO2HNH2HO2CCHON CO2HCO2HN CO2H NAD+NH2, CO2,H2OL-tryptophan ( 4.1 ) seratonin melatoninN -formylkynurenine ( 4.2 ) kynurenine kynurenic acid3-hydroxykynurenine 3-hydroxyanthranillic acid 2-amino-3-carboxymuronic semialdehydequinolinic acidpicolinic acidor  127 pathogens.147 Human pathogens including, Salmonella typhimurium, Listeria monocytogenes and Toxoplasma gondii, have been shown to illicit an IDO induced response.141, 147  The mechanism of survival through which most cells escape local Trp (4.1) depletion caused by IDO is complex.  In normally functioning cells, Trp (4.1) is transported into the cell and bound to tRNA for protein synthesis by the enzyme tryptophanyl-tRNA-synthetase (TTS).147 Once tryptophanyl tRNA is synthesized, it is protected from degradation by IDO and other catabolic enzymes.  TTS expression and upregulation is driven in keratinocytes, monocytes, epithelial cells and fibroblasts by the same regulators as IDO expression, including the pro-inflammatory cytokines, IFN-γ, and TNF-α.7 Therefore, even in cells with active IDO expression, docking of Trp (4.1, Figure 4.2) to tRNA remains efficient in periods of extracellular Trp depletion caused by IDO.147 4.1.1 IDO in Disease and Immune Evasion  The essential aspects of IDO in the human body are juxtaposed with its role in a large number of human diseases including chronic inflammatory diseases and cancer. In both of these diseases, IDO can play a central role due its effects on the immune system, specifically T lymphocytes (T-cells).147   In normally functioning T-cells from healthy individuals, TTS is not up-regulated to the same degree as in other cell types.  This makes T-cells very sensitive to Trp (4.1, Figure 4.2) depletion in their microenvironments.147 The body uses the IDO Trp depletion mechanism as a means to help retard a T-cell response, such as after a pathogenic invasion.145, 147 In a normal immune response, the activation of IDO is such that IDO Trp depletion does not reach inhibitory levels for T-cells until the pathogen has been eliminated.141, 147 However, aberrant levels of TTS expression in T-cells, causing resilience to an IDO-induced Trp-limited microenvironment, promotes a continuing immune response after a pathogen has been eliminated.  This TTS overexpression has been observed from T-cells in patients with Graves’ disease and rheumatoid arthritis and is implicated in many autoimmune diseases including Lupus.141, 147, 148  T-lymphocytes also provide an important mechanism through which the body maintains and clears aberrant cancerous cells from the body.  The expression of IDO by some tumors has been implicated as a mechanism for immune escape and tumor virulence.149, 150 In 2008, Prendergast implicated the dormant nature of some tumors to a balanced antagonism between   128 uncontrolled cancer growth and the immune system’s T-cell surveillance.150 This balance can be broken by providing selection pressure for the production of tumor immunological escape through mechanisms like expression of IDO.150, 151   IDO has been shown to have a complex and important relationship to immune system regulation. Therefore, small molecule hIDO inhibitors or promoters could be of significant clinical importance to treatment of either cancer or autoimmune diseases.151  4.1.2 IDO Inhibitors The discovery of inhibitors of IDO has been an area of active research for several years starting soon after the implication of IDO as a mechanism of tumor immune evasion.142 Early IDO inhibitors, such as 1-methyl tryptophan (4.3, Figure 4.3) (1-MT, Ki ~ 30µM), were simple derivatives of the enzyme’s native substrate.142 Large-scale screening of synthetic combinatorial libraries has produced a wide variety of synthetic inhibitors such as β-carbolines (e.g., 3-butyl-β-carboline Ki = 3 µM) or phenyl imidazole, which show a higher degree of activity than 1-MT (4.3, Figure 4.3).142 However, due to the large body of preclinical data for 1-MT (4.3, Figure 4.3), it is the first proof of principle IDO inhibitor, and is currently in phase I clinical trials.152 There is a great deal of preclinical data that shows the effectiveness of 1-MT (4.3) in relation to cancer treatment.  One such study showed the combination of 1-MT (4.3) with paclitaxel (1.19, Chapter 1), a well-known tubulin depolymerizing antimitotic used in the clinic.  In this study, the combination significantly enhanced tumor shrinkage over the levels achieved with paclitaxel alone in a mouse breast cancer model (MMTV-Neu mice).151 However, due to a poor inhibition constant, and many off target effects, 1-MT (4.3, Figure 4.3) is likely only a first step in a long road towards finding a viable clinical candidate.142, 153    129  Figure 4.3 Marine natural product inhibitors of IDO and synthetic analogues Due to the promising proof of principle results obtained using 1-MT (4.3), a large number of reports have come forward describing ever more potent IDO inhibitors.142, 150, 154–157 A number of these reports relating to the use of marine derived natural products are from the Andersen lab. To date, the most potent natural product inhibitors of recombinant hIDO in vitro to be reported have been exiguamine A (4.4)(Ki ~ 41 nM)154, 157 and annulin B (4.5)(Ki ~ 120 nM) (Figure 4.3).142, 155 Subsequently, structure activity relationship (SAR) studies found that the indole quinone moiety 4.6 (Figure 4.3) was the essential pharmacophore in the exiguamines.157  A similar SAR study performed on the annulin B (4.5, Figure 4.3) scaffold found the abbreviated scaffold of the novel synthetic inhibitor 4.7 (Figure 4.3) as one of the most potent in vitro inhibitors (Ki ~ 61 nM) reported to date.16 However, in vivo activity of 4.7 (Figure 4.3) in a cell based assay for IDO inhibition proved disapointing.156 Modifications to increase the bioavailability of this compound and related inhibitors should be realized before any clinical evaluation can be considered.156  One issue associated with widespread administration of an IDO inhibitor is the off target effects an inhibitor would incur.141, 142, 158 For instance, if a potent IDO inhibitor was O OHOOOMeO O SAR studies OOO NHnBuOHNHO O NH NH2OONMeNO O SAR studies NH NH2OONMeMeNOONMeNH2CO2H1-methyl-tryptophan ( 4.3 )annulin B ( 4.5 )exiguamine A ( 4.4 ) 4.64.7  130 administered, tumor produced IDO would be inhibited.  However, the patient’s nascent IDO would also be inhibited, possibly causing the deleterious mis-regulation of immune responses.  Therefore, there may be a need for IDO inhibitor therapies that are targeted to the microenvironment directly surrounding a tumor.  Recently, a report detailing the use of a conjugate 1-MT tumor antigen vaccine presented the effectiveness of a targeted therapy.158 The results of this study showed that not only did antigen ligated 1-MT show significant tumor reduction as compared to controls, but also that side effects associated with immunotherapy and free 1-MT (4.3, Figure 4.3) was not encountered.158 This was most notable in gravid mice, where 1-MT (4.3, Figure 4.3) treatment caused miscarriage, but ligated 1-MT-antigen had no effects on fertility.158 From this report it is clear that selective inhibition of tumor presented IDO can be achieved, and use of more potent IDO inhibitors as conjugates would be the next logical step.  4.2 Xestoquinones The xestoquinone class of compounds are proposed meroterpenoids composed of a drimane terpene portion, which has been degraded and oxidized, and a polyketide aromatic portion that in some cases has been reduced to varying degrees.159 The first member of the class, halenaquinone (4.8, Figure 4.4), was isolated in 1983 and at that time was proposed to be of polyketide origin.160 By 1985, xestoquinone (4.9, Figure 4.5), was isolated in the Hirata lab161 and by 1988 several more members of this class were isolated, including the thiazine containing compounds named adociaquinones A (4.11, Figure 4.5) and B (4.12, Figure 4.4).162 This family of compounds is a chemotaxonomic hallmark of the smaller members of the Xestospongia genus (i.e. X. sapra and X. exigua),159 but they have also been found in certain species of the genus Petrosia.163 In 1988, the Harada group accomplished the first total synthesis.164  This total synthesis established the absolute configuration of halenaquinone (4.8, Figure 4.4), and by analogy, many additional members of this family of compounds.164  The proposed biosynthetic origin and overview of biological activity will be presented below.   131  Figure 4.4 All known carbon skeletons, represented by a known natural product, related to the xestoquinone family of compounds.159–163, 165–167 4.2.1 Xestoquinone Biogenesis The isolation of the xestoquinones in the early 1980’s spurred an interest in how these compounds were produced biosynthetically by the sponge. In 1993, Crews et. al. suggested a mixed polyketide terpene biosynthesis of the xestoquinones from an intermediate similar to zonarone (4.13, Figure 4.5),159 a meroterpene produced by brown seaweed in the genus Dictyopteris.168 In 1996, a cyclized and aromatized version of zonarone (4.13, Figure 4.5), cyclozonarone (4.14, Figure 4.5), was isolated from Dictyopteris undulata, giving further support to Crews’s biosynthetic proposal.169 Recently, isolation of neopetrosiaquinone A163 (4.15, Figure 4.5) and orhaloquinone (4.16, Figure 4.5)170, present a clear biogenesis to the xestoquinones from the proposed meroterpenoid building blocks.  Figure 4.5 below shows a proposed biogenesis for O O OOHO OOO NHS NHSO OO OHO O OOO OO NHSO OOxestoquinolide B O O OO OHO O OO OH14-hydroxymethylxestoquinone15-hydroxymethylxestoquinoneN OOO SO3 OHOSO3HHO exiguaquinolhalenaquinone ( 4.8 )xestosaprol N ( 4.10 ) adociaquinone B ( 4.12 )OONO O OO NHNnoelaquinone  132 the xestoquinone class of compounds, with isolated compounds shown to support proposed intermediates. These intermediates are not likely true intermediates in the biogenesis in the strictest sense, but rather offer a logical stepwise route to furnish the final compound.  Figure 4.5 Proposed biogenesis159, 163, 168–170  The fungal natural products epoxyphomalin A (4.17, Figure 4.6) and macrophorin A (4.18, Figure 4.6) also contain several structural moieties found in the xestoquinones and hint at a possible fungal origin.174 Further evidence was provided in 2009, when epoxyphomalin A174 (4.17, Figure 4.6) was isolated from a sponge associated marine fungus (Phoma sp.).  Subsequently, the previously reported compound macrophorin A (4.18, Figure 4.6) was found in 2012 along with two new related compounds from a marine derived Penicillium purpurogenum.175 These compounds closely resemble zonarol and zonarone (4.13, Figure 4.5), and suggest a sponge associated fungal origin for these meroterpenes and by analogy the xestoquinones. OO OOO OO O OOO OOO HOO OOO zonarone (4.13) cyclozonarone (4.14)neopetrosiaquinone A (4.15 ) orhaloquinone (4.16 )O xestoquinone (4.9 )adociaquinone A (4.11)SHNO O[O]cyclization [O]O OO-CO2HO HO-H2OtaurineOHOH [O]  133   Figure 4.6 Fungal produced related compounds.171, 172, 174, 175 4.2.2 Known Xestoquinone Biological Activity The xestoquinones, as a family, have demonstrated strong inhibitory effects on a wide variety of biological targets. For instance, halenaquinone (4.8, Figure 4.4) shows Staphylococcus aureus and Bacillus subtilis inhibitory activity.160 Together with the publication of the structure of xestoquinone, cardiotonic activity was also reported. Subsequently, biological activities associated with these natural products include: antifungal, cytotoxic176 and antimalarial activity.177, 178  Due to the large array of phenotypic responses to these compounds by mammalian cells and microorganisms, several groups have tried to identify their molecular targets.  Members of this class of compounds have shown inhibitory activity against myosin Ca2+ ATPase,179 Cdc25B, MKP-1 and 3 phosphatases, 171, 181 pp60v-src protein tyrosine kinase (though not an inhibitor of all PTKs),159 topoisomerase II, 182 hypoxia-inducible factor-1,165 and Helicobacter pylori glutamate racemase.165  In reports of the biological activity associated with these compounds the naphthoquinone moiety is implicated as the primary pharmacophore. The furan moiety, present in many of these structures, has previously been manipulated or excluded synthetic derivatives with only minor changes in activity.159, 170 The quinone, however, is often implicated in biological activity, due to its’ propensity to act as a Michael acceptor towards a biological nucleophile.170 Coupled to this is a lipophilic conjugated aromatic ring system, allowing these compounds to easily pass through lipid membranes. The combination of these two attributes likely accounts for the diverse biological activity observed from this family of compounds.159, 166   In several assays, adociaquinones A (4.11, Figure 4.4) and B (4.12, Figure 4.5), or members of the family with a thiazine ring system, have been tested alongside xestoquinone (4.9, Figure 4.5) and halenaquinone (4.8, Figure 4.4). The activities associated with these heterocyclic HO OHO OH HO OHO OHmacrophorin A (4.18)epoxyphomalin A (4.17)HO  134 compounds are often strikingly more selective, inhibiting fewer in vivo and in vitro targets.159, 166, 170, 178 This is likely attributable to the thiazine “blocked” quinone in the adociaquinones that prevents nucleophilic attack. An article from the Nagle lab in 2012, reports that 4.11 (Figure 4.4) and 4.12 (Figure 4.5) are strong inhibitors (< 1µM) of hypoxia inducible factor (HIF-1) under iron chelator (chemical hypoxia) induced HIF-1 activation in a T47D cell-based reporter assay.168 The authors suggested that the thiazine ring has an important role in establishing selective biological activity, although the mechanism still remains uncertain.166, 168  4.3 Isolation of Xestolactone A, Xestosaprol O and P As part of an ongoing screening program designed to find novel and potent (<10 µM) IDO inhibitors, a library of marine invertebrate extracts was screened in vitro using purified recombinant human IDO.  The crude methanolic extract of Xestospongia vansoesti showed potent inhibition in the assay. Prompted by the IDO inhibition bioassay data, large portions of freeze-dried X. vansoesti material were exhaustively extracted using MeOH.  The resulting MeOH extracts were combined and dried in vacuo to give a dark red/brown glass. This extract was then partitioned between hexanes and H2O and then between EtOAc and H2O.  The EtOAc partition proved most active in the bioassay and was subsequently separated using LH20 Sephadex eluting with 20% DCM/MeOH. Extensive HPLC purification then provided the known compounds 4.11 and 4.22–4.26 (Figure 4.7) and the novel compounds xestolactone A (4.19), xestosaprol O (4.20), and xestosaprol P (4.21) (Figure 4.7).   135 Figure 4.7 Xestoquinones isolated from X. vansoestii160, 162, 183, 184 4.4 Structure Elucidation of Novel Xestoquinone Analogues 4.4.1 Structure Elucidation of Xestolactone A Xestolactone A (4.19, Figure 4.7) was isolated as dark red optically active crystals that gave an [M+H]+ ion in the HRESITOFMS at m/z 325.1070, appropriate for the molecular formula C19H16O5 (calcd. 325.1076) requiring 12 sites of unsaturation. LRESIMS in MeOD showed the exchange of one deuterium for hydrogen giving the [M+D]+ ion at m/z 327.1, suggesting the presence of a single exchangeable proton.  Analysis of the 1H (Figure 4.8), 13C (Figure 4.9) and gHSQC (Figure 4.32) NMR spectra of 4.19 (Figure 4.7) showed the presence of two carbonyl resonances [C-16: δ 169.0; C-9: δ 169.7], three aromatic methines [C-1: δC 148.7, H-1: δH 8.00 s; C-11: δC 125.8, H-11: δH 8.61 s; C-14: δC 121.1, H-14: δH 8.10 s] and seven quaternary aromatic carbons [C-2: δ 127.8; C-7: δ 147.0; C-8: δ 144.9; C-10: δ 135.8; C-12: δ 126.3; C-13: δ 156.0; C-15: δ 158.6], accounting for seven sites of unsaturation and suggesting that 4.19 is pentacyclic. OO OOHO O O OHOHO OOHOHO OOHOHO1 34 567891011121314161819201 2 9 1315 16171819 116O OOO O O OOHO OHHO OHOHO OHHO OHOOH3HO OOOHO SHNOOSHNOOA B C D E F1 202117104.194.204.214.224.234.114.244.254.26  136  Figure 4.8 1H NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6.   Figure 4.9 13C NMR spectrum of xestolactone A (4.19) recorded at 150 MHz in acetone-d6. Using 1D and 2D NMR data (Figures 4.8, 4.9 and 4.11–4.13) for 4.19 (Figure 4.7) several structural features were immediately apparent.  In the 1H NMR spectrum (Figure 4.8) of 4.19, an oxymethine quartet δ 5.74 (H-17, J = 7.0 Hz) (C-17: δ 79.1) showed correlations in the 2030405060708090100110120130140150160170180190200 ppm  137 gCOSY spectrum (Figure 4.11) to a methyl doublet at δ 1.71 (H-18, J = 7.0 Hz)(Figure 4.10A).  In addition, H-17 (δ 5.74) showed correlations in the gHMBC spectrum (Figure 4.12) to C-18 (δ 20.9), a carbonyl resonance at δ 169.0 (C-16), and a quaternary aromatic resonance at δ 156.0 (C-13).  Strong HMBC correlations from an aromatic methine singlet at δ 8.61 (H-11) and from Me-18 (δ 1.54) to C-17 (δ 79.1) and C-13 (δ 156.0) warranted attachment of methine C-17 (δ 79.1) to C-13 (δ 156.0). With no COSY correlation from an exchangeable resonance to H-17, an ester linkage was proposed from carbonyl C-16 (δ 169.0) to C-17 (δ 79.1) (Figure 4.10 A). HMBC correlations from the aromatic resonance at δ 8.10 (H-14) to four quaternary aromatic resonances at δ 135.8 (C-10), δ 126.3 (C-12), δ 156.0 (C-13) and δ 158.6 (C-15), which were also shown to have correlations from a second aromatic methine δ 8.61 (H-11) suggested the presence of a tetra-substituted benzene ring (Figure 4.10 A). H-11 (δ 8.61) also showed HMBC correlations to a carbonyl resonance at δ 169.0 (C-16), suggesting lactone carbonyl attachment to C-12 (δ 126.3) accounting for two of the four substitutions on the benzene ring. A third substitution was suggested by strong HMBC correlations from H-11 (δ 8.61) to a second carbonyl resonance at δ 169.7 (C-9) implying attachment at C-10 (δ 135.8) (Figure 4.10 A). A methyl singlet resonating at δ 1.54 (H-19) was found to give HMBC correlations to the quaternary carbon resonance at δ 38.7 (C-6), two aromatic quaternary carbon resonances at δ 158.6 (C-15) and δ 147.0 (C-7), and a methylene carbon resonance at δ 29.0 (C-5).  This suggested that the quaternary carbon C-6 (δ 38.7) was directly attached to the previously described tetrasubstituted aromatic ring at C-15 (δ 158.6) as the fourth and final substituent (Figure 4.10 B).   A three carbon spin system, established using gCOSY60 (Figure 4.11) and gHSQC experiments, starting with two diastereotopic methylenes H-5 a/b (δ 2.55/ 2.08; C-5: δ 29.0), H-4 a/b: (δ 2.59/ 2.12; C-4: δ 31.3) and culminating in oxymethine H-3 (δ 5.06; C-3: δ 60.2) was also observed.  H-3 (δ 5.06) showed COSY correlations to the one resonance, δ 4.30 (OH-3), in the proton spectrum that showed no coupling partner in the gHSQC spectrum, confirming hydroxyl attachment at C-3. HMBC correlations from the diastereotopic methylene resonances at δ 2.12/2.59 (H-4a/b) to the carbon resonances at δ 127.8 (C-2) and δ 38.7 (C-6) coupled with HMBC correlations from a second set of diastereotopic methylene resonances δ 2.55/2.09 (H-  138 5a/b) to C-7 (δ 147.0) and C-6 (δ 38.7), suggested a methyl and hydroxyl substituted cyclohexene ring as shown in Figure 4.10 B.     Figure 4.10 Selected HMBC and COSY correlations for xestolactone A (4.19).             OO OH-17(! 5 . 7 4)H-18(! 1 . 7 1)HHH-14(! 8.10) HO H-4B(! 2.12) H-19(! 1 . 5 4)OH-3(! 4.30) OOOHO OHH-1(! 8.00) (! 147.0)(! 144.9)COSYHMBCA BC H H H-4A(! 2.59)C-2(! 127.8)1 3 6 9 1114 1 71 6 1 81 9HH 317131114 12 15H-11(! 8.61)15 10 H4 5 67 152(! 169.7)C-9 (! 169.0)C-16 C-7C-8  139 Table 4.1 1H and 13C NMR chemical shifts for 4.19, 4.20 and 4.21.  xestolactone A(4.19) xestosaprol O (4.20) xestosaprol P (4.21) atom # δ C, type δ H (J in Hz) δ C, typea δ H (J in Hz) δ C, type δ H (J in Hz) 1 148.7, CH 8.00, s 30.5, CH2 a. 3.23 bd  (13.6)  b. 2.34 dd (13.6, 4.0) 147.8, CH 8.13, s 2 127.8, C  66.0, CH 4.10, bm 126.8, C  3 60.2, CH 5.06, bs 27.9, CH2 a. 2.01 m b. 1.72 m 58.0, CH 4.90, m 4 31.3, CH2 a. 2.59, m b. 2.12, m 32.8, CH2  a. 2.28 m  b. 1.75 m 29.7 CH2 a. 2.45, obs b. 1.92, bd (14.7) 5 29.0, CH2 a. 2.55, m b. 2.08, m 40.1, C  27.5, CH2 a. 2.42, m b. 1.79, td, (3.8, 12.6) 6 38.7, C  137.5, C  36.2, C  7 147.0, C  144.1, C  147.1, C  8 144.9, C  177.0, C  143.3, C  9 169.7, C  130.5, C  170.8, C  10 135.8, C  123.9, CH 8.63, s 136.4 C  11 125.8, CH 8.61, s 133.5, C  126.4, CH 8.45, s 12 126.3, C  173.0, C  145.9, C  13 156.0, C  111.1, C  65.8, CH 4.90, m 14 121.1, CH 8.10, s 146.8, C  31.9, CH2 a. 2.28, m b. 2.02, m 15 158.6, C  178.5, C   35.7, CH2 a. 2.77, m b. 2.71, m 16 169.0, C  131.8, C  197.1, C  17 79.1, CH 5.74, q (7.0) 124.7, CH 8.33, s 132.8, C  18 20.9, CH3 1.71, d (7.0) 155.8, C  122.8, CH 8.09, s 19 33.5, CH3 1.54, s 26.2, C 1.49, s 149.4, C  20   39.5, CH2 3.87, m 31.9, CH3 1.38, s 21   48.5, CH2 3.41, m   OH-3  4.30, bd  (4.0)  Not Observed  Not Observed OH-7  /  8.72 s  / OH-13  /  Not observed  Not Observed NH-20  /  9.23, bt (3.4)  /  a. Resonances in column determined from 1D and 2D NMR experiments        140  Figure 4.11 Inset of gCOSY NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6.  Figure 4.12 Insets of olefinic region of the gHMBC NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in DMSO-d6. H-17/ Me-18  H-17/ Me-18  H-17 H-3 H-3-OH H-3/ H-4  H-3/ H-4  H-3/ H-3-OH  H-3/ H-3-OH  H-17/ C-13  H-11 / C-16 H-11/ C-9 H-1/ C-13  H-1/ C-8  H-1/ C-7  H-14/ C-10  H-14/ C-12  H-11/ C-13  H-17/ C-16  H-11/ C-15  Me-19/ C-7  Me-19/ C-15  Me-18/ C-13  H-5a/ C-7  H-4b/ C-2  Me-19/ C-14    141  Figure 4.13 Insets of aliphatic region of the gHMBC NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in DMSO-d6. The remaining aromatic methine resonating at δ 8.00 (H-1) showed strong HMBC correlations to C-3 (δ 60.2) and to a quaternary aromatic carbon resonance at δ 127.8 (C-2), suggesting a Δ1,2 double bond (Figure 4.10 C). Further HMBC correlations from H-1 (δ 8.00) to resonances at δ 147.0 (C-7) and δ 144.9 (C-8) were also observed. With all 19 carbons and 16 protons observed in the 1D NMR spectra of 4.19 (Figure 4.7) accounted for, one additional oxygen atom was required to satisfy the molecular formula suggested by the HRESIMS. An ether linkage between C-1 and C-8 coupled with attachment of C-7 (δ 147.0) to C-8 (δ 144.9) to give a tri-substituted furan and cyclohexenone ring system supplied the pentacycle requisite by the chemical formula, and matched nicely with the observed 1D and 2D NMR and HRESIMS results, shown in Figure 4.10 C. tROESY correlations observed between Me-19 (δ 1.54), H-4a (δ 2.59) and H-5a (δ 2.55) and from H-5a (δ 2.55) to H-3 (δ 5.06) established the relative configuration as shown in Figure 4.10 C.  However, no correlations could be observed which would establish the configuration at C-17. Luckily, xestolactone A (4.19) was found to crystallize from acetone.  Single crystal X-ray diffraction analysis obtained for xestolactone A (4.19, Figure 4.14) confirmed the constitution obtained from NMR analysis and established the absolute configuration of 4.19 as 3S, 6R, and 17S. H-14/ C-17  H-14/ C-6  H-17/ C-18  Me-18/ C-17  Me-19/ C-6  H-4ab + H5ab / C-5  Me-19/ C-5    142  Figure 4.14 ORTEP diagram of 4.19. 4.4.2 Structure Elucidation of Xestosaprol O  Xestosaprol O (4.20, Figure 4.7) was isolated as an optically active yellow amorphous solid that gave the [M+Na]+ ion in the HRESITOFMS at m/z 452.0774, appropriate for the molecular formula C21H19NO7S (calcd. 452.0780) requiring 14 sites of unsaturation. The molecular formula and the 1H and 13C NMR spectra were nearly identical to those reported for xestosaprol N (4.10).185 Analysis of the 1D and 2D NMR data (Figures 4.16–4.19) for 4.20 (Figure 4.15) revealed all of the same structural features present in xestosaprol N185 (4.10, Figure 4.4): a diosphenol [C-5: δ 40.1; C-6: δ 137.5; C-7: δ 144.1, 7-OH: δ 8.72 s; C-8: δ 177.0; C-9: δ 130.5; C-18: δ 155.8], a naphthoquinone moiety [C-9: δ 130.5; C-10: δ 123.9, H-10: δ 8.63; C-11: δ 133.5; C-12: δ 173.0; C-13: δ 111.1; C-14: δ 146.8; C-15: δ 178.5; C-16: δ 131.8; C-17: δ 124.7, H-17: δ 8.33; C-18: δ 155.8], a dioxythiazine ring [NH-20: δ 9.23 bs; C-21: δ 39.5, H-21: δ 3.87 m; C-22: δ 48.5, H-22: δ 3.41 m], and a methyl cyclohexyl ring system [C-1: δ 30.5, H-1a/b δ 3.23 bd (J = 13.6 Hz)/ δ 2.34 dd (J = 13.6, 4.0 Hz); C-2: δ 66.0, H-2: δ 4.10 bm, 2-OH: Not observed; C-3: δ 27.9, H-3a/b: δ 2.01 m/ δ 1.72 m; C-4: δ 32.8, H-4a/b: δ 2.28 m/ δ 1.75 m; C-5: δ 40.1; C-6: δ 137.5; C-19: δ 26.2, H-19: δ 1.49 s].46 However, careful inspection of the 2D NMR data (Figures 4.18–4.19) revealed one noticeable difference between these compounds. The aromatic methine H-17 (δ 8.33) showed strong HMBC correlations to the most de-shielded carbonyl resonance δ 178.5 (C-15), while the other aromatic methine resonance H-10 (δ 8.63) showed HMBC correlations to the diosphenol carbonyl resonance at δ 177.0 (C-8) and the more shielded quinone carbonyl resonance at δ 173.0 (C-12) (Figure 4.19).  This suggests that the orientation of the thiazine ring system present in xestosaprol N185 (4.10, Figure 4.4) was reversed in xestosaprol O (4.20, Figure 4.15). A weak HMBC correlation from the N-H (δ 9.23) on the   143 thiazine ring system to C-15 (δ 178.5), confirms the regiochemistry shown in Figure 4.15 for xestosaprol O (4.20).  This structure is in agreement with the HRESIMS result and the molecular formula. By comparison with known compound xestosaprol N185 (4.10), and interpretation of the tROESY data the absolute configuration of 4.20 (Figure 4.7) was set as 2S and 5R.  Figure 4.15 Structure of xestosaprol O (4.20) showing selected HMBC and COSY correlations  Figure 4.16 1H NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6.  HO OOOHO SNO OHHH C-15(! 178.5)(! 173.0)C-12(! 177.0)C-8C-7(! 144.1) 14 COSYHMBC(! 137.5)C-62 171019 20211 13 NH-20         (! 9.23)18 H-17(! 8.33)H-10(! 8.63)  144  Figure 4.17 13C NMR spectrum of xestosaprol O (4.20) recorded at 150 MHz in DMSO-d6.    Figure 4.18 gCOSY 60 spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6. 20253035404550556065707580859095100105110115120125130135140145150155160165170175180185190195 ppmNH-20/ H-20 NH-20/ H-20 H-20/ H-21 H-2/ H-1a H-2/ H-1b   145  Figure 4.19 gHMBC spectrum expansions of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6. 4.4.3 Structure Elucidation of Xestosaprol P  Xestosaprol P (4.21, Figure 4.7) was isolated as an optically active yellow amorphous solid that gave the [M+H]+ ion in the HRESITOFMS at m/z 339.1234, appropriate for the molecular formula C20H19O5 (calcd. 339.1232) requiring 12 sites of unsaturation. The LRESIMS recorded in MeOD gave the [M+D]+ ion at 342.2 indicating the presence of 2 exchangeable protons in 4.21. The 1D and 2D NMR data showed a great deal of similarity to that of 3, 13-Dideoxo-1, 2, 14, 15-tetrahydro-3, 13-dihydroxyhalenaquinone162 (4.23, Figure 4.7) and differed by only 2 mass units in the HRESIMS. The 1H-NMR (Figure 4.21) of 4.21 showed the presence of three aromatic methine singlets δ 8.13 (H-1), δ 8.45 (H-11), and δ 8.09 (H-18), one more than seen in the 1H NMR spectrum for 4.23.162 The presence of a furan in 4.21, [H-1: δ 8.13; C-1: δ 147.8; C-2: δ 126.8; C-7: δ 147.1; C-8: δ 143.3] in place of the dihydrofuran seen in 4.23 (Figure 4.7) accounts for the difference in two mass units and for the added degree of unsaturation. Through interpretation of the 1D and 2D NMR data of 4.21, the presence of a tetra-substituted aromatic ring [C-10: δ 136.4; C-11: δ 126.4; C-12: δ 145.9; C-17: δ 132.8; C-18: δ 122.8; C-19: δ 149.4] linked through a cyclohexenone ring [C-6: δ 36.2, C-7: δ 147.1; C-8: δ 143.3; C-9: δ 170.8; C-10: δ 136.4; C19: δ 149.4] to the above mentioned tri-substituted furan as in 4.19  C-16 C-18  H-10  H-17 OH-7 C-12 C-8 C-8 C-7 C-6 C-5 C-15 C-11 C-9   146 (Figure 4.7) was established as shown in Figure 4.20. All further features, including the constitution of rings B (Figure 4.21) [C-2: δ 126.8; C-3: δ 58.0, H-3: δ 4.90; C-4: δ 29.7, H-4a/b: a. δ 2.45 obscured/ b. δ 1.92, bd (J = 14.7 Hz); C-5: δ 27.5, H-5 a/b: a. δ 2.42 m/ δ 1.79 td (J = 3.8, 12.6 Hz), C-6: δ 36.2; C-7: δ 147.1] and E [C-12: δ 145.9; C-13: δ 65.8, H-13: δ 4.90 m; C-14: δ 31.9, H-14 a/b: δ 2.28 m/ δ 2.02 m; C-15: δ 35.7, H-15 a/b: δ 2.77 m/ δ 2.71 m; C-16: δ 197.1; C-17: δ 132.8] were found to be identical to those seen in 4.23 (Figure 4.7). The proposed structure 4.21 is consistent with the 1D and 2D NMR data and the HRESIMS result.   Figure 4.20 Xestosaprol P (4.21) showing selected gHMBC and gCOSY correlations   Figure 4.21 The 1H NMR spectrum of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. O OOHOHOHH-1(! 8.13) HHH-18(! 8.09)H-11(! 8.45) C-16(! 197.1)(! 170.8)C-9(! 147.1)C-7C-8(! 144.9) (! 65.8)C-13 COSYHMBC2031 6 12 H H15 H-15a(! 2.77)1019 H-13(! 4.90)  147  Figure 4.22 The 13C NMR spectrum of xestosaprol P (4.21) recorded at 150 MHz in DMSO-d6.  Figure 4.23 gHMBC spectrum expansion of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. H-11/ C-13 H-11/ C-17 H-11/ C-19 H-11/ C-9 H-18/ C-10 H-18/ C-12 H-18/ C-6 H-18/ C-16 H-1/ C-2 H-1/ C-8 H-1/ C-7 H-18/ C-9   148  Figure 4.24 gHMBC spectrum expansion of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6.   Figure 4.25 gCOSY spectrum expansion of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6. H-15ab/ C-16 H-11/ C-16 H-14a/ C-12 H-20/ C-7 H-20/ C-6 H-20/ C-5 H-4b/ C-2 H-14b/ C-16 H-15ab/ C-13 H-14a/ C-13 H-5b/ C-7 H-20/ C-19 H-14a/ C-15 H-4b / H-3 H-14b / H-13 H-14a / H-13 H-4a / H-3 H-14b / H-15b H-14a / H-15a H-4b / H-5b   149  In order to determine the absolute configuration at all of the stereocenters in 4.21 (Figure 4.20), we first employed a tROESY NMR experiment (Figure 4.37).  This experiment unfortunately was unable to establish the configuration at C-13 in 4.21 (Figure 4.7), as no tROESY correlations were observed from any moieties of known configuration (e.g. Me-20) to any of the E ring resonances (Figure 4.37). Furthermore, several attempts at crystallization of 4.21 were unsuccessful. Subsequently, 4.23 isolated as part of this work, was shown to possess an alcohol moiety at C-13. 4.23 (Figure 4.7), was found to crystallize and possessed all stereocenters present in 4.21. Therefore, due to a presumed common biosynthetic origin, 4.23 represents a good model compound for 4.21.162 An X-ray diffraction analysis was undertaken with crystals of 4.23, and an ORTEP diagram obtained (Figure 4.26). After comparison with the ORTEP diagram of 4.23 (Figure 4.26), the absolute configuration of 4.21 (Figure 4.7) can be established as 3S, 6R, and 13R.  Figure 4.26 ORTEP diagram of 3,13-dideoxo-1, 2,14,15-tetrahydro-3, 13-dihydrohalenaquinone (4.23)162 4.5 Proposed Biosynthetic Origin of Xestolactone The lactone ring found in xestolactone A (4.19, Scheme 4.2) is unique within the xestoquinone family of compounds. We propose this moiety could arise biogenetically from degradation and cyclization of a meroterpenoid starting material similar to a reduced version of xestoquinone (Scheme 4.2). First, Bayer-Villiger oxidation at C-13, hydrolysis, oxidation to the acid and decarboxylation would furnish the correct carbon skeleton (Scheme 4.2).  Subsequent   150 oxidation at C-3, reduction of the ketone at C-16, and lactone formation would furnish xestolactone A (4.19).  Figure 4.27 Proposed biogenesis for xestolactone (4.19) carbon skeleton. 4.6 Structure Revision of Adociaquinone A and B Based on Isolated Material  Several known compounds (4.22–4.26) were isolated along with the three new xestoquinone analogues described above (Figure 4.7).  One of these compounds afforded the mass spectral and 1D NMR data matching those of adociaquinone B (4.12, Figure 4.27).  However, the 2D NMR data showed conflicting results as to which regio-isomer, A (4.11) or B (4.12), it represented.  In the first report of these structures by Bloor and Schmitz,162 the  regiochemistry was assigned using proton-carbon NOE data.  Their interpretation gave the carbon chemical shifts shown below (Figure 4.27) for both adociaquinone A (4.11, Figure 4.27) and B (4.12, Figure 4.27).  After detailed analysis of the isolated compound and the reported data, it became evident that the literature assignment of the dioxythiazine ring moiety and OO OOHOO O OO O O OOO O OO OHO O OO OH -CO2O O OOHOH -H2OHydrolysis[O] [O][H] 16 4.19OO OH13[O] 3  151 quinone differed for those we assigned to those positions.162 Based on our 2D NMR results, it is clear that original interpretation for these two structures should be revised so that the chemical shifts reported for adociaquinone A (4.11) are now those reported for adociaquinone B (4.12) and vice versa (Figure 4.27).  Figure 4.28 1H NMR spectrum of adociaquinone A (4.11) recorded at 600 MHz in DMSO-d6.  Figure 4.29 13C NMR spectrum of adociaquinone A (4.11) recorded at 150 MHz in DMSO-d6   152  Figure 4.30 Structure assignment of isolated 4.11 and the reported 13C NMR chemical shifts of 4.11 and 4.12.162  O OOO NS OOHHHHO O OO HNS OO! 39.7!  48.3 H HNH-21(! 9.23)!  147.0!  110.7!  178.4! 173.0H-18(! 8.26)H-11(! 8.70)! 138.1 ! 131.7!  169.3 ! 130.9! 154.2! 147.3! 146.2!121.6 !143.0 ! 122.0 !125.3! 146.2 ! 37.2! 147.1!  142.9 ! 169.3! 135.9 ! 125.4! 128.5 ! 177.8 ! 111.7! 147.8! 173.7! 134.4! 122.9!  157.4! 31.8 ! 48.2! 40! 146.1 ! 37.2! 147.1! 143.1 !  169.4! 137.9 ! 124.8! 131.8 ! 173.7 ! 111.4! 147.9! 178.3!  130.9! 123.4! 154.6!  31.6 ! 48.2! 40O O OO NHSO Oadociaquinone B (4.12)adociaquinone A (4.11)Isolated adociaquinone A (4.11)! 37.3H-1(!  8.01) H-21     (!  3.82)  153   Figure 4.31 gHMBC NMR spectrum of adociaquinone A (4.11) recorded at 600 MHz in DMSO-d6  4.7 The Xestoquinones as Inhibitors of IDO Compounds 4.11, 4.19, 4.20, and 4.22–4.26 were all assayed for in vitro inhibition of purified recombinant human IDO (Figure 4.31). It was not possible to assay 4.21 (Figure 4.7), as sufficient amounts could not be obtained for the assay.     154        Figure 4.32 Nonlinear regression curves were based on a sigmoidal dose-response equation and used to calculate the corresponding IC50 values shown in the table. The data are shown as an average of quadruplicates with error bars representing SD. 4.8 Conclusion  From the extract of Xestospongia vansoesti, three new members of the xestoquinone family of compounds 4.19, 4.20 and 4.21 (Figure 4.7) were isolated.  The carbon skeleton for xestolactone A (4.19) has only been observed once before in noelaquinone (Figure 4.4), which we propose to be the product of degradation and reduction of a precursor similar to xestoquinone (4.8, Figure 4.4).  Isolation of xestosaprol O (4.20) and re-isolation adociaquinone A (4.11) (Figure 4.7) illuminated a discrepancy in the literature surrounding the regiochemistry of the thiazine moiety on the quinone ring.  Sigmoidal dose-response-2 -1 0 1 2 3050100150Xesto-1Xesto-2Xesto-3Xesto-4Xesto-5Xesto-6Xesto-7Xesto-8 log [inhibitor] ( µ M)[Kynurenine] (µM)4.23!4.20!4.19!4.22!4.11!4.27!4.26!4.25!Compound IC50 4.23 3,13-Dideoxo-1, 2, 14, 15-tetrahydro-3, 13-dihydroxyhalenaquinone     123 ± 1 µM 4.26 xestosaprol B 127 ± 1 µM 4.22 13, 14, 15, 16- tetrahydroxestoquinol 6 ± 1 µM 4.11 adociaquinone A 2 ± 1 µM 4.25 xestosaprol A 41 ± 1 µM 4.26 xestosaprol D 12 ± 1 µM 4.19 xestolactone A 81 ± 1 µM 4.20 xestosaprol O 4 ± 1 µM   155  In vitro testing of these compounds (Figure 4.31) against recombinant hIDO has revealed that adociaquinone A (4.11) and xestosaprol O (4.20) are potent inhibitors of hIDO. Due to the activity observed from these compounds, it is likely that the furan present in adociaquinone A (4.24) is not necessary for IDO inhibition. It is also likely that the alcohol at C-3 in xestosaprol O (4.20) has led to a reduced binding affinity for xestosaprol O (4.20) compared to adociaquinone A (4.11) (Figure 4.7).  Further support can be seen in the drop in hIDO inhibition observed between 4.26 (IC50 = 12 µM) and 4.25 (IC50 = 41 µM) (Figure 4.31) where the only difference in structure is an alcohol at the C-3 in 4.25 (Figure 4.7). Due to the potent hIDO inhibitory activity of xestosaprol O (4.20, Figure 4.7) and adociaquinone A (4.11, Figure 4.7), a synthetic supply of these compounds or simplified analogues would be of great value for continued testing and future clinical development.   4.9 Experimental Section 4.9.1 IDO Inhibition Assays Enzyme activity assays were performed with human recombinant IDO (rh-IDO) expressed in E. coli as previously described.186 The activity of rh-IDO in the absence and presence of inhibitory compounds was determined using an end-point assay as previously described187 with the following changes: The assay was performed in a 100 mM potassium phosphate buffer (pH 6.5), 10 mM sodium ascorbate (Sigma), 1.25 µM methylene blue (Sigma), 10 µg/mL catalase (Sigma), 400 µM L-tryptophan (Sigma). The reaction was started by the addition of rh-IDO (100 nM) and allowed to progress at 37 °C for 60 min before termination with 30% (w/v) trichloroacetic acid. The samples were further incubated for 15 min at 60 °C prior to addition of 2% (w/v) 4-(dimethylamino)benzaldehyde (Sigma). After 5 min at room temperature the absorbance at 480 nm was measured with an infinite M200 TECAN plate reader, and the kynurenine concentration was determined from the extinction coefficient (15820 M-1cm-1) for kynurenine.188 Nonlinear regression of the enzyme activity assays and calculations of IC50 values were performed using GraphPad Prism 4. 4.9.2 General Experimental Procedures Melting points were taken using a Fisher-Johns apparatus, and the reported values are uncorrected. Optical rotations were measured using a Polarimeter with sodium light (589 nm). UV spectra were recorded with a Dual λ Absorbance Detector. The 1H and 13C NMR spectra   156 were recorded on a 600 MHz spectrometer with a 5 mm cryoprobe. 1H chemical shifts are referenced to either residual DMSO-d6 (δ 2.49 ppm), acetone-d6 (δ 2.05 ppm) or MeOD-d4 (δ 3.31 ppm) and 13C chemical shifts are referenced to the DMSO-d6 (δ 39.5 ppm) or acetone-d6 (δ 29.92 ppm) solvent peak. Low and high resolution ESI-QIT-MS were recorded on a LC system mass spectrometer. Merck Type 5554 silica gel plates and Whatman MKC18F plates were used for analytical thin layer chromatography. Reversed-phase HPLC purifications were performed on a Binary HPLC Pump attached to a Dual λ Absorbance Detector. All solvents used for HPLC were Fisher HPLC grade and were filtered through a 0.45 µm filter prior to use.   4.9.3 Extraction and Isolation  The freeze-dried sponge sample (300 g) was extracted exhaustively with MeOH (3 x 200 mL). The combined MeOH extracts were concentrated in vacuo to afford a dark red/brown glass. The extract was partitioned between H2O (150 mL), and hexanes (3 x 75 mL) and then H2O and EtOAc (3 x 75 mL) and the combined EtOAc extract evaporated under reduced pressure to give a dark red/ brown solid (500 mg).  The EtOAc soluble material was then subjected to Sephadex LH20 chromatography in MeOH: DCM (4:1) yielding 5 fractions based on NP-TLC. The second fraction (43mg), which was most active in the bioassay, was subsequently subjected to rigorous RP-HPLC.  Initial separation of the active fraction using a C8 reversed-phase HPLC using a Phenomenex Luna C8 100 Å, 5 µm 250 × 10 mm, column eluting with 38% ACN/H2O gave a complex mixture from 6-12 min which were further separated as described below, and purified xestosaprol B (4.27) (2.1 mg, 13.5 min), 13, 14, 15, 16-tetrahydroxestoquinol (4.22) (0.9 mg, 14.2 min), xestolactone A (4.19) (2.4 mg, 17 min), xestosaprol D (4.26) (2.3 mg, 24 min), adociaquinone A (4.11) (1.9 mg, 27 min), 3, 13-Dideoxo-1, 2, 14, 15-tetrahydro-3, 13-dihydroxyhalenaquinone (4.23) (1.7 mg, 33.5 min), xestosaprol A (4.25) (1.1 mg, 34.5 min) and partially purified xestosaprol O (4.20) (12 min). Subsequently the fraction representing 6–12 min was subjected to RP-HPLC at 20% ACN/H20, giving xestosaprol O (4.20) (0.6 mg, 31.5 min). Xestosaprol P (4.21) was purified via C18 reversed-phase HPLC using a CSC-Inertsil 150A/ODS2, 5 µm, 25 x 0.94 cm column, eluting with 20% ACN/H2O at 41.5 min (0.7 mg).   157 Xestolactone A (4.19): Dark red crystals from acetone; mp 182 ºC decomp; [α]20D -23.5° (c 0.46  MeOH); UV (MeOH) λmax (ε) 234 (7600), 308 (3900) nm; 1H NMR and 13C NMR, see Table 4.1; positive ion HRESIMS [M+H]+ m/z 325.1070 (calcd. for C19H17O5, 325.1076). Xestosaprol O (4.20): Amorphous yellow solid; [α]20D -6.5° (c .001, MeOH); UV (MeOH); λmax (ε) 212 (3528), 278 (3430) nm ; 1H NMR and 13C NMR, see Table 4.1; positive ion HRESIMS [M+Na]+ m/z 452.0774 (calcd. for C21H19NO7SNa, 452.0780). Xestosaprol P (4.21): Amorphous brown solid; [α]20D -26.4° (c .014, MeOH); UV (MeOH) λmax (ε) 206 (1580), 268 (1140), 313 (1230) nm; 1H NMR and 13C NMR, see Table 4.1; positive ion HRESIMS [M+H]+ m/z 339.1234 (calcd. for C20H19O5, 339.1232). 4.10 2D NMR Spectra  Figure 4.33 gHSQC NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6.  ppm1.01.52.02.53.03.54.04.55.05.56.06.57.07.58.08.59.0 ppm160150140130120110100908070605040302010  158  Figure 4.34 tROESY NMR spectrum of xestolactone A (4.19) recorded at 600 MHz in acetone-d6  Figure 4.35 gHSQC NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6   159  Figure 4.36 gHMBC NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6  Figure 4.37 tROESY NMR spectrum of xestosaprol O (4.20) recorded at 600 MHz in DMSO-d6   160  Figure 4.38 gHSQC spectrum of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6  Figure 4.39 tROESY spectrum of xestosaprol P (4.21) recorded at 600 MHz in DMSO-d6.    161 Chapter 5: Synthesis of Analogues of Xestosaprol N and the Adociaquinones; Structure Activity Relationship Studies and Novel Inhibitors of Indolamine 2,3-Dioxygenase. 5.1  Previous Syntheses of Xestoquinone and Analogues  Since their discovery, the xestoquinones have attracted a great deal of attention from synthetic organic chemists due to their fused polycyclic structure and various biological activities. The first total synthesis in 1988 by the Harada group164 provided (+)-halenaquinone (4.8, Figure 5.1) and (+)-halenaquinol starting from optically pure (8 a R)-(–)- Wieland-Miescher ketone in 14 and 15 steps respectively.  Shortly thereafter, Harada provided the first total synthesis of (+)-xestoquinone (4.9, Figure 5.2) and (+)-xestoquinol, this time in 13 steps. Along with their original isolation, adociaquinone A (4.11, Figure 5.1) and B (4.12, Figure 5.1) were synthesized from isolated xestoquinone (4.9, Figure 5.2) as the starting material by Bloor et al.162 The only total synthesis of adociaquinone A (4.11, Figure 5.1) and B (4.12, Figure 5.1) was published in 1995 by Harada et al., and it relied on the methodology used by Bloor et al. to install the thiazine ring system onto synthetic xestoquinone (4.9, Figure 5.2).162, 189  Figure 5.1 Several previously synthesized members of the xestoquinone family of compounds.162, 164, 189–195 To date, nine total syntheses of members of the xestoquinone family have been accomplished.164, 190–198 The majority of these were completed in the 1990’s and focused on xestoquinone (4.9, Figure 5.2) as the target compound.  The most efficient synthesis in terms of yield and number of synthetic operations is the eight step synthesis by R. Carlini et al., which furnished (±)-xestoquinone in 7.4% overall yield.195 This synthesis began with a (Bis(trifluoroacetoxy)iodo)benzene (PIFA) mediated addition of (E)-2,4-pentadienol into 2-methoxy-4-methylphenol as shown in Figure 5.2. Next, intramolecular Diels-Alder reaction with the monoketal formed the A, B, and C rings of xestoquinone.  The D and E rings were O OOO adociaquinone A (4.11)SHNO O O O OO NHSO Oadociaquinone B (4.12)O OOhalenaquinone (4.8 )OO  162 subsequently formed through a second Diels-Alder reaction with 4, 7-dimethoxyisobenzofuran (Figure 5.2).195  Figure 5.2 R. Caralini synthesis of (±)-xestoquinone.195 5.2 Synthesis of Xestosaprol N Analogues Many of the approaches discussed above hinged on the construction of the furan moiety present in both the adociaquinones and xestoquinones.190–198 Taking into account the SAR relationship revealed by the analogues isolated and tested in the IDO assay (shown in chapter 4), an abbreviated pharmacophore (5.1, Figure 5.3) found in xestosaprol O (4.20) (Figure 5.3) became the focus of my synthetic efforts.  Figure 5.3 Xestosaprol O (4.20) and synthetic target (5.1) Modification of several previously reported synthetic methodologies precluded themselves as possible routes towards the synthesis of this abbreviated scaffold.190–197 This was largely due to the fact that many of the previous syntheses focused on the formation of the furan present in the xestoquinones, which is absent in our synthetic target.190, 193–197 Therefore, it became clear that there was a need to develop our own methodology in order to reach our desired target (5.1, Figure 5.1) efficiently.  5.2.1 Photocyclization Towards the Carbon Framework of Xestosaprol O  Examination of the literature revealed two papers by Sato et al., detailing a light catalyzed silver cation mediated cyclization process,199, 200 which could furnish direct access to a significant portion of the xestosaprol carbon skeleton in one synthetic operation (Figure 5.4). OH OMe O OO O O O HH OOO O OOOxestoquinone (4.9)+PIFAOOO OHOHxestosapro l O ( 4.20 )S OO NH OOO OHR1R R= NH, R 1=SO 2R= SO 2, R 1=NH5.1  163 Though the photochemical process appeared to be a novel reaction, and offered entry into a wide variety of terpenoid natural product scaffolds, no application of this methodology towards natural product synthesis could be found in the literature.  Figure 5.4 Silver and light promoted cyclization reported by Sato and Tamura.199, 200 5.2.2 Retrosynthetic Analysis A retrosynthetic analysis of the desired end product is presented in Figure 5.5. The first disconnection relies on the formation of the thiazine ring from hypotaurine, followed by a proposed ceric ammonium nitrate oxidation of the dimethoxyquinol to the quinone and O2 oxidation to the diosphenol. The final disconnection in this analysis relies on the Sato methodology shown in Figure 5.4, to give an intermediate like 5.7 and the commercially available methyl cyclohexene (5.3, Figure 5.5).200  Figure 5.5 Retrosynthetic analysis 5.2.3 Initial Synthetic Trials To begin our synthesis we needed to generate an intermediate similar to 5.7 (Figure 5.5). From the literature, we found methodology to generate 5.9 and 5.10 (Scheme 5.1) by Friedel-Crafts acylation of 1, 4-dimethoxynaphthalene (5.8) with acetic anhydride using aluminium chloride.201 The acylation gave the opposite regio-chemical outcome 5.9 (Scheme 5.1) as the O Cl AgOTfbenzene, h! O H+5.2 5.3 5.4O XOOOOOOOR1R OH OH OOOOR= NH, R1=SO2R= SO2, R1=NH5.1 5.5 5.65.7 5.3  164 primary product and the desired regio-isomer 5.10 as the minor product.200 Subsequent attempts at α-keto halogenation under various conditions failed, giving only the electrophilic aromatic substitution products.  Acylation of 5.8, with chloroacetic anhydride, provided the desired product 5.11 (Scheme 5.1) in poor yields. Attempts at the photo-cyclization reaction with 5.11 provided a complex mixture of products, none of which resembled the desired tetracycle 5.6 (Figure 5.5).    Scheme 5.1 1, 4-dimethoxynaphthalene trials. We then attempted the Sato methodology using commercially available 2-bromo-2'-acetonaphthone (5.13) (Scheme 5.2) as starting material, only to produce intractable mixtures of undesired products. Subsequently, chromate oxidation was used to access 2-bromo-2'-acetonaphthoquinone (5.14, Scheme 5.2). This proved no different as a starting material in the photocyclization reaction, giving complex mixtures of undesired products.  Finally, the use of 5.15 (Scheme 5.2) gave promising results, ultimately providing a product 5.16 (Scheme 5.2) in low (~26%), but consistent yields.  Though 5.16 (Scheme 5.2) was reduced as compared with 5.6 (Figure 5.5), it did possess the desired carbon framework needed in the final product. The decision was made to carry forward with 5.16 as starting material for the rest of the synthesis. OO O OO OO OO OOAlCl3,  DCE, 0 ° C – 60 ° C5.8 5.9 5.10+( 3 :1) (3:1)OO O ClOO O ClOOClAlCl3,  DCE, 0 ° C – 60 ° C OO O Cl+5.8 5.11 5.12  165  Scheme 5.2 Successful photocyclization reactions and non-productive naphthyl starting materials 5.2.4 Halogen Effects on Photocyclization The decision to use bromo-acetophenone derivatives, as compared with the chloro-acetophenones used by Sato et al. (Figure 5.4), was originally made due to higher reported yields for bromination versus chlorination of acetophenone.200, 202, 203 The low yield of the cyclization caused us to revisit the halogen substituent effects on the reaction, as there was no report by the authors in the original paper regarding this issue.  The model compound 3’, 4’-dimethylacetophenone was subsequently chlorinated 5.17A, brominated 5.17B, and iodinated 5.17C (Scheme 5.3).  These three compounds were then compared in the cyclization reaction to identify the optimal halide substituent for the reaction.   When comparing the reaction mixtures we found that the amount of side products, mainly 3’, 4’-dimethylacetophenone, appeared to increase from chloro- to iodo-substitution. The iodo-dimethylacetophenone 5.17C gave the most 3’, 4’-dimethylacetophenone, apparently to the detriment of product yield (~19%) (Scheme 5.3).  However, little difference in product yield was OO BrOMeOMe O ClBrO No ProductNo ProductBrOOO No Product5.115.135.145.15 5.165.3 , AgOTf benzene , h!5.3 , AgOTf benzene , h!5.3 , AgOTf benzene , h!5.3 , AgOTf benzene , h!, 26%  166 observed between the 5.17A (35%) and 5.17B (33%). We surmised that the increase in byproducts correlated with a decrease in energy required for homolysis (C-Cl > C-Br > C-I) as a rationale for this trend.203 Weighing yield differences in bromination versus chlorination of the acetophenone starting material (5.25, Scheme 5.6), we decided to carry forward with the bromo acetophenone 5.15 (Scheme 5.2).   Scheme 5.3 Halogen effects on photocyclization 5.2.5 Synthetic Efforts Continued  Oxidation of 5.16 (Scheme 5.4) at the benzylic position using KMnO4 in ACN gave the two regio-isomers 5.19 and 5.20 (Scheme 5.4) in a near 1:1 mixture.  An interesting note is that KMnO4-mediated oxidations using either MgSO47H2O or MnSO4H2O as additives in acetone, provided the same yields, but gave a slight excess of 5.20.204, 205 The ketones 5.19 and 5.20 (Scheme 5.4) were easily separated by silica gel flash chromatography and subsequently oxidized under an O2 atmosphere with KOtBu in tBuOH to furnish 5.21 and 5.22 (Scheme 5.4), containing a hydroxyquinone and diosphenol in a single step.207 The oxidation of 5.19 and 5.20 to the hydroxyquinone as compared to the putative intermediate 5.5 (Figure 5.5) was seen as an advantage, allowing for regiospecific addition of hypotaurine onto the quinone. Interestingly, ketone 5.19 (Scheme 5.4) in tBuOH gave two products, the quinone 5.5 (Scheme 5.4) and the hydroxyquinone 5.21 in a 1:1 ratio from the same oxidation.  This is likely due to similarities in pKa values for the protons at the benzylic position and the protons alpha to the carbonyl (Scheme 5.4). Deprotonation, then oxidation could furnish the quinone 5.5 as a competing reaction product as shown in Scheme 5.5. The quinone (5.5) was not detected in the reaction mixture from ketone 5.20 (Scheme 5.4), likely stemming from less resonance stabilization for benzylic proton abstraction.   Using glyme as solvent at –10 °C minimized 5.5 formation, to give mainly 5.21 (Scheme 5.5), though formation of other uncharacterized side products still occurred. O XX= Cl  5.17A      Br  5.17B      I    5.17C O5.18  From   A = 35 %                     B = 33 %                     C = 19 %5.3, AgOTf benzene, h!  167  Scheme 5.4 Oxidations to form quinone products  Scheme 5.5 Two proposed reaction pathways leading to 5.21 and 5.5 Once both hydroxyquinones were in hand, addition of hypotaurine to construct the thiazine ring present in 5.1 (Figure 5.3) was attempted. The addition of the amine of hypotaurine regiospecifically onto the hydroxyl-bearing carbon of the hydroxyquinone moiety in 5.21 and 5.22 (Scheme 5.6) required much harsher conditions than are usually employed for this OO KMnO4 ACN, 40%O OOOOOHO OOO OOOHOKOt-Bu, O2t-BuOH, 43% KOt-Bu, O2t-BuOH, 39%OH OH OH5.165.19 5.205.21 5.5 5.22glyme-10 ° C41%KOt-BuOOHH H H OO OO OOO OHOOOHO OH5.19 5.215.5O OO OO t- BuO t- Bu  168 transformation on an un-substituted quinone (i.e. adociaquinone A 4.11 and B 4.12 from xestoquinone 4.9, Figure 5.1 and 5.2).189 Refluxing temperatures, an oxygen-enriched atmosphere, and a catalytic amount of TFA were employed to reduce the reaction time (Scheme 5.6) and give the final products 5.23 and 5.24 (Scheme 5.6) in moderate yields (~40%).  Scheme 5.6 Thiazine formation. Throughout the synthesis, the presence of highly oxidized products (Figure 5.6) reduced yields of all oxidative reactions in the sequence.  Though many of these were present as intractable mixtures, a large amount of a lactol was consistently formed in every oxidative step performed. Shown in Figure 5.6 are two examples of the lactol side products 5.26 and 5.27, which decreased yields in this synthesis.  Figure 5.6 Representative over oxidized side products This synthetic route offers several advantages over known routes into the scaffold of the xestosaprols and adociaquinones.  First, it is short; five formal synthetic steps from inexpensive commercially available reagents. Second, the reactions performed require relatively innocuous conditions and reagents. Lastly, the synthetic route offers flexibility, providing access to many analogues of the xestoquinone family with varying degrees of oxidation or quinone substitution. All of these factors make this route a useful tool to efficiently provide material for further OOO OHHO H2N SO 2HACN/ EtOH/ H 2OTFA, O 2, 45% OOO OHSNHO O5.22 5.24OOOHO OH H2N SO 2HACN/ EtOH/ H 2OTFA, O 2, 42% OOO OHHNSO O5.21 5.23OO OHO 5.27 OOOOHO5.26 OH  169 evaluation of the hIDO inhibitory abilities of these compounds.  Preliminary testing involving these analogues, and the other analogues outlined below, is presented in sections 4 and 5 of this chapter.  Scheme 5.7 Synthetic route overview 5.3 Further Synthesis of Xestoquinone Analogues Along with the above synthetic approach towards xestosaprol N and O analogues 5.24 and 5.23, several other compounds were synthesized in order to determine the minimal pharmacophore of the adociaquinones. In addition to this, we also ventured to assess the minimal pharmacophore of tetrahydroxestoquinol (4.22, Figure 5.7) through SAR as well. The proposed synthetic targets are shown in Figure 5.7.  The synthetic efforts towards all of these compounds will be presented with the exception of 5.31, which has been synthesized previously.207 O O BrNH4BrOxone!  , MeOHreflux, 3 hrs OOO OOO2, t-BuOKrt, 4hrsOOOHOOOOHOOOOHNSOO OOOSNH OHOHOHOHO O ACN/ EtOH/ H2O, reflux, TFA3 daysH2N SO2Hrt, 24 hrst-BuOHO2, t-BuOK-10 ° C, 4 hrsglyme5.15 5.165.3, AgOTfbenzene, h" +5.25 5.215.225.245.23 5.195.20KMnO4 ACN  170  Figure 5.7 Proposed abbreviated synthetic targets and tetrahydroxestoquinol (4.22) 5.3.1 Attempted Synthesis of 5.28 and 5.29 We proposed compound 5.28 (Figure 5.7) as a model to test the necessity of the quinone present in 5.23 and 5.24 (Scheme 5.7) for inhibition of IDO.  Construction of 5.28 again relied on photocyclization methodology shown previously to give the known compound 5.4 (Figure 5.4).  Any number of oxidative methodologies could then conceivably give the diosphenol product 5.28 (Scheme 5.7).  This was supported in the literature by the isolation or synthesis of several simple diketo or diosphenol abitane terpenoids, such as 6-hydroxysalvinolone (5.34) and 6-hydroxy-5,6-dehydrosugiol (5.35) and xanthoperol (5.36) (Figure 5.9).209–211 Figure 5.8 Diosphenol and diketone abitane diterpenoids  Various oxidation conditions were attempted in order to form an alpha diketone, which would spontaneously tautomerize to the diosphenol (5.28, Scheme 5.7). We first tried oxygen-mediated oxidations analogous to the methodology used earlier (Figure 5.6), only to receive lactol 5.38 (Scheme 5.7). Chromate oxidation also gave no sign of the desired diosphenol, giving starting material and complex mixtures of undesired products. Formation of the α,β-unsaturated ketone (5.29) (Scheme 5.8) using IBX mediated dehydrogenation, or the acyloin (5.39) (Scheme OOHOH OHO OH HNS OOOOOS OOOO ONHS OOO ONH O5.28 5.29 5.305.31 5.32 5.33 OOOHOH 4.22OHOOH6-hydroxy-5,6-sugiol( 5.35 ) OHOOxanthoperol( 5.36 )OHOOHHO H6-hydroxysalvinolone( 5.34 ) O OHOOHcupresol ( 5.37 )  171 5.8) through Rubottom oxidation were easily achieved, but all attempts at oxidation of the acyloin (5.39) gave intractable mixtures of byproducts and lactol 5.38 (Scheme 5.8).212 Trost-Riley oxidations were also attempted with selenium dioxide, but only 5.40 (Scheme 5.8) could be isolated from the complex reaction mixture.213  One possible explanation for the inability to make the simple diosphenol could be due to the electron density of the adjacent aromatic ring.  In the natural product diterpenoids (5.34– 5.36) (Figure 5.8), the oxygen substituent para to the ketone provides electron density, making the ketone, and likely the diketone or diosphenol less reactive to attack by a nucleophile. However, even in the natural products, several examples exist for formation of the lactol, cupresol (5.37) (Figure 5.8) being the most notable.214 It is possible that some of the desired 5.28 (Scheme 5.8) may form in the reaction mixture, however, stabilizing substituents are not present, making the diosphenol more electrophilic and possibly more prone to degradation.   Scheme 5.8 Synthetic efforts towards 5.28. 5.3.2 Attempted Synthesis of 5.30 Compound 5.30 shown in Figure 5.7, as with 5.28 was proposed to determine if the quinone present in the xestoquinones and adociaquinones was necessary for IDO inhibition. This O O TMSO OH O OHXIBX or DMPmCPBA,DCM, 0 ° C,K2CO370%TMSOTf, DCMNEt3, 0 ° CO2, KOt-But-BuOHOO OH XIBX, DMSOOO ODioxane,Reflux, 5% OHSeO2, AcOH,80 ° C, 64%5.29 5.45.285.38 5.395.4 5.40  172 compound was to mimic tetrahydroxestoquinol (4.22, Figure 5.7), without the nucleophilic furan moiety. Initial formation of the carbon skeleton 5.18 (Scheme 5.3), as described previously, relied on the Sato methodology.200 Bromination of the benzylic methyl groups using NBS, ABCN in CCl4 at reflux proceeded, though 5.41 (Scheme 5.9) decomposed rapidly. Carrying forward with only partial purification of the product minimized this issue. Displacement of bromine on 5.41 by acetate gave the diacetate 5.42 (Scheme 5.8).  All attempts at this point to oxidize alpha to the ketone failed, as was seen with 5.4.  Several different methodologies were attempted, with only diol 5.43 (Scheme 5.9) being formed.  This compound was used in place of 5.30 (Scheme 5.9) for biological testing.    Scheme 5.9 Attempted synthesis of 5.30 5.3.3 Thiazine Containing Analogues  The final derivatives made were 5.44, 5.45, 5.48 and 5.49 (Scheme 5.10).  The tetracycles 5.44 and 5.45 (Scheme 5.10) were a fortuitous discovery from the reaction of 5.14 (Scheme 5.10) with hypotaurine, to create an intermediate towards synthesis of other analogues.  Intriguingly, these compounds were potent hIDO inhibitors and also represent the first examples of a thiazine ring being formed from the addition of hypotaurine to an α-halo ketone.   A similar route was used to access compounds 5.32 and 5.33 (Scheme 5.10), which were obtained as an inseparable mixture of regioisomers.  The mixture showed very potent activity against IDO (Table 5.1) surpassing even 5.23 and 5.24 (Scheme 5.10) as inhibitors. This result makes this scaffold a prime candidate for the minimal pharmacophore of the adociaquinones. Continued HPLC purification will hopefully lead to pure 5.32 and 5.33 (Scheme 5.10) for further testing. O OBrBrNBS, ABCNCCl4 , reflux KOAc, DMF40 ° C, 35%over 2 steps OOAcOAcO2, KO t- Bu,t- BuOH, 80%OOHOHOOHOH OH x5.30 5.41 5.425.435.18  173   Scheme 5.10 Reaction sequence for proposed minimal pharmacophore analogues 5.4 Biological Activity of Adociaquinone and Xestosaprol O (4.20) Derivatives The graphics below contain preliminary hIDO inhibition data for all of the synthetic derivatives discussed above.  These values will be discussed further in the conclusion section of this chapter.   Figure 5.9 Comparison of the effect of 5.24 (filled circles) and 5.23 (hollow circles) on the enzymatic activity of IDO (100 mM potassium phosphate buffer, pH 6.5, 100 nM IDO, 37 °C).       OO O Br HNS OOO O HN S OOH2N SO 2H5.14 5.44 5.45NHS OOOO HN S OOACN, EtOH,H2O +OO O HNSOO OOO5.47 5.33 NHSOO O O5.32O+H2N SO2HACN, EtOH,H2O  174 # IC50 # IC50 5.5 6.8 µM 5.38 >100 µM 5.21 10.4 µM 5.39 >100 µM 5.22 30 µM 5.40 40 µM 5.23 1.41 µM 5.43 >100 µM 5.24 0.114 µM 5.44 0.20 µM 5.29 >100 µM 5.45 0.059 µM 5.31 3.6 µM 5.48/5.49 0.039 µM     Figure 5.10 IC50 values for synthetic xestoquinone analogues with SAR diagram of xestosaprol O colored to reflect findings. Black bonds/ atoms represent the minimal pharmacophore; green bonds/ atoms are neutral additions or untested; and blue bonds/ atoms appear deleterious for hIDO inhibition. 5.5 Conclusion The above route (Scheme 5.7) to deshydroxy-xestosaprol N (5.23) and O (5.24) represents the first synthesis of this natural product scaffold. The route is the first example of a regio-selective cyclization of hypotaurine onto a hydroxyquinone and the products 5.23 and 5.24 offer further support for the structure assignments of xestosaprol O (4.20) (Figure 5.3).  This route is also the first use of the photocyclization methodology towards a natural product scaffold.199, 200 The Sato methodology200 provided the carbon framework of xestosaprol O (4.20) in one synthetic transformation from easily accessible starting materials, highlighting the efficiency of this route. The biological activity of the synthetic analogues 5.24 (Figure 5.10) and 5.45 (Figure 5.10) are two of the most potent hIDO inhibitors in vitro reported to date, surpassing the inhibitory potency of the natural products from this family, adociaquinone A (4.11) (IC50 = 2 µM) and xestosaprol O (4.19) (IC50 = 4 µM)(Figure 5.1 and 5.3). The bioassay results illuminate the thiazine ring regiochemistry in 5.24 (IC50 = 114 nM) and 5.45 (IC50 = 54 nM) (Figure 5.10) as the most potent for IDO inhibition. In comparison, the corresponding regioisomers 5.23 (IC50 O OOHOHO xestosaprol O ( 4.20 )SO OHN preferredregiochemistry  175 =1.41 µM) and 5.44 (IC50 =130 nM) (Figure 5.11) are an order of magnitude less potent. The inactivity of compounds 5.29, 5.39, and 5.43 (Figure 5.11) support the naphthoquinone moiety as the pharmacophore.  However, the decreased inhibitory activity of 5.31 (IC50 = 3.6 µM)(Figure 5.11) as compared to the natural products, suggest the need for an electron withdrawing substituent on the benzene ring of the naphthoquinone as is seen in 5.32, 5.33, 5.44, and 5.45 (Figure 5.11).  Initial bioassay results of the minimal pharmacophore analogue 5.32/ 5.33 (as a mixture) (Figure 5.11) and the two analogues 5.23 and 5.24 (Figure 5.11) have verified this class of compounds as potent in vitro inhibitors of hIDO.  Future testing of these compounds in both cell and animal models of IDO will reveal if this class has any potential for future use in the clinic.    Figure 5.11 Analogues of the xestoquinones tested. OOOHO OOO OOOHO OHOHOH5.215.55.22 HNS OOOOOSOOOO ONH S OOO ONH 5.315.325.33HNS OOO O HN SOO5.44 5.45NHS OOOO HN SOOOOHOH 5.43O OH5.39O5.29O OH5.40OOO OHSNHO O 5.24OOO OHHNSO O 5.23  176 5.6 Experimental Section 5.6.1 General Experimental Parameters for the IDO Assay Human indoleamine 2, 3-dioxygenase expressed in E. coli was purified as described elsewhere.215 The inhibitory effect of the compounds on the activity of this enzyme was assayed in vitro based on methods developed by Takikawa et al. with some modifications.187 For IC50 value determinations, assays were conducted in 96-well plates (60 µL reaction volume) at 37 °C. Reaction mixtures contained ascorbic acid (10 mM), L-Trp (400 µM), methylene blue (1.25 µM), bovine catalase (tetramer: 40 nM), IDO (100 nM), DMSO (1%) and varying inhibitor concentrations in potassium phosphate buffer (100 mM, pH 6.5).216 The reactions were initiated by simultaneous addition of the ascorbic acid and L-Trp last. The progress of Trp oxidation in reactions lacking inhibitor was monitored spectrophotometrically at 321 nm (Tecan Infinite M200 plate reader). All reactions in a plate were terminated by addition of 30 % TCA and incubation at 65 °C (covered) for 15 minutes as soon as the uninhibited reactions ceased to progress linearly (∼45 min). Once cooled, 2% para-(dimethylamino)benzaldehyde dissolved  in glacial acetic acid was added, and the absorbance of the  product was measured at 480 nm. 5.6.2 General Experimental Procedures All non-aqueous reactions were carried out in oven dried Pyrex® glassware under an Ar atmosphere unless otherwise noted. Air and moisture sensitive reagents were manipulated using airtight dry syringes.  Anhydrous solvents were all obtained from commercial sources and all reagents were obtained from commercial sources without further purification.  All 1H and 13C NMR spectra were recorded at 600 and 150 MHz respectively as indicated and referenced to the internal residual solvent peak denoted in the experimental.  Flash chromatography was performed using silica gel (230–400 mesh) with the solvent system indicated.  All UV reactions were performed in a photo-reactor with a water-cooled Pyrex® filtered 450 W medium pressure mercury lamp.        177 5.6.3 Synthetic Procedures and Methodology Preparation of 5.11:  To a round bottom flask containing dimethyl naphthoquinol (5.8) (0.5 g, 2.65 mmol) in DCE (20 mL) at 0 °C was added chloroacetyl anhydride (0.547 g, 3.2 mmol) and AlCl3 (0.777 g, 5.83 mmol) under N2 atmosphere.  This mixture was heated to 60 °C and stirred together for 2 hours yielding a dark liquid. This was then poured over ice (~80 ml) to give a bright yellow precipitate.  The melted ice water and diluted reaction mixture was extracted with DCM (3 X 20 mL). The combined organic layers were then dried with brine, MgSO4, filtered, and concentrated by rotoevaporation.  The crude reaction mixture was then fractionated using silica gel chromatography (eluting with 20% DCM/toluene) to give purified 2-chloro-1-(5, 8-dimethoxynaphthalen-2-yl)ethanone (5.11) (0.082 g, 11.7%) as a bright yellow solid, and the undesired regioisomer 5.12 (0.203 g, 29%). Characterization of 5.11:  1H NMR (600 MHz, acetone-d6) δ 8.87 (s, 1H), 8.26 (d, J = 9.0 Hz, 1H), 8.05 (dd, J = 9.0, 2.0 Hz, 1H), 7.03 (d, J = 8.7, 6.9 Hz, 1H), 5.15 (s, 2H), 4.00 (s, 3H), 3.98 (s, 3H); 13C NMR (150 MHz, acetone-d6) δ 190.5, 149.7, 148.6, 131.3, 127.7, 124.8, 123.6, 123.3, 122.0, 106.5, 104.3, 54.9, 46.1 (X2); positive ion HRESIMS [M+H]+ m/z 265.0631 (calcd. for C14H14O3Cl, 265.0631). OO O ClOO O ClOOClAlCl3,  DCE, 0 ° C – 60 ° C OO O Cl+5.8 5.11 5.12  178   Figure 5.12 1H and 13C NMR spectra of 5.11 in acetone-d6 at 600 MHz and 150 MHz    179    Figure 5.13 1H and 13C NMR spectra of 5.12 in MeOD-d4 at 600 MHz and 150 MHz   180 Preparation of 5.15:  To a stirred solution of 5.25 (2 g, 11.5 mmol) and MeOH (60 mL) in a round bottom flask was added NH4Br (1.24 g, 12.6 mmol), and Oxone® (7.746 g, 12.6 mmol) at room temperature open to air.  This was heated to reflux, stirred for 3 hours, and monitored by TLC for complete consumption of the starting material.  Once all starting material was consumed, the reaction was filtered to remove excess Oxone®. The residue was washed once with cold MeOH.  The MeOH wash and reaction liquid were combined and dried under reduced pressure to give a crude paste, which was purified directly by silica gel column chromatography (hexanes/toluene/diethyl ether (4:1.5:0.5)) as eluent giving 2-bromo-1-(5, 6, 7, 8-tetrahydronaphthalen-2-yl)ethanone (5.15) (2.61 g, ~89.5%) as an off white solid. 1H NMR (600 MHz, CDCl3) δ 7.66 (s, 1H), 7.65 (d, J = 7.5 Hz, 1H), 7.15 (d, J = 7.5 Hz, 1H), 4.40 (s, 2H), 2.79 (bm, 4H), 1.79 (bm, 4H); 13C NMR (150 MHz, CDCl3) δ 191.4, 144.5, 138.0, 131.5, 129.9, 129.7, 126.1, 31.3, 29.9, 29.5, 23.0, 22.8; positive ion HRESIMS [M+H]+ m/z 253.0236 (calcd. for C12H14OBr, 253.0228).       O O BrNH4BrOxone!  , MeOHreflux, 3 hrs 5.155.25  181  Figure 5.14 1H and 13C NMR spectra of 5.15 in CDCl3 at 600 MHz and 150 MHz   182 Preparation of 5.16:  To a Pyrex® test-tube containing 5.15 (0.25 g, 0.98 mmol) under N2 atmosphere, was added silver triflate (0.253 g, 0.98 mmol), methyl cyclohexene (5.3)(0.23 mL, 1.96 mmol), and benzene (100 mL). N2 was then allowed to bubble through the reaction to remove oxygen.  The test tubes were cooled using an ice bath and irradiated using a medium pressure mercury lamp for 2 hours while stirring.199, 200 After irradiation, 10 mL of a 5% NaHCO3 solution was added and allowed to stir for 30 min, after which time the reaction mixture was filtered over a silica gel frit to remove the silver bromide precipitate and washed using EtOAc.  The filtrate was then concentrated under reduced pressure and purified directly using silica gel column chromatography (hexanes/toluene/diethyl ether (4:1:1) as eluent), yielding 12b-methyl-1, 3, 4, 4a, 5, 8, 9, 10, 11, 12b-decahydrotetraphen-6(2H)-one (5.16) as a clear yellow oil (0.067 g, 26%). 1H NMR (600 MHz, CDCl3) δ 7.74 (s, 1H), δ 6.99 (s, 1H), 3.00 (dd, J = 17.6, 5.0 Hz, 1H), 2.77 (m, 4H), 2.38 (dd, J = 17.6, 3.2 Hz, 1H), 2.32 (bd, J = 14.1 Hz, 1H), 1.95 (m, 1H), 1.59-1.49 (m, 3H), 1.41 (m, 1H), 1.32 (m, 1H), 1.29 (s, 3H), 1.22 (m, 2H); 13C NMR (150 MHz, CDCl3) δ 198.6, 146.2, 144.5, 135.3, 129.6, 128.1, 126.7, 43.2, 42.0, 37.7, 32.0, 30.2, 29.5, 29.0, 26.1, 23.2, 23.1, 22.5; positive ion HRESIMS [M+H] + m/z 269.1905 (calcd for C19H25O, 269.1905).      O Br O5.15 5.165.3 , AgOTfbenzene, h!  183   Figure 5.15 1H and 13C NMR spectra of 5.16 in CDCl3 at 600 MHz and 150 MHz   184 Preparation of 5.17A:  To a round bottom flask containing 3’, 4’-dimethyl acetophenone (1.0 g, 6.75 mmol) and MeOH (50 mL) was added NH4Cl (0.722 g, 13.5 mmol) and Oxone® (4.56 g, 7.42 mmol) sequentially.  This was stirred vigorously at room temperature overnight and monitored by TLC for consumption of starting material.  When all of the starting material had been consumed (~24 hours), the reaction was filtered to remove undissolved solids and washed with cold MeOH.  The wash and filtrate were combined and concentrated to give a thick paste, which was purified using silica gel chromatography (eluting with (8:1.5:0.5) hexanes/toluene/diethyl ether) to give 2-chloro-1-(3, 4-dimethylphenyl)ethanone (5.17C) as an off white solid (0.957 g, 78%). 1H NMR (600 MHz, CDCl3) δ 7.71 (s, 1H), δ 7.66 (d, J = 7.4 Hz, 1H), δ 7.22 (d, J = 7.4 Hz, 1H), δ 4.67 (s, 2H), δ 2.31 (s, 3H), δ 2.30 (s, H); 13C NMR (150 MHz, CDCl3) δ 191.1, δ 144.0, δ 137.6, δ 132.3, δ 130. 3, δ 129.8, δ 126.4, δ 46.3, δ 20.4, δ 20.0; positive ion HRESIMS [M+H]+ m/z 183.0578 (calcd. for C10H12OCl, 183.0577).       OO ClNH 4 Cl, Oxone !MeOH 5.17A  185   Figure 5.16 1H and 13C NMR spectra of 5.17A in CDCl3 at 600 MHz and 150 MHz   186 Preparation of 5.17B:   To a round bottom flask containing 3’, 4’-dimethyl acetophenone (1.0 g, 6.75 mmol) and MeOH (50 mL) was added NH4Br (0.730 g, 7.4 mmol) and Oxone® (4.56 g, 7.42 mmol) sequentially.  This was stirred vigorously at room temperature overnight and monitored by TLC for consumption of starting material.  When all of the starting material had been consumed (~24 hours), the reaction was filtered to remove undissolved solids and washed with cold MeOH.  The wash and filtrate were combined and concentrated to give a thick paste, which was purified using silica gel chromatography (eluting with (8:1.5:0.5) hexanes/toluene/diethyl ether) to give 2-bromo-1-(3, 4-dimethylphenyl)ethanone (5.17B) (1.40 g, 92%) as a white solid. 1H NMR (600 MHz, acetone-d6) δ 7.82 (s, 1H), 7.78 (d, J = 7.9 Hz, 1H), 7.30 (d, J = 7.9 Hz, 1H), 4.71 (s, 2H), 2.33 (s, 6H); 13C NMR (150 MHz, acetone-d6) δ 190.3,143.0, 136.8, 131.8, 129.4, 129.3, 126.1, 31.6, 18.7, 18.4; positive ion HRESIMS [M+Na]+ m/z 248.9895 (calcd. for C10H11ONaBr, 248.9891).       OO BrNH4Br, Oxone!MeOH 5.17B  187   Figure 5.17 1H and 13C NMR spectra of 5.17B in acetone-d6 at 600 MHz and 150 MHz   188 Preparation of 5.17C:    5.17B  (0.1 g, 0.44 mmol) and acetone (10 mL) were stirred together with potassium iodide (0.73 g, 4.4 mmol) in a round bottom flask overnight.  After 18 hours, the remaining undissolved KI was removed using suction filtration and the reaction mixture was concentrated under a stream on N2.  The product was diluted in DCM and passed through a silica gel frit to give 2-iodo-1-(3, 4-dimethylphenyl)ethanone (5.17C) (0.108 g, 90% yield) as a brown amorphous solid without the need of further purification. 1H NMR (600 MHz, CDCl3) δ 7.73 (s, 1H), 7.68 (d, J = 8.1 Hz, 1H), 7.19 (d, J = 8.1 Hz, 1H), 4.30 (s, 2H), 2.30 (s, 3H), 2.29 (s, 3H); 13C NMR (150 MHz, CDCl3) δ 192.9, 143.8, 137.4, 131.4, 130.2, 130.1, 126.9, 20.3, 19.9, 2.2; positive ion HRESIMS [M+Na]+ m/z 296.9741 (calcd. for C10H10ONaI, 296.9752).          O Br O IKI, Acetone 5.17C5.17B  189   Figure 5.18 1H and 13C NMR spectra of 5.17C in CDCl3 at 600 MHz and 150 MHz   190 Preparation of 5.18:   To a Pyrex® test-tube containing 5.17B (0.1 g, 0.44 mmol) in benzene (45 mL) was added silver triflate (0.113 g, 0.44 mmol) and methyl cyclohexene (5.3)(0.078 mL, 0.66 mmol).199, 200 N2 was then allowed to bubble through the reaction to remove oxygen.  The test tubes were cooled using an ice bath and irradiated using a medium pressure mercury lamp for 2 hours while stirring.199, 200 After irradiation, 10 mL of a 5% NaHCO3 solution was added and allowed to stir for 30 min, after which time the reaction mixture was filtered over a silica gel frit to remove the silver bromide precipitate and washed using EtOAc.  The filtrate was then concentrated under reduced pressure and purified directly using silica gel column chromatography (hexanes/toluene/diethyl ether (4:1:1) as eluent) to give purified 4a, 6, 7-trimethyl-2, 3, 4, 4a, 10, 10a-hexahydrophenanthren-9(1H)-one (5.18) as a waxy yellow solid (0.035 g, 33%). 1H NMR (600 MHz, CD2Cl2) δ 7.74 (s, 1H), 7.13 (s, 1H), 2.99 (dd, J = 18.3, 4.4 Hz, 1H), 2.36–2.32 (m, 2H), 2.31 (s, 3H), 2.26 (s, 3H), 1.97 (m, 1H), 1.59 (m, 1H), 1.54 (m, 2H), 1.44 (m, 1H), 1.35 (m, 1H), 1.31 (s, 3H), 1.22–1.15 (m, 2H); 13C NMR (150 MHz, CD2Cl2) δ 198.2, 147.3, 144.3, 135.1, 130.3, 128.3, 127.8, 43.5, 42.3, 38.0, 32.0, 30.3, 30.0, 26.5, 23.0, 20.8, 19.5; positive ion HRESIMS [M+H]+ m/z 243.1741 (calcd. for C17H23O, 243.1749).    O Br AgOTf, h !benzene O5.17B 5.18  191   Figure 5.19 1H and 13C NMR spectra of 5.18 in CD2Cl2 at 600 MHz and 150 MHz    192 Preparation of 5.20 and 5.19:  5.16 (0.05 g, 0.18 mmol) was added to a round bottom flask and dissolved in 4 mL ACN and cooled to 0 °C.  KMnO4 (0.177 g, 1.12 mmol) was then added to the reaction mixture in three portions over an hour and allowed to warm to room temperature.  This mixture was allowed to stir open to air for 24 hours, at which time it was filtered. Excess KMnO4 in the filtrate was removed using a 10% metapersulphate solution (~0.2 mL) and further filtered to remove MnO2 precipitate. The filtrate was then concentrated and separated using silica gel chromatography (EtOAc/hexanes (1:5 and 1:4) as eluent), yielding both 12b-methyl-1, 3, 4, 4a, 5, 9, 10, 12b-octahydrotetraphene-6, 11(2H, 8H)-dione (5.20) (0.011 g, 21%) and 12b-methyl-1, 3, 4, 4a, 5, 10, 11, 12b-octahydrotetraphene-6, 8(2H, 9H)-dione (5.19) (0.009 g, 19%) as off white solids.  Characterization of 5.20: 1H NMR (600 MHz, CDCl3) δ 8.02 (s, 1H), 7.91 (s, 1H), 3.06 (dd, J = 17.3, 4.8 Hz, 1H), 2.97 (m, 2H), 2.67 (t, J = 6.6 Hz, 2H), 2.45 (m, 2H), 2.13 (m, 2H), 2.00 (m, 1H), 1.57 (m, 2H), 1.53 (m, 1H), 1.46 (td, J = 12.8, 3.8 Hz, 1H), 1.37 (m, 1H), 1.32 (s, 3H), 1.14 (m, 2H); 13C NMR (150 MHz, CDCl3) δ 198.5, 198.5, 147.3, 141.9, 136.5, 135.05, 128.1, 125.6, 42.9, 42.1, 39.5, 38.5, 37.7, 31.9, 29.6, 29.4, 26.1, 23.2, 22.5; positive ion HRESIMS [M+Na]+ m/z 305.1517 (calcd. for C19H22O2Na, 305.1517).  O OKMnO 4ACN, 0 ° C– rt OOO +5.16 5.195.20  193    Figure 5.20 1H and 13C NMR spectra of 5.20 in CDCl3 at 600 MHz and 150 MHz    194 Characterization of 5.19:  12b-methyl-1, 3, 4, 4a, 5, 10, 11, 12b-octahydrotetraphene-6, 8(2H, 9H)-dione (5.19): 1H NMR (600 MHz, CDCl3) δ 8.68 (s, 1H), 7.21 (s, 1H), 3.03 (dd, J = 16.8, 5.6 Hz, 1H), 2.99 (q, J = 5.8 Hz, 2H), 2.64 (t, J = 6.4 Hz, 2H), 2.44 (dd, J = 17.7, 3.4 Hz, 1H), 2.37 (bd, J = 14.7 Hz, 1H), 2.14  (m, 2H), 1.99 (m, 1H), 1.61–1.55 (m, 3H), 1.48 (td, J = 13.5, 3.3 Hz, 1H), 1.36 (obs, 1H), 1.33 (s, 3H), 1.16 (m, 2H); 13C NMR (150 MHz, CDCl3) δ 197.4, 197.2, 154.3, 150.0, 131.2, 131.0, 127.5, 126.7, 42.9, 41.8, 39.2, 39.0, 37.8, 31.8, 30.5, 29.7, 26.1, 23.1, 22.6; positive ion HRESIMS [M+H]+ m/z 283.1698 (calcd. for C19H23O2, 283.1698).            OO 5.19  195   Figure 5.21 1H and 13C NMR spectra of 5.19 in CDCl3 at 600 MHz and 150 MHz    196 Preparation of 5.22:  O2 gas was bubbled through a vial containing ketone 5.20 (0.02 g, 0.07 mmol), in tBuOH (2 mL) at 25 °C.  After 5 minutes, KOtBu (0.039 g, 0.354 mmol) was added into the reaction vessel in several small aliquots over 5 minutes, turning the reaction mixture green, then dark red. After 4 hours, the O2 was removed and the reaction quenched by addition of approximately 1.5 mL of 1M HCl. The crude reaction mixture was concentrated under a stream of nitrogen and fractionated using C18 reversed phase column chromatography (eluting with 100% H2O, (4:1) H2O/MeOH and (1:1) H2O/MeOH), yielding partially purified product ((1:1) H2O/MeOH fraction).  This was carried forward in its partially purified state for the final reaction. A second reaction was purified using reversed phase C18 HPLC (eluting with 40% ACN/H2O (0.05% TFA) retention time: 41 min), to give 5, 10-dihydroxy-12b-methyl-1, 3, 4, 12b-tetrahydrotetraphene-6, 8, 11(2H)-trione (5.22) as a yellow solid (0.009 g, 39%). 1H NMR (600 MHz, DMSO-d6) δ 11.86 (bs, 1H), 8.81 (s, H), 8.58 (s, 1H), 8.32 (s, 1H), 6.24 (s, 1H), 3.16 (obs, 1H), 2.54 (obs, 1H), 2.18 (td, J = 13.6, 5.6 Hz, 1H), 1.97 (bd, J = 13.4 Hz, 1H), 1.89 (m, 1H), 1.72 (bd, J = 13.4 Hz, 1H), 1.53 (s, 3H), 1.32 (td, J = 13.3, 4.0 Hz, 1H), 1.22 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 183.7, 180.8, 177.6, 160.2, 155.9, 142.3, 139.6, 132.6, 132.5, 130.2, 124.5, 123.5, 111.6, 40.5, 39.6, 26.1, 26.0, 23.1, 21.4; positive ion HRESIMS [M+H] + m/z 323.0915 (calcd. for C19H15O5, 323.0919).  OO OOOHOKO t- Bu , O 2t- BuOH OH5.20 5.22  197   Figure 5.22 1H and 13C NMR spectra of 5.22 in DMSO-d6 at 600 MHz and 150 MHz   198 Characterization of 5.26:  4a-hydroxy-12b-methyl-2, 3, 4, 4a, 9, 10-hexahydro-1H-naphtho[2, 3-c]chromene-6, 11(8H, 12bH)-dione. Isolated as a product from reaction conditions for 5.22. C18 reversed phase HPLC (retention time of 57 min eluting with (2:5) ACN/H2O) gave (5.2 mg). 1H NMR (600 MHz, CDCl3) δ 8.06 (s, 1H), 8.03 (s, 1H), 3.36 (bs, 1H), 2.99 (t, J = 6.0 Hz, 2H), 2.68 (t, J = 6.0 Hz, 2H), 2.15 (m, 2H), 1.98 (m, 1H), 1.89 (m, 1H), 1.74 (m, 2H), 1.68 (bm, 3H), 1.58 (m, 1H), 1.52 (bs, 3H); 13C NMR (150 MHz, CDCl3) δ 198.1, 164.5, 142.9, 136.7, 131.1, 127.4, 124.1, 122.3, 105.8, 41.6, 39.3, 34.8, 29.3, 23.1, 22.8, 22.3, 20.8, 20.5; positive ion HRESIMS [M+Na]+ m/z 323.1257 (calcd. for C18H20O4Na, 323.1259).            OO OHO 5.26  199   Figure 5.23 1H and 13C NMR spectra of 5.26 in CDCl3 at 600 MHz and 150 MHz   200 Characterization of 5.27:  Isolated from reaction conditions for 5.22 using C18 RPHPLC (retention time = 16 minutes eluting with (2:5) ACN/H2O) provided 4a, 10-dihydroxy-12b-methyl-2, 3, 4, 4a-tetrahydro-1H-naphtho[2, 3-c]chromene-6, 8, 11(12bH)-trione (5.27) as a yellow solid (6.3 mg).  It should be noted that this compound degrades quickly upon purification. 1H NMR (600 MHz, DMSO-d6) δ 11.90 (bs, 1H), 8.42 (s, 1H), 8.07 (s, 1H), 7.54 (s, 1H), 6.22 (bs, 1H), 1.93 (bm, 1H), 1.87 (bm, 1H), 1.69 (bm, 2H), 1.56 (m, 1H), 1.54 (bm, 2H), 1.51 (bs, 3H), 1.45 (bm, 1H); 13C NMR (150 MHz, DMSO-d6) δ 183.3, 180.5, 163.0, 154.7, 154.3, 152.3, 138.7, 134.2, 130.1, 128.3, 111.2, 105.4, 41.9, 39.4, 32.8, 22.3, 20.1, 19.8; negative ion HRESIMS [M-H]- m/z 327.0881 (calcd. for C18H15O6, 327.0869).        5.27 OOOOHO OH  201  Figure 5.24 1H and 13C NMR spectra of 5.27 in DMSO-d6 at 600 MHz and 150 MHz   202   Figure 5.25 gHMBC and gHSQC NMR spectra of 5.27 in DMSO-d6 at 600 MHz   203 Preparation of 5.21:  O2 gas was bubbled through a vial containing 5.19 (0.02 g, 0.07 mmol) in 2 mL glyme at -10 °C.  After 5 minutes, KOtBu (0.039 g, 0.35 mmol) was added into the reaction in several small aliquots over 5 minutes, turning the reaction mixture dark red. After 4 hours, the O2 was removed and the reaction quenched by addition of approximately 1.5 mL of 1M HCl. The crude reaction mixture was concentrated under a stream of nitrogen and fractionated using C18 reversed phase column chromatography (Sep-Pak) (eluting with H2O, (4:1) H2O/MeOH and (1:1) H2O/MeOH), yielding partially purified product ((1:1) H2O/MeOH fraction). This was then purified using C18 reversed phase HPLC (eluting with 40% ACN/H2O with 0.05% TFA (retention time: 46 min)) giving 5, 9-dihydroxy-12b-methyl-1, 3, 4, 12b-tetrahydrotetraphene-6, 8, 11(2H)-trione (5.21) as a light yellow solid (0.009 g, 41%). 1H NMR (600 MHz, DMSO-d6) δ 11.94 (bs, 1H), 8.80 (s, 1H), 8.61 (s, 1H), 8.24 (s, 1H), 6.25 (s, 1H), 3.15 (bd, J = 14.4 Hz, 1H), 2.49 (obs, 1H), 2.17 (td, J = 13.8, 4.6 Hz, 1H), 1.95 (bd, J = 13.8 Hz, 1H), 1.88 (m, 1H), 1.70 (bd, J = 14.4 Hz, 1H), 1.51 (s, 3H), 1.31 (td, J = 13.8, 4.6 Hz, 1H), 1.21 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 184.1, 180.3, 177.6, 160.1, 157.1, 142.2, 139.5, 133.8, 131.9, 129.0, 124.2, 123.8, 111.8, 40.6, 39.6, 26.1, 26.0, 23.1, 21.3; negative ion HRESIMS [M-H]- m/z 307.0973 (calcd. for C19H15O4, 307.0970).  OO OOOHO OHO2, t -BuOK-10 ° C, 4hrsglyme 5.215.19  204    Figure 5.26 1H and 13C NMR spectra of 5.21 in DMSO-d6 at 600 MHz and 150 MHz    205 Preparation of 5.21 and 5.5:  O2 gas was bubbled through a vial containing 5.19 (0.02 g, 0.07 mmol) in 2 mL tBuOH at room temperature.  After 5 minutes, KOtBu (0.039 g, 0.35 mmol) was added into the reaction in several small aliquots over 5 minutes turning the reaction mixture green, then dark red. After 4 hours, the O2 was removed and the reaction quenched by addition of approximately 1.5 mL of 1M HCl. The crude reaction mixture was concentrated under a stream of nitrogen and fractionated using C18 reversed phase column chromatography (Sep-Pak)(eluting with H2O, (4:1) H2O/MeOH and (1:1) H2O/MeOH), yielding partially purified product ((1:1) H2O/MeOH fraction). The products were then purified using C18 reversed phase HPLC (eluting with (2:5) ACN/H2O with 0.05% TFA) giving two products 5.21 (0.004 g, 21%) and 5-hydroxy-12b-methyl-1, 3, 4, 12b-tetrahydrotetraphene-6, 8, 11(2H)-trione (5.5) (0.005 g, 22%) as a yellow solid (HPLC retention time: 48 min for 5.5). 1H NMR (600 MHz, DMSO-d6) δ 11.94 (bs, 1H), 8.56 (s, 1H), 8.24 (s, 1H), 6.39 (s, 1H), 6.25 (s, 1H), 2.69 (td, J = 13.0, 6.5 Hz, 1H), 2.52 (obs, 2H), 2.01 (bd, J = 13.0 Hz, 1H), 1.91 (m, 1H), 1.72 (bd, J = 13.0 Hz, 1H), 1.54 (s 3H), 1.39 (td, J = 13.0, 4.4 Hz, 1H), 1.34 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 183.9, 182.3, 180.4, 170.2, 160.2, 157.2, 134.1, 132.8, 129.1, 123.8, 123.2, 124.1, 111.9, 41.6, 40.1, 32.7, 27.4, 25.7, 21.3; negative ion HRESIMS [M-H]- m/z 323.0917 (calcd. for C19H15O5, 323.0919). OO OOOHO OH OOO OH+KO t -Bu, O2t -BuOH5.19 5.21 5.5  206    Figure 5.27 1H and 13C NMR spectra of 5.5 in DMSO-d6 at 600 MHz and 150 MHz    207 Preparation of 5.23:    To a round bottom flask was added 5.21 (0.005 g, 0.015 mmol) and hypotaurine (0.003 g, 0.023 mmol) dissolved in 0.5 mL of H2O, with a drop of TFA. To this 2 mL of ACN and EtOH (1:1) were added and the reaction mixture was stirred while heated to 80 °C under O2 atmosphere.  After 36 hours, the reaction mixture showed complete consumption of starting material, and was concentrated under a stream of N2 and then purified directly on reversed phase C18 HPLC (eluting with (2:5) ACN/H2O with 0.05% TFA (retention time: 22 min)), yielding 5-hydroxy-14b-methyl-3, 4, 11, 12-tetrahydro-1H-tetrapheno [9, 10-b] [1, 4]thiazine-6, 8, 13(2H, 10H, 14bH)-trione 9, 9-dioxide (5.23) as a bright yellow solid (0.003 g, 42%).  1H NMR (600 MHz, DMSO-d6) δ 9.23 (bs, 1H), 8.83 (bs, 1H), 8.63 (s, 1H), 8.32 (s, 1H), 3.87 (m, 2H), 3.39 (t, J = 6.0 Hz, 2H), 3.15 (bd, J = 9.4 Hz, 1H), 2.5 (obs, 1H), 2.17 (td, J = 13.7, 4.9 Hz, 1H), 1.95 (bd, J = 12.7 Hz, 1H), 1.87 (q, J = 12.7 Hz, 1H), 1.71 (bd, J = 12.7 Hz, 1H), 1.52 (s, 3H), 1.31 (td, J = 13.7, 3.9 Hz, 1H), 1.21 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 178.0, 177.5, 173.7, 158.1, 147.3, 142.3, 139.5, 134.4, 131.6, 128.4, 124.7, 124.2, 111.9, 48.2, 40.7, 40.0, 39.3 (obs), 26.2, 26.1, 23.1, 21.4; negative ion HRESIMS [M-H]- m/z 412.0859 (calcd. for C21H18NO6S, 412.0855).     OOOHO OH H2N SO 2HACN/ EtOH/ H 2OTFA, O 2 OOO OHHNSO O5.21 5.23  208    Figure 5.28 1H and 13C NMR spectra of 5.23 in DMSO-d6 at 600 MHz and 150 MHz    209 Preparation of 5.24:  In a small conical flask, 5.22 (0.007 g, 0.022 mmol), and hypotaurine (0.004 g, 0.032 mmol) were dissolved in 0.5 mL of H2O and 2.5 mL of ACN/EtOH (1:1) with one drop of TFA. The reaction mixture was stirred while heating to 80 °C under O2 atmosphere.  After 36 hours the reaction mixture showed complete consumption of starting material, and was concentrated under a stream of N2 and then purified directly on reversed phase C18 HPLC eluting with (2:5) ACN/H2O with 0.05% TFA (retention time: 24 min) yielding 5-hydroxy-14b-methyl-3, 4, 10, 11-tetrahydro-1H-tetrapheno [10, 9-b][1, 4] thiazine-6, 8, 13(2H, 9H, 14bH)-trione 12, 12-dioxide (5.24) as a bright yellow solid (0.004 g, 45%). 1H NMR (600 MHz, DMSO-d6) δ 9.32 (bs, 1H), 8.81 (bs, 1H), 8.61 (s, 1H), 8.29 (s, 1H), 3.87 (bm, 2H), 3.41 (obs, 2H), 3.15 (d, J = 14.0 Hz, 1H), 2.17 (td, J = 13.9, 5.6 Hz, 1H), 1.96 (bd, J = 14.6 Hz, 1H), 1.89 (bd, J = 14.0 Hz, 1H), 1.73 (bd, J = 13.2 Hz, 1H), 1.52 (s, 3H), 1.32 (td, J = 13.9, 4.2 Hz, 1H), 1.24–1.19 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 177.9, 177.5, 173.8, 158.1, 147.3, 142.3, 139.6, 134.5, 131.7, 128.5, 124.7, 124.3, 111.9, 48.2, 40.7, 40.0, 26.2, 26.1, 23.1, 21.4; positive ion HRESIMS [M+H]+ m/z 436.0828 (calcd. for C21H19NO6SNa, 436.0831).    OOO OHHO H2N SO 2HACN/ EtOH/ H 2OTFA, O 2 OOO OHSNHO O5.22 5.24  210   Figure 5.29 1H and 13C NMR spectra of 5.24 in DMSO-d6 at 600 MHz and 150 MHz   211 Preparation of 5.4:  To a Pyrex® test-tube containing 2-bromoacetophenone  (0.1 g, 0.5 mmol), benzene (45 mL), silver triflate (0.129 g, 0.50 mmol), and methyl cyclohexene (5.3) (0.09 mL, 0.75 mmol).199, 200 N2 was then allowed to bubble through the reaction to remove oxygen.  The test tubes were cooled using an ice bath and irradiated using a medium pressure mercury lamp for 2 hours while stirring.199, 200 After irradiation, 10 mL of a 5% NaHCO3 solution was added and allowed to stir for 30 minutes, after which time the reaction mixture was filtered over a silica gel frit to remove the silver bromide precipitate and washed using EtOAc.  The filtrate was then concentrated under reduced pressure, and purified directly using silica gel column chromatography (hexanes/toluene/diethyl ether (4:1:1) as eluent) to give purified 4a-methyl-2, 3, 4, 4a, 10, 10a-hexahydrophenanthren-9(1H)-one (5.4) as a yellow clear oil (0.047 g, 44%). 1H NMR (600 MHz, CDCl3) δ 7.98 (d, J = 8.0 Hz, 1H), 7.56 (t, J = 7.5 Hz, 1H), 7.39 (d, J = 8.0 Hz, 1H), 7.29 (t, J = 7.5 Hz, 1H), 3.04 (dd, J = 17.4, 4.7 Hz, 1H), 1.80 (dd, J = 17.4, 3.2 Hz, 1H), 2.38 (bd, obs, 1H), 2.02 (m, 1H), 1.61–1.52 (m, 3H), 1.48 (m, 1H), 1.37 (m, 1H), 1.34 (s, 3H), 1.23–1.15 (m, 2H); 13C NMR (150 MHz, CDCl3) δ 198.3, 149.8, 134.5, 132.4, 127.6, 126.8, 126.5, 43.3, 42.2, 38.8, 38.0, 31.9, 29.9, 26.4, 22.9; positive ion LRESIMS [M+H]+ m/z 215.1 (calcd. for C15H19O, 215.1).199, 200         O Br AgOTfbenzene, h! O H+ 5.3 5.4  212 Preparation of 5.29:  To a solution of 5.4 (0.05 g, 0.23 mmol) in DMSO (2 mL), was added IBX (0.196 g, 0.7 mmol) and the mixture was stirred and heated to 80 °C for 24 hours.  The reaction mixture was then diluted in 30 mL of H2O and extracted using EtOAc (3 X 10 mL), the organic layers were combined, washed with brine, dried over MgSO4, filtered, and then concentrated under reduced pressure to give a yellow waxy solid.  This was subjected to silica gel chromatography (eluting with EtOAc/hexanes (1:9)), to give 4a-methyl-2, 3, 4, 4a-tetrahydrophenanthren-9(1H)-one (5.29) as a light yellow oil (0.031 g, 64%). 1H NMR: (600 MHz, CD2Cl2); δ 8.12 (d, J = 8.0 Hz, 1H ), 7.58 (m, 1H), 7.56 (t, J = 7.4 Hz, 1H), 7.39 (t, J = 7.4 Hz, 1H), 6.25 (d, J = 1.3 Hz, 1H), 2.63 (td, J = 13.5, 5.8 Hz, 1H), 2.45 (dpent, J = 13.5, 2.0 Hz, 1H), 2.38 (dpent, J = 13.5, 2.0 Hz, 1H), 2.04 (dm, J = 14.0 Hz, 1H), 1.89 (qt, J = 14.0, 4.4 Hz, 1H), 1.76 (dm, J = 14.0 Hz, 1H), 1.50 (s, 3H), 1.36–1.48 (m, 3H); 13C NMR (150 MHz, CD2Cl2) δ 184.8, 168.5, 152.5, 136.4, 132.6, 126.8, 126.3, 126.2, 124.2, 41.32, 40.9, 33.7, 28.4, 26.5, 22.3; positive ion HRESIMS [M+H]+ m/z 213.1283 (calcd. for C15H17O, 213.1279).      OIBX, DMSOO 80 ° C 5.295.4  213   Figure 5.30 1H and 13C NMR spectra of 5.29 in CD2Cl2 at 600 MHz and 150 MHz   214 Preparation of 5.40:  To a round bottom flask was added 5.4 (0.125 g, 0.58 mmol), dioxane (4 mL), H2O (0.1 mL), and 1 drop of AcOH.  To this solution selenium dioxide (0.083 g, 0.76 mmol) was added.  The solution was heated to reflux for 16 hours and then cooled to room temperature.  The crude was filtered through a silica gel frit and washed with 10 mL of EtOAc.  The wash and filtrate were then combined and concentrated under reduced pressure to give a crude red solid.  This was purified by silica gel chromatography (eluting with EtOAc/hexanes (1:4)), to give 1-hydroxy-4a-methyl-2, 3, 4, 4a-tetrahydrophenanthren-9(1H)-one (5.40) as a red oil (0.006 g, 5%). 1H NMR (600 MHz, acetone-d6) δ 8.10 (d, J = 7.8 Hz, 1H), 7.72 (d, J = 7.8 Hz, 1H), 7.63 (t, J = 7.8 Hz, 1H), 7.42 (t, J = 7.8 Hz, 1H), 6.67 (s, 1H), 4.63 (m, 1H), 2.82 (bs, OH), 2.43 (bd, J = 10.7 Hz, 1H), 2.28 (bs, 1H), 1.99 (m, 1H), 1.79 (m, 1H), 1.51 (s, 3H), 1.33–1.40 (m, 2H); 13C NMR (150 MHz, acetone-d6) δ 183.3, 169.1, 151.5, 131.9, 129.9, 126.0, 125.9, 125.2, 119.0, 67.3, 40.9, 39.6, 36.8, 25.7, 19.5 (Some peaks observed only in gHMBC spectrum); positive ion HRESIMS [M+H]+ m/z 229.1225 (calcd. for C15H17O2, 229.1229).      O ODioxane,Reflux, 5% OHSeO2, AcOH,5.4 5.40  215   Figure 5.31 1H and 13C NMR spectra of 5.40 in acetone-d6 at 600 MHz and 150 MHz   216  Figure 5.32 gHMBC NMR spectrum of 5.40 in acetone-d6 at 600 MHz Preparation of 5.38:  In a small vial, 5.4 (0.05 g, 0.23 mmol) in glyme (3 mL) was stirred at 0 °C for 5 minutes while bubbling O2 through the solution.  After 5 minutes, KOtBu (128.8 g, 1.15 mmol) was added in three separate aliquots over 5 minutes.  After addition of base, O2 was again bubbled through the solution for 2 hours at 0 °C. Addition of 1 mL of 10% HCl solution halted the reaction and silica gel column chromatography (eluting with 15% EtOAc/hexanes) provided 0.02 g (37%) of 4a-hydroxy-10b-methyl-2, 3, 4, 4a-tetrahydro-1H-benzo[c]chromen-6(10bH)-one (5.38). 1H NMR (600 MHz, CD2Cl2) δ 8.07 (d, J = 7.3 Hz, 1H), 7.62 (td, J = 7.3, 1.2 Hz, 1H), 7.45 (d, J = 7.3, 1H), 7.38 (td, J = 7.3, 1.2 Hz, 1H), 3.53 (bs, 1H), 1.98 (m, 1H), 1.91 (bm, 1H), 1.77–1.74 (m, 2H), 1.71–1.67 (m, 2H), 1.58 (m, 1H), 1.51 (obs, 1H), 1.49 (s, 3H); 13C NMR (150 MHz, CD2Cl2) δ 165.1, 134.9, 134.6, 130.4, 127.4, 125.5, 123.9, 105.7, 42.1, 35.1, 20.9, 21.2, 23.3 (2 X C); positive ion HRESIMS [M+H]+ m/z 213.1283 (calcd. for C15H17O, 213.1279). O2 , KO t- Buglyme OO OHO5.4 5.38  217   Figure 5.33 1H and 13C NMR spectra of 5.38 in CD2Cl2 at 600 MHz and 150 MHz   218 Preparation of 5.39:  To a round bottom flask a solution of 5.4 (0.02 g, 0.09 mmol) in DCM (2 mL) under N2 atmosphere at 0 °C was stirred. Triethyl amine (0.02 mL, 0.17 mmol), and then trimethylsilyl triflate (0.02 mL, 0.11 mmol) were sequentially added dropwise over 5 min.  Once the reaction was complete (~1 hour) the reaction mixture was diluted with 2 mL DCM and quenched using 0.5 mL of a saturated NaHCO3 solution, then concentrated under a stream of N2. This was then triturated using dry diethyl ether to remove the product from the insoluble triflate.  The combined ether layers were then concentrated in vacuo and resuspended in DCM (1.5 mL).  To this solution was added NaHCO3 (0.007 g, 0.09 mmol) and mCPBA (0.014 g, 0.081 mmol) at 0 °C. The solution was stirred for 4 hours at which time all starting material had been consumed.  1M HCl (~1 mL) was added and the reaction concentrated. H2O (30 mL) was added and the reaction mixture was extracted using EtOAc (3 X 10 mL).  The organic layers were combined, washed with brine, dried over MgSO4, filtered, concentrated, and purified using silica gel chromatography (15 % EtOAc/hexanes), to give 10-hydroxy-4a-methyl-2, 3, 4, 4a, 10, 10a-hexahydrophenanthren-9(1H)-one  (5.39) as a white solid (0.014 g, 70%). 1H NMR (600 MHz, CDCl3) δ 7.95 (dd, J = 7.8, 1.6 Hz, 1H), 7.57 (td, J = 7.8, 2.0 Hz, 1H), 7.41 (d, J = 7.8 Hz, 1H), 7.31 (td, J = 7.8, 2.0 Hz, 1H), 4.58 (d, J = 13.7 Hz, 1H), 3.78 (bs, 1H), 2.16 (d, J = 14.8 Hz, 1H), 1.96 (d, J = 13.8 Hz, 1H), 1.81 (tt, J = 14.8, 3.7 Hz, 1H), 1.74 (dd, J = 14.8, 4.6 Hz, 1H), 1.67–1.62 (m, 3H), 1.56 (m, 2H), 1.48 (s, 3H); 13C NMR (150 MHz, CDCl3) δ 200.9, 153.6, 134.9, 127.2, 127.0, 126.7, 70.9, 46.1, 38.1, 36.7, 26.2, 22.4, 22.0, 19.9; positive ion HRESIMS [M+Na]+ m/z 253.1201 (calcd. for C15H18O2Na, 253.1204). O TMS O OHm C PBA, K 2 CO 3DCM, 0 ° C ,TMSOTfDCM, NEt 3 , 0 ° CO5.4 5.39  219    Figure 5.34 1H and 13C NMR spectra of 5.39 in CDCl3 at 600 MHz and 150 MHz    220 Preparation of 5.42:  To a round bottom flask was added a mixture of 5.18 (0.02 g, 0.08 mmol) and NBS (0.044 g, 0.248 mmol) in CCl4 (3.2 mL) under Ar atmosphere. Then a single flake of 1,1'-azobis(cyclohexanecarbonitrile) was added and stirred while heated to reflux for 1 hour.  The reaction was cooled to room temperature, concentrated under reduced pressure and fractionated using silica gel chromatography (eluting with EtOAc/hexanes (1:8)) to give partially purified 5.41.  The product was then concentrated in vacuo and diluted with DMF (5 mL) under Ar atmosphere. KOAc (0.017 g, 0.17 mmol) was added in to the solution and this was stirred at 50 °C for 24 hours.  The reaction was cooled, H2O (50 mL) was added, and the mixture was extracted using EtOAc (3 X 20 mL).  The organic layers were combined, washed with brine, dried over MgSO4, filtered, concentrated under reduced pressure, and purified using silica gel chromatography (eluting with EtOAc/Hex (1:5)) to give (4b-methyl-10-oxo-4b, 5, 6, 7, 8, 8a, 9, 10-octahydrophenanthrene-2, 3-diyl)bis(methylene) diacetate (5.42) as a white solid (0.010 g, 35%). 1H NMR (600 MHz, acetone-d6) δ 7.98 (s, 1H), 7.56 (s, 1H), 5.25 (d, J = 4.1 Hz, 2H), 5.21 (s, 2H), 3.08 (dd, J = 17.8, 5.4 Hz, 1H), 2.47 (bd, J = 13.8 Hz, 1H), 2.37 (dd, J = 18.3, 3.5 Hz, 1H), 2.07 (s, 3H), 2.06 (s, 3H), 2.05 (m, 1H), 1.60 (bm, 2H), 1.53 (bm, 2H), 1.39 (obs, 1H), 1.38 (s, 3H), 1.15 (m, 2H); 13C NMR (150 MHz, acetone-d6) δ 196.9, 170.8, 170.7, 150.1, 141.8, 133.9, 132.6, 128.8, 128.5, 64.2, 63.8, 43.5, 42.2, 39.1, 38.1, 31.8, 30.1, 26.5, 23.2, 20.83, 20.8; positive ion HRESIMS [M+Na]+ m/z 381.1676 (calcd. for  C21H26O5Na, 381.1678). O OBrBrNBS, ABCNCCl4, reflux KOAc, DMF40 ° C, 35%over 2 steps OOAcOAc5.41 5.425.18  221   Figure 5.35 1H and 13C NMR spectra of 5.42 in acetone-d6 at 600 MHz and 150 MHz    222 Preparation of 5.43:  O2 gas was bubbled through a solution of 5.42 (0.01 g, 0.028 mmol) and tBuOH (~1 mL) in a round bottom flask for 5 minutes and then KOtBu (0.006 g, 0.56 mmol) was added into the reaction mixture in three aliquots. O2 was then reintroduced and bubbled through the mixture for 4 hours at which point the reaction was stopped and saturated NH4Cl (0.5 mL) was added. The reaction was concentrated under a stream of N2, 30 mL of H2O was added and the mixture extracted using EtOAc (3 X 10 mL).  The organic layers were combined, concentrated and the product purified using silica gel chromatography (eluting with EtOAc/hexanes (1:1)) to give 6, 7-bis(hydroxymethyl)-4a-methyl-2, 3, 4, 4a, 10, 10a-hexahydrophenanthren-9(1H)-one (5.43) as a white solid (0.005 g, 65 %). 1H NMR (600 MHz, acetone-d6) δ 7.98 (s, 1H), 7.58 (s, 1H), 4.78 (d, J = 5.1 Hz, 2H), 4.70 (d, J = 5.1 Hz, 2H), 4.43 (t, J = 5.1 Hz, 1H), 4.30 (t, J = 5.1 Hz, 1H), 3.05 (dd, J = 17.4, 4.4 Hz, 1H), 2.47 (bd, J = 13.1 Hz, 1H), 2.34 (d, J = 17.5 Hz, 1H), 1.59–1.47 (m, 4H), 1.39 (obs, 1H), 1.36 (s, 3H), 1.94–1.43 (bm, 2H); 13C NMR (150 MHz, acetone-d6) δ 197.3, 148.9, 147.2, 138.3, 131.5, 127.0, 126.0, 62.6, 62.5, 43.8, 42.4, 39.0, 38.3, 32.0, 30.4, 26.6, 23.3. positive ion HRESIMS [M+H]+ m/z 275.1648 (calcd. for C17H23O3, 275.1647).     OOAcOAc O2, KO t -Bu,t -BuOH OOHOH 5.435.42  223    Figure 5.36 1H and 13C NMR spectra of 5.43 in acetone-d6 at 600 MHz and 150 MHz    224 Preparation of 5.31:  To a round bottom flask with a west condenser attached to the top, a solution of 2-hydroxy-1, 4-napthoquinone (0.02 g, 0.12 mmol) and 5 mL of ACN/EtOH (1:1), was added hypotaurine (0.012 g, 0.12 mmol) dissolved in H2O (0.2 mL).  A drop of TFA was added to the reaction mixture, and a balloon of O2 was affixed to the top of a west condenser placed onto the reaction flask.  The reaction mixture was refluxed for three days, with another equivalent of hypotaurine added on the second day, after which time the reaction was cooled and concentrated under reduced pressure to yield a dark red glass.  This was fractionated using C18 reversed phase column chromatography (eluting with a step gradient from H2O to MeOH with 10% MeOH steps) to give purified 3, 4-dihydro-2H-naphtho[2, 3-b][1, 4]thiazine-5, 10-dione 1, 1-dioxide (5.31) (0.014 g, 46%). 1H NMR (600 MHz, DMSO-d6) δ 8.98 (bs, 1H), 7.95 (d, J = 3.8 Hz, 2H), 7.88 (t, J = 6.8 Hz, 1H), 7.77 (t, J = 6.8 Hz, 1H), 3.85 (m, 2H), 3.38 (m, 2H); 13C NMR (600 MHz, DMSO-d6) δ 178.5, 174.8, 146.8, 135.1, 133.1, 132.4, 129.8, 126.3, 125.9, 111.0, 48.2, 39.3; negative ion LRESIMS [M-H]- m/z 262.1 (calcd for C12H8NO4S, 262.0).208             S OOO ONH 5.31  225 Preparation of 5.14:  To a round bottom flask 5.13 (0.5 g, 2.0 mmol) was combined with AcOH (2 mL) and a solution of 45% v/v AcOH/H2O (4.1 mL) and CrO3 (1 g, 1.0 mmol) were added. The reaction mixture was stirred for 2 days after which point it was filtered over a silica gel frit and washed using DCM. The filtrate and wash were then concentrated under reduced pressure.  Silica gel column chromatography (eluting with (3:20) EtOAc/hexane), gave unreacted starting material (5.13) (0.283 g) and 6-(2-bromoacetyl)naphthalene-1, 4-dione (5.14) (0.071 g, 30% BRSM). 5.14 degraded very quickly and as such was used with only 1H NMR characterization. Both 13C NMR and mass spectral experiments gave results showing the material had begun to degrade or decompose. 1H NMR (600 MHz, CDCl3) δ 8.61 (bs, 1H), 8.34 (dd, 2.0, J = 8.4 Hz, 1H), 8.21 (bd, J = 7.9 Hz, 1H), 7.06 (d, J = 4.1 Hz, 2H), 4.50 (s, 2H); 13C NMR (150 MHz, DMSO-d6) δ 191.1, 184.2, 184.1, 139.1, 139.0, 137.9, 134.4, 133.6, 131.9, 126.6, 125.9, 34.4.     OO O BrO Br CrO3, AcOHH2O 5.145.13  226    Figure 5.37 1H NMR spectrum in CDCl3 and 13C NMR spectrum in DMSO-d6 of 5.14 at 600 MHz and 150 MHz    227 Preparation of 5.44 and 5.45:  To a round bottom flask was added 5.14 (0.02 g, 0.07 mmol) in 4 mL of ACN/EtOH (1:1).  To this was added hypotaurine (0.011 g, 0.105 mmol) in 0.3 mL of H2O and the reaction was allowed to stir at room temperature overnight.  After 18 hours the reaction was concentrated and fractionated using C18 reversed phase column chromatography (eluting with a stepwise gradient from H2O to MeOH with increases in 10% MeOH for each fraction).  The (1:5) MeOH/H2O fraction contained the desired product and C18 reversed phase HPLC (20% ACN/H2O) gave purified 5.45 (retention time: 13.5 min, 1.1 mg) and 5.44 (retention time: 14.7 min, 2.5 mg) as yellow solids. Characterization for 5.45:  8-(1, 1-dioxido-3, 4-dihydro-2H-1, 4-thiazin-5-yl)-3, 4-dihydro-2H-naphtho-[2, 3-b][1, 4]-thiazine-5, 10-dione 1, 1-dioxide (5.45); 1H NMR (600 MHz, DMSO-d6) δ 9.29 (s, 1H), 8.11 (s,1H), 8.04 (s, 2H), 7.68 (bs, 1H), 5.56 (s, 1H), 3.86 (m, 2H), 3.80 (m, 2H), 3.39 (m, 2H), 3.18 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 178.3, 173.8, 149.4, 146.4, 138.7, 133.3, 132.9, 130.4, 126.2, 124.1, 94.0, 48.0, 46.4, 39.5, 39.1 (all resonances from 2D NMR data); negative ion HRESIMS [M-H]- m/z 393.0217 (calcd. for C16H13O6N2S2, 393.0215).     OO O Br HNS OOO O HN S OOH2 N SO 2 H5.14 5.44 5.45NHS OOOO HN S OOACN, EtOH,H2 O +  228   Figure 5.38 1H and gHSQC NMR spectra of 5.45 in DMSO-d6 at 600 MHz   229  Figure 5.39 gHMBC NMR spectrum of 5.45 in DMSO-d6 at 600 MHz Characterization of 5.44:  7-(1, 1-dioxido-3, 4-dihydro-2H-1, 4-thiazin-5-yl)-3, 4-dihydro-2H-naphtho[2, 3-b][1, 4] thiazine-5, 10-dione 1, 1-dioxide (5.44). 1H NMR (600 MHz, DMSO-d6) δ 9.29 (s, 1H), 8.12 (d, J = 2.3 Hz, 1H), 8.04 (d, J = 8.0 Hz, 1H), 7.90 (dd, J = 8.0, 1.7 Hz, 1H), 7.69 (bs, 1H), 5.56 (s, 1H), 3.87 (m, 2H), 3.81 (m, 2H), 3.39 (m, 2H), 3.19 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 178.1, 173.9, 149.9, 147.0, 141.3, 132.6, 131.0, 130.7, 126.9, 123.8, 111.2, 94.4, 48.2, 46.6, 40.0, 39.32; negative ion HRESIMS [M-H]- m/z 393.0212 (calcd. for C16H13N2O6S2, 393.0215). HNS OOO O HN SOO5.44  230   Figure 5.40 1H and 13C NMR spectra of 5.44 in DMSO-d6 at 600 MHz and 150 MHz    231 Preparation of 5.47:  A solution of 5.46 (0.1 g, 0.588 mmol), CrO3 (0.294 g, 2.94 mmol), AcOH (1 mL) and 45% v/v AcOH/H2O (2.5 mL) was stirred together in a round bottom flask. The reaction mixture was stirred for 2 days after which point it was filtered over a silica-gel frit and washed using DCM. The filtrate and wash were then concentrated under reduced pressure.  The reaction mixture was purified using silica gel column chromatography (eluting with (3:20) EtOAc/hexanes) to give unreacted 5.46 (0.06 g) and 6-acetylnaphthalene-1, 4-dione (5.47) as a dark yellow solid (0.015 g, 32.5% BRSM). The product quinone was found to spontaneously degrade upon exposure to solvents for long periods (~ 4 hours). 1H NMR (600 MHz, DMSO-d6) δ 8.32 (s, 1H), 8.29 (d, J = 8.5 Hz, 1H), 8.01 (d, J = 8.6 Hz, 1H), 7.10 (s, 2H), 2.65 (s, 3H); 13C NMR (150 MHz, DMSO-d6) δ 197.1, 184.2, 184.1, 140.4, 139.0, 138.9, 133.9, 133.0, 131.7, 126.5, 125.1, 27.1; HREIMS [M]+ m/z 200.04757 (calcd. for C12H8O3, 200.04734).        OO OO CrO 3, AcOHH2O5.46 5.47  232   Figure 5.41 1H and 13C NMR spectra of 5.47 in DMSO-d6 at 600 MHz and 150 MHz   233 Preparation of 5.32 and 5.33:  A round bottom flask containing 5.47 (0.01 g, 0.05 mmol), hypotaurine (0.008 g, 0.07 mmol) in H2O (0.5 mL) and 3 mL ACN/EtOH (1:1) was allowed to stir at room temperature overnight.  After 18 hours the reaction was concentrated and fractionated directly via C18 RP-HPLC (eluting with (1:4) ACN/H2O (retention time = 22 min)) to give a so far inseparable mixture of regioisomers 5.32 and 5.33 (0.003 g, 15.1%). 1H NMR (600 MHz, DMSO-d6) δ 9.26 (bs, 1H), 8.45 (d, J = 1.7 Hz, 1H), 8.27 (dd, J = 8.0, 1.8 Hz, 1H), 8.13 (d, J = 8.14 Hz, 1H), 3.87 (m, 2H), 3.39 (m, 2H), 2.70 (s, 3H); 13C NMR (150 MHz, DMSO-d6) not recorded; positive ion HRESIMS [M+Na]+ m/z 328.0256 (calcd. for C14H11NO5NaS, 328.0256).           OO O HNSOO OOO5.47 5.33 NHSOO O O5.32O+H2N SO2HACN, EtOH,H2O  234  Figure 5.42 1H NMR spectrum of 5.32/ 5.33 mixture in DMSO-d6 at 600 MHz           235 Chapter 6: Chemical Probe Synthesis, Cellular Target Identification, and Structure Activity Relationship Studies of Latonduine A: A Trafficking Corrector of the Cystic Fibrosis Transmembrane Conductance Regulator (CFTR) for the Treatment of Cystic Fibrosis. 6.1  Cystic Fibrosis Cystic Fibrosis (CF) is the most common fatal genetic disease in people of European descent, occurring in one in every 2,500 live births.217 It is an autosomal recessive genetic disease first characterized by Dorothy Andersen and Guido Fanconi in the 1930’s.219 In homozygous individuals, life expectancy is severely reduced (~48.5 years) with the disease adversely affecting the mucosal membranes in the lungs, intestines, liver, pancreas, and several other organs including the testes, making males and often females infertile.219 The most notorious symptoms associated with CF are those related to the lungs. In CF patients, there is a decrease in the volume of airway surface liquid (ASL) that leads to the collapse of lung and airway cilia necessary for airway clearance.  Therefore, microorganisms inhaled into the lungs cannot be cleared and cause infection, inflammation, and fibrosis. It is estimated that in some populations, one in every 25 individuals is carrying one aberrant copy of the gene responsible for CF.219 Heterozygotes, however, show no symptoms and have normal life expectancies, leading some to postulate a heterozygous advantage against various bacterial pathogens as a mechanism for persistence of CF in the gene pool.220, 221 As of 2014, no direct advantage has been found for heterozygotes with a specific pathogen, though some general advantages have been noted.220, 222 6.1.1 The CFTR Protein The cause of CF is a mutation in the gene encoding for the Cystic Fibrosis Transmembrane Conductance Regulator (CFTR), a 1,480-residue member of the ATP binding cassette (ABC) family of proteins.217 It is the only member known to act as an ion channel, transporting chloride and bicarbonate ions for water homeostasis at epithelial surfaces.217 As with all ABC family members, CFTR contains two transmembrane domains (TMD) and two nucleotide-binding domains (NBD), but is unique in its extracellular regulatory region (R) and intracellular N- and C- terminal extensions.217 The protein forms a pore at the epithelial apical cell surface with the two TMDs, each containing six α-helices.217 The active transport of anions out of the cell occurs after the binding of two molecules of ATP into the NBDs, causing a   236 conformational shift to open the pore.217 The correct orchestration of the dynamic portions of CFTR architecture leads to the need for a high degree of quality control in production and folding of this peptide.217 In normal cells only 30% of the synthesized polypeptide is correctly folded and trafficked, using protein chaperones and co-chaperones (i.e. calnexin, HSP-70), to the cell surface to reach functional maturity.217, 218 The remaining 70% is lost due to protein quality control mechanisms present mainly in the Endoplasmic Reticulum (ER) where CFTR is folded.217 If CFTR is properly folded and survives ER protein quality control mechanisms (i.e. ERAD), then, it is shipped off to the Golgi apparatus via COP-I/II vesicles for maturation. Final trafficking to the plasma membrane then occurs via the endosome network (Figure 6.1).217  The protein is stable for up to 24 hours, after which point it is removed, again by the endosomal network, and degraded by lysosomes.217 Due to the size of CFTR, and the complexity of processing, nearly 2000 mutations, causing issues with the polypeptide itself or its processing, are known to cause CF.217, 218 The most common mutation, present in ~90% of patients, is the deletion of a phenylalanine residue at position 508 in CFTR (delF508).219 CFTR mutations are classified into five different groups based on their protein expression: class I mutations are characterized by lack of CFTR protein synthesis; class II, by inadequate protein processing; class III, defective regulation of CFTR; class IV, abnormal CFTR conductance at the cell surface; and class V mutations, reduced quantity of CFTR. Not all classes have similar phenotypic results, and classes I–III are often associated with the most severe disease phenotypes.223 delF508-CFTR is a class II mutation causing major issues with trafficking to the cell surface, often resulting in degradation in the ER likely as a result of prolonged association with protein chaperones such as calnexin.217, 223 delF508-CFTR also shows a shorter half-life if it is able to escape the ER, being ubiquitinylated and degraded by “periphery quality control” mechanisms.219 Despite the issues associated with the F508del mutant and others, many mutant proteins (including delF508-CFTR) have been shown to function if allowed to reach the cell surface and escape degradative enzymes.224, 225     237  Figure 6.1 Trafficking of WT and mutant CFTR in the cell.217–223 6.2 Cystic Fibrosis Treatments 6.2.1 Current Clinical Treatments The majority of the current clinically approved treatments for CF are geared towards alleviating the symptoms associated with a disease phenotype.  As such, various anti-inflammatory treatments such as α1-antitrypsin have been developed.223 Other symptom alleviating treatments such as solutions of inhaled mannitol or hypertonic saline solutions have been shown to improve the ASL fluid layer by creating an osmotic gradient.223   Treatments also Mutant CFTRWild Type CFTR ERGolgiPlasma MembraneEndosome EndosomeRemoval lysosomeGlycosylation/ post-translational modification Translation Initial FoldingCOPI/II vesicles COPI/II vesiclesUbiquitinylationGlycosylation/ post-translational modificationERAD ProteasomeUnfolded CFTR PolypeptideNucleus  238 include the use of inhaled antibiotics for patients suffering from severe and repeated bacterial infections.  Antibiotics like tobramycin (6.1, Figure 6.2) are frequently used to treat the chronic infections of Pseudomonas aeruginosa that occur in 60–80% of CF patients.223   Recently, a new approach towards treating CF came to fruition with the approval of Ivacaftor (Kalydeco®) (6.2, Figure 6.2).217–219 Ivacaftor is a potentiator approved for use in patients presenting G551D-CFTR, a mutation which presents normal trafficking to the cell surface, but fails to provide anion transport.219, 223  This compound binds to G551D-CFTR present on the cell surface and restores anion transport ability, allowing the protein to function and alleviating the disease phenotype.  The G155D mutation is relatively rare however, accounting for roughly 5% of those patients with CF.217, 219 Spurred on by the success of Ivacaftor (6.2, Figure 6.2), the search for new treatments aimed at correction of the underlying cause of CF are currently in preclinical and clinical development. 6.2.2 CFTR Trafficking Correctors Over the past decade a number of cell-based assay screens were developed in the hopes of finding compounds that could correct the underlying protein folding and trafficking issues associated with mutant CFTR.  These screens identified a large number of chemical motifs, including some that have already attained clinical approval [e.g. sildenafil (6.3, Figure 6.2)], which displayed correction activity.219, 226 However, many of these compounds appear to act through unknown mechanisms.  Some of the correctors of CF, as outlined by Birault et al., have mechanisms of action which are known and these can be defined in two broad categories: pharmacological chaperones and proteostasis modulators (PM).219  Pharmacological chaperones are compounds, which are proposed to bind directly to mutant CFTR and stabilize its structure, allowing it to pass through protein quality control mechanisms and be transported to the cell surface. Examples of these are shown in Figure 6.2 and include the correctors CF-Corr-4a (6.4), VRT 325 (6.5), VX-809 (Lumacaftor) (6.6) currently in clinical trials, and VX-661 (6.7).219, 223  All of these chaperones are proposed to interact with CFTR, and likely do not act as chaperones with other misfolded proteins. Exceptions to this rule of course are proteins containing similar structure motifs.    Proteostasis modulators are compounds that permit expression of mutant CFTR through putative evasion of protein quality control mechanisms.  These compounds make up the majority   239 of those identified, and have offered insight into many of the post translation modifications necessary in protein maturation. A brief discussion of several promising members of this group will be presented below.  One class of PMs found to effectively correct CFTR trafficking, are histone deacetylase (HDAC) inhibitors. Histone deactylases are enzymes involved in a myriad of different cellular functions from gene expression to regulation of protein function.  SAHA (6.8, Figure 6.2), a clinically approved HDAC inhibitor, has been shown to be a corrector of the delF508-CFTR, likely inhibiting HDAC6, which has a regulatory role involved in protein quality control.219, 223  Another important class of PMs that have shown correction in high through put screening are phosphodiesterase (PDE) inhibitors.219 Due to the fact that several PDE5 inhibitors have already been approved for clinical use, such as sildenafil (6.3, Figure 6.2), an open label study has begun to determine if they can decrease inflammation in patients with CF and provide correction.219, 227 Cyclooxygenase inhibitors, as a class, also show CFTR trafficking correction.219 These compounds have been used for many years in the clinic to reduce inflammation in patients with CF and have been known to slow the progression of the disease when taken for long periods.219 Several other classes of compounds, including protein kinase inhibitors and cardiac glycosides, have been implicated as PMs, but many of these show severe therapeutic side effects that preclude their use.219     240  Figure 6.2 Various treatments of CF in the clinical or preclinical stage of development219, 223 6.3 Elucidation of the Enzymatic Target of Latonduine A for the Correction of F508del-CFTR In 2003, the oroidin (Chapter 1, Fig. 1.11) related marine alkaloid latonduine A (6.9, Scheme 6.1) was isolated from Stylissa carteri by David E. Williams along with two analogues of latonduine B, as methyl and ethyl esters.78 These three compounds were isolated without biological activity, and in order to confirm the proposed structures, Roger Linington accomplished a total synthesis of latonduine A (6.9).78 OHO NH2 OH2N O O NH2OHOH OHNH2H2N HOtobramycin (6.1)NHO NHO OHivacaftor (6.2)S NHNOONN NNOOsildenafil (6.3) OOFF O NH OHOlumacaftor (6.6)OOFF O NH NF OHOHOHVX661 (6.7 )NNNNO SO O OVRT-325 (6.5)MeO HNCl NS NS NHOCF-Corr 4a (6.4) HN O NHO OHSAHA (6.8 )  241  Scheme 6.1 Latonduine A (6.9) and B (6.10) and the previous total synthetic route for latonduine A (6.9).78 Within a year of their discovery, Drs. Graeme W. Carlile, David Y. Thomas, and John W. Hanrahan at McGill University screened the latonduines, along with several other purified marine natural products, in a trafficking assay for delF508-CFTR.79 This assay, which has been used to screen diverse compound libraries, measures expression of mutant delF508-CFTR.226 In order to quantify expression of CFTR on the cell surface, the mutant CFTR gene is encoded with a triple hemagglutinin (3-HA) tag in the fourth extracellular loop.226  The tag was not found to interfere with the functioning or expression of the CFTR protein, either WT or mutant forms, at the cell surface.226 The 3-HA tag if correctly trafficked to the cell surface on the mutant CFTR protein, offers a mechanism by which a fluorescent dye (3, 3’, 5, 5’-tetramethylbenzidine) can be applied and expression can be quantified at the cell surface.226  This assay then allows for a direct quantifiable method for high-throughput screening (HTS) of small molecule correctors of trafficking.226 The halide efflux assay is often used in conjunction with the HTS assay, and it measures iodine efflux from treated cells as an approximate measure of halide ion channel expression on the cell surface.226 Latonduine A (6.9, Scheme 6.1) showed a considerable ability to traffic mutant CFTR to the cell surface in the HTS assay, and provided functional halide efflux in the iodine efflux assay, reaching levels close to 45% of those seen in wild type cells.79 Subsequent studies in H2N OEtOEtACN, NEt398% HN NHBrBr OMeSO3H, 35 ° C, 7 days65-69% catechol borane rt, THF, LiBH484% NH NHBrBr O OHDMPTHFNH NHBrBr O Oethyl-orthoformateTFA, reflux35%NH NHBrBr O OOEtNH NHBrBr O NN NH2 K2CO3, H2O/THF 85%HNBrBr CCl3O HNBrBr ONH OEtOEtNH NHBrBr O NN NH2 NH NHBrBr O NN NH2RO2C R= Me      Et90%6.9 6.106.9 6.116.12guanidine  242 delF508-CFTR mouse models confirmed the cell-based studies.  Mice fed with 50 mg/kg of latonduine A (6.9) by oral gavage once daily for 2 days showed a salivary increase to 9% of those seen in WT mice, while mice treated with vehicle (DMSO) alone showed only 0.12% of salivary levels seen in WT mice.79 These results established a definitive physiological response to the latonduines and a need for further studies on these compounds. Once the activity of latonduine A (6.9, Scheme 6.1) in vivo was established, determining the mechanism of action was necessary for further pre-clinical development. Dr. Graeme W. Carlile performed several assays in order to elucidate a plausible mechanism of action for latonduine. Pathways for correction such as CFTR gene modulation, direct binding to CFTR protein, or modulation of CFTR synthesis were all ruled out as possible modes of action.79   Due to the lack of evidence for a target, either within CFTR or the cell, we embarked on a plan to synthesize latonduine analogues to be used as activity based protein profiling (ABPP) probes (Figure 6.3).79, 228–231 This technology had been previously used to characterize the members of an enzymatic class (i.e. cysteine proteases) present in a given cell type through either covalent linking, or strong binding affinity to an inhibitory probe.228, 229 Figure 6.3 General conceptual design of an ABPP tool and “Pull Down” experiments to separate an enzyme linked product.229, 231   243 Due to a lack of highly electrophilic or nucleophilic moieties present on the latonduines, we surmised they would not likely bind covalently to a putative enzyme target, though their inhibitory activity was potent (~EC50 = 8 nM), hinting at a specific enzyme interaction.79 The need for a mechanism to covalently link latonduine to its target led us to envision using a photo-inducible covalent linker.  “Photoaffinity elements” such as benzophenones (Scheme 6.4) are generally unreactive in vivo, but become reactive in the presence of light, making them benign additions to probe molecules.230, 231 In addition, an alkyne amendment to the latonduine scaffold would provide a “click element”, to which biotin linked to an azide could be attached through the copper (I) mediated “click” [3+2] cyclization reaction to furnish a triazole.229 Biotinylation offers a mechanism for separation of the enzyme/latonduine complex from the rest of the cellular milieu (Figure 6.3).  This separation can be achieved using streptavidin affinity column chromatography followed by SDS-Page, to furnish latonduine ABPP labeled protein for further analysis.229, 231   6.3.1 Construction of Latonduine A ABPP Probes The original synthesis by Roger Linington in the Andersen lab provided an efficient route to milligram quantities of latonduine A (6.9, Scheme 6.1), however, the synthetic route offers relatively few opportunities for amendments or substitutions.78 As such, only a small number of analogues had been constructed up to the time I began working on this project.  Figure 6.4 below shows the various substitutions that were made and their impacts on the biological activity of latonduine A (6.9).  Figure 6.4 Latonduine scaffold SAR N NHBrBr ON NNH2 HN NHBrBr ON NH2NR1R isolatonduine A ( 6.13 )InnactiveRegiochemistry requiredSubstitution with CO2MeCO2Et toleratedMethylation toleratedbiotinylaion reduced trafficking  244 As can be seen from Figure 6.4, both methylation of the pyrrole nitrogen or carboxylate substitution on the pyrimidine were the only known, at that time, amendments that were tolerated. Intriguingly, isolatonduine A (6.13, Figure 6.4) showed no corrector activity despite a high degree of similarity with latonduine A (6.9, Scheme 6.1).  Due to the near complete retention of activity with pyrimidine substitution in latonduine B (6.10, Scheme 6.1), early efforts for probe construction focused on modifications on the pyrimidine.  Initially, we devised a synthetic route to access compounds 6.14 and 6.15 (Scheme 6.3) for Heck coupling between the pyrrole and bromo-pyrimidine seen in Scheme 6.2.  This, we hoped, would provide a general route towards latonduine and an ABPP analogue (6.16). We began with the necessary starting materials for synthesis of latonduine A (6.9, Scheme 6.1). Starting with dibromination of 4-methyl pyrimidine (6.17, Scheme 6.2), followed by nucleophilic displacement of the primary bromide in 6.18 by azide, gave 6.19 (Scheme 6.2). Subsequent Staudinger reduction and haloform reaction furnished the desired Heck reaction starting material 6.14. Displacement of the primary bromide in 6.18 by pyrrole-2-carboxylic acid (Scheme 6.3) furnished 6.15 in high yields.  Both of these substrates would be used to screen conditions for Heck coupling. We had hoped that once Heck coupling conditions were in hand we could employ a 6-substituted-4-methyl aminopyrimidine starting material  (Scheme 6.2) to access latonduine B (6.10, Scheme 6.1), and proposed ABPP analogue 6.16 (Scheme 6.2).  Unfortunately, all attempts at realizing the Heck coupling for ring closure proved unsuccessful using both substrates 6.14 and 6.15. The reaction conditions screened for the Heck reactions are summarized in Table 6.1  Scheme 6.2 Proposed Heck coupling to form latonduine A (6.9) and analogues. HN XO N NNH2Br NH XON NNH2X = O, N R RR = CO2Me,     H,     COPh N NHON NNH2OBrBrProposed latonduine ABPP ( 6.16 )N NNH2BrR Br  245    Scheme 6.3 Synthesis of 6.14 and 6.15  Table 6.1 Conditions screened for Heck coupling using 6.14 or 6.15 as starting materials. Metal/ Ligand Solvent Base Temp Products 10-20 % Pd(dba)2 “ “ “ “ DMF “ “ DMAc Dioxane AgCO3 KOAc NEt3 “ “ 140° C “ “ “ 100° C NA NA DesBr DesBr ~10 % DesBr 10% Pd(dppf)Cl2 . DCM DMF NEt3 140° C ~10 % DesBr 10% Pd(OAc)2 DMF NEt3 “ ~10 % DesBr 10% Pd(TFA)2 DMF NEt3 “ ~10 % DesBr  Due to disappointing results from our cross coupling attempts, we decided on alkylation of the pyrrole NH to introduce ABPP functionality onto latonduine A (6.9, Scheme 6.1). Dr. Javier Garcia from our lab had previously designed a benzophenone photo-affinity probe (6.20, Scheme 6.4).  We proposed attachment of this probe (6.20) at the latonduine pyrrole nitrogen would provide a latonduine ABPP probe (6.23, Scheme 6.4).79  The route to formation of the latonduine probe (6.23, Scheme 6.4) is presented in Scheme 6.4. Once the latonduine probe 6.23 was constructed, our collaborators at McGill University showed that the probe (6.23, Scheme 6.4) presented no trafficking activity in the whole cell HTS assay, likely owing to its size and inability to penetrate the cell membrane (Figure 6.5).79 N NNH2 Br2, AcOH2 hrs N NNH2Br Br NaN3 N NNH2Br N3DMFHN OHOK2CO3,DMF, rtAr88%HN OO N NNH2Br1). i. PPh3, THF, ! ii. NH4OH, !, 2hrs2). HN CCl3ONEt3, ACN30% over 2 steps98% HN NHO N NNH2Br6.15 6.146.17 6.18 6.19  246  Scheme 6.4 Latonduine probe synthesis   Figure 6.5 Assay for trafficking correction of latonduine photo-affinity probe (6.23) The ABPP pull-down experiments were performed on cell lysates in spite of the lack of CFTR trafficking in whole cells, and proved successful at separating protein targets.  An overview of the experimental procedure used by Dr. Graeme Carlile is available in the original manuscript.79 Protein purification and mass spectrometry experiments identified poly [ADP-ribose] polymerase-1 (PARP-1) as a likely target. Subsequent testing of PARP-1, -2, -3, -4, -5a, and -5b with latonduine A (6.9, Scheme 6.1), showed latonduine A (6.9) was a weak µM HO O OH BrDMF, K2CO3rt, 18hrs HO O OBr BrDMF, K2CO3rt, 18hrsBr O O ONHN O BrBrNNH2N K2CO3,  DMFrt, 1 day 6.206.21 6.2236.9O O O6.23Test to determine whether azido-latonduine enteredthe cell and had an effect100µM 10µM 1µM 100nM 10nM 1nM 100pM 10pM-1.0-0.50.00.51.01.52.02.53.03.5LatonduineAzido-latonduineTreatment6.23 !(6.9)   247 inhibitor of PARP-1, -2, -4, -5a, and -5b, but a selectively potent inhibitor of PARP-3 (EC50 = 400 pm).79  It was also shown through competitive binding studies that latonduine A (6.9) acted as an inhibitor of the PARP family in the same site as ABT-888 (6.24, Figure 6.6). ABT-888 has been shown in crystallographic studies to be an NAD mimic, binding to the NAD+ pocket of the catalytic active site.79 This suggests that the latonduines also bind to PARP enzymes in the NAD+ binding site.  Figure 6.6 ABT-88879  The selectively strong inhibitory activity of latonduine A (6.9, Scheme 6.1) against PARP-3 was linked to CFTR correction through a small interfering-RNA (si-RNA) experiment.79 In this experiment, cells were treated with si-RNA for either PARP-1, -3, or -5a for two days, after which, they were treated with latonduine A (6.9) in a dose-dependent manner.79 The results showed that the knockdowns of PARP-1 and -5a presented no increased sensitivity to latonduine A (6.9).79 PARP-3 knockdown cells, however, showed a highly increased sensitivity for correction by latonduine A (6.9), dropping the EC50 from 8 nM to 200 pM in the HTS assay.79 Several other experiments, not listed here, were performed in order to provide corroborating evidence for a mechanism of trafficking by latonduine A (6.9, Scheme 6.1) that required PARP-3 inhibition.79 Through these experiments, PARP-3 inhibition was shown to be a novel mechanism by which delF508-CFTR could be trafficked to the cell surface and provide relief of the disease phenotype.79  However, further SAR studies around the latonduine scaffold, and an improved synthesis of latonduine A (6.9) or analogues are still needed to provide sufficient material for further development.   6.4 PARP Inhibitors As Trafficking Correctors of F508del-CFTR 6.4.1 The PARP Family of Enzymes Poly [ADP-ribose] polymerases (PARPs) are a group of 17 enzymes that catalyze the attachment of ADP ribose to a protein through loss of nicotinamide from NAD+ (Figure 6.7).232, NHNOH2N HNABT-888 ( 6.24 )  248 233 Several members of the family, PARPs/ARTDs -1–6 (tankyrase-2), catalyze attachment of polymeric chains of ADP-ribose of various lengths to different substrates within the cell (Figure 6.7).232, 233  All other members of the PARP family (PARP-7–17) are either presumed or proven to only attach one ADP-ribose unit at a time; this has led to a reclassification of the PARP family as ARTDs (diphtheria toxin like mono or poly ADP-ribosyltransferase (mART)).232, 233    Figure 6.7 Mechanism of ADP-ribose attachment to protein substrate  The PARP/ ARTD enzyme family has been linked to many important physiological roles within cells.232 PARP-1 and -2 have been shown to be involved in repair of single strand breaks in DNA, recruiting DNA polymerase-β and DNA ligase-III, and in DNA base excision repair.232 PARP-1 and -2 have also been shown to be involved in chromatin reorganization and transcriptional regulation, which are important functions related to mitosis.232 This function makes PARP-1 and -2 essential enzymes for the repair of DNA damage and the lack of PARP activity in cells and in knockout mice was shown to cause increased sensitivity to ionizing radiation and alkylating reagents.232, 233 The essential nature of these enzymes has led to the development of several PARP inhibitors as chemotherapeutic agents to be used in cancer treatment.232 Currently, there are no clinically approved PARP inhibitors available, but there are several, such as ABT-888 (6.24, Figure 6.6), in clinical trials.234, 235 6.4.2 PARP/ ARTD-3 PARP-1 and -3 are closely related, and have been shown to interact with each other in the cell, being implicated in telomere integrity and mitotic spindle stabilization.236 PARP-3 is a NNNN NH2O OHOHOPO OOPOO NO OHOHO O NH2 PARP/ ARTD NNNN NH2O OHOHOPO OOPOON O OHOHOO NH2- ProteinNAD+ ADP-Ribosylated protein  249 60kDa protein expressed primarily in the “neuroglial cells in the brain and spinal cord and epithelial cells forming the ducts of the prostate, salivary glands, liver, and pancreas.”232, 236 It has a N-terminal domain rich in tryptophan, glycine, and arginine (WGR domain) and a C-terminal catalytic domain.236 The catalytic domain of PARP-3 has a high degree of similarity to that of PARP-1 and -2.233 Evidence for involvement of PARP-3 in DNA repair has been reported though its role is not clear.232, 233, 236 In one study, si-RNA treated cells for PARP-3 behave in the same manner as wild type cells when presented with a single strand DNA damaging agent.232 However, more recent studies have linked PARP-3 with double strand break repair in concert with PARP-1.236 It has also been shown that PARP-3 associates with histones H3C and H2BE. All of these associations point to PARP-3 also being involved in gene expression or silencing though direct evidence for this function is absent.237 Lastly, the distribution in the cell of PARP-3 has been reported as either localized to the daughter centriole or the polycomb group bodies, both of which reside within the nucleus.238, 239  6.4.3 PARP 16 and Other PARP Family Members The vast majority of PARPs/ ARTDs have varied functions, some involved in protein regulation and expression (PARP-10 and -15) and others are still unresolved or catalytically unproductive (PARP-9 and -13).233, 240 An example of protein regulation being governed by a PARP family member can be seen with PARP-16 (ARTD-15).  PARP-16 is an endoplasmic reticulum membrane associated protein, with an N-terminal cytosolic PARP catalytic domain, a hydrophobic trans-membrane domain, and a C-terminal ER-luminal domain.241 PARP-16 is the first member of this family of enzymes shown to catalyze ADP-ribosylation of serine and threonine. It has been shown to transduce ER stress from the C-terminal ER domain to the cytosolic domain, and as such, regulates important protein kinases IRE1-α and PERK through ADP-ribosylation.241 ADP-ribosylation of these ER associated protein kinases induces up-regulation and activation, which transduces ER stress from improper protein folding to the nucleus.241 The downstream result of this is the regulatory mechanism known as the unfolded protein response (UPR).241 This translates to PARP-16 playing an important role in cell protein folding regulation and homeostasis.241   250 6.4.4 Synthetic PARP Inhibitors Working with Glaxo-Smith-Kline (GSK), our collaborators at McGill screened a series of synthetic PARP inhibitors to investigate the role of PARP inhibition as a mechanism for correction. In the process they discovered that PARP-16 was also potently inhibited by latonduine A (6.9, Scheme 6.1). PARP-16’s role as a pivotal enzyme in protein regulation gave rise to the proposal that it might be another target of latonduine responsible for CFTR trafficking.  In order to identify if PARP-16 inhibition, PARP-3 inhibition, or both were responsible for the trafficking effects observed, we began construction of a series of analogues related to latonduine A (6.9, Scheme 6.1). Our aim was to synthesize a diverse array of compounds related to the latonduine scaffold in order to increase PARP selectivity while enhancing trafficking correction. From the beginning, we observed that the dihydrobenzazepinone nucleus, present in one potent trafficking corrector 6.25 (Figure 6.8), and known PARP inhibitor rucaparib (6.26, Figure 6.8), was synthetically more accessible than the dibromopyrrole-azepinone core 6.11 (Figure 6.8) seen in the latonduines. The use of both the dibromopyrrole-azepinone and the dihydrobenzazepinone nucleus were both subsequently explored through our synthetic efforts.  Figure 6.8 Latonduine bromopyrrole azepinone and benzaepinone for comparison78, 234 Our screening approach was to test all of the compounds constructed directly in the HTS assay to measure their trafficking abilities.  Subsequently, we would test for PARP-1, -3, and -16 inhibitions regardless of trafficking correction.  Dr. Graeme Carlile at McGill University performed all biological assays including the PARP inhibition assays. These were performed on the catalytic domains of PARP-1, -3, and -16 as isolated proteins in vitro. NH NH NHO OBr Br NH NHBr Br O NN NH2HN NHOHN 6.25ru ca p a ri b 6.26 6.11 6.9  251 6.5 Synthesis of Novel Trafficking Correctors of F508del-CFTR 6.5.1 Pyrrole Azepine SAR The latonduine scaffold was relatively untested in SAR studies for trafficking activity. It was assumed that the pyrrole, azepinone, pyrimidine, and their orientation to one another were requisite attributes for biological activity (see Figure 6.4).  However, this was at the time a relatively untested hypothesis. Initial screening of latonduine synthetic intermediates showed alcohol 6.27 (Figure 6.9) as a promising CFTR trafficking corrector (Table 6.4). 6.27 was isolated as a side product from the hydroboration reaction used to synthesize intermediate 6.12 (Scheme 6.1) in the latonduine A total synthesis. 6.11 (Figure 6.8) also showed trafficking correction (Table 6.4), challenging our previously held assertion that latonduine A represented a minimal pharmacophore.  Figure 6.9 Alcohol 6.27 and olefin 6.11 To further probe the latonduine A (6.9, Figure 6.8) scaffold we wanted to test the orientation of the pyrrole nitrogen relative to the amide in the azepinone.  From previous bioassay results, we observed that the N-H was not necessary for activity (Figure 6.4). Therefore, it is not clear how the pyrrole is functioning in the PARP NAD+ binding pocket.  To test if pyrrole orientation had an effect on trafficking ability, we synthesized simplified analogues 6.31 and 6.32 from 6.28 (Scheme 6.5) using Schmidt rearrangement conditions and bromination (Scheme 6.5). Both showed trafficking correction, illustrating that pyrrole regiochemistry is inconsequential in some cases. As shown in Table 6.4, these compounds also displayed selectivity for PARP-3 and -16 inhibitions, though less effectively than 6.9 (Figure 6.8). Notably, the intermediates 6.29 and 6.30 (Scheme 6.5) showed neither correction activity, nor strong PARP inhibition, underlining the importance of the bromine substituents (Table 6.4). NH NHBrBr OHO6.27 NH NHOBrBr 6.11  252  Scheme 6.5 Pyrrole azepinone regio-isomer synthesis Given the challenging nature of generating multiple dibromopyrrole analogues in an efficient manner, we next opted to explore elaboration of the dihydrobenzazepinone nucleus in 6.25 and 6.26 (Figure 6.8).  6.5.2 Benzazepinone Trafficking Corrector Synthesis In order to synthesize analogues of 6.25 (Figure 6.8), we sought methodology that could provide a diverse array of substituents on the dihydroazepinone portion of the molecule, while we systematically tested the aryl substitution for positive trafficking effects.  We then surmised that combination these two amendments onto a single molecule would furnish a more potent corrector. In the literature, methodology utilizing a light activated phthalimide ring expansion reaction to form substituted [2]-benzazepine-1, 5-diones (e.g. 6.34, Scheme 6.6) appeared promising.243, 244 This reaction type involves the singlet excited state of phthalimide, which then reacts with an alkene (Reaction type I in Scheme 6.6) or tethered ethyl trimethyl silane (6.33, Reaction type II Scheme 6.6) to form an azetidine as shown in Scheme 6.6. The azetidine then undergoes transannular ring expansion to form the benzazepine-1, 5-dione (6.34). 243, 244 These benzazepine-1, 5-diones could then be further functionalized as needed.  Several reports have been published using these reactions to generate many diverse structural motifs.244  NaN 3  (2 eq)HClNH O NH NNH HN N N NO + NH NN N NBr BrNH HNOBr Br NBS, DCM6.28 6.29 6.306.31 6.32  253 Type I  Type II   Scheme 6.6 Phthalimide photochemistry243–245 6.5.3 Ethyl Silyl Tethered Phthalimides Produce Dihydrobenzazepinone  At first, we chose to employ reaction type II, following methodology originally discovered in the Mariano lab.245 Two [2]-benzazepine-1, 5-diones, 6.34 and 6.37 (Scheme 6.7), were synthesized which showed limited delF508-CFTR trafficking (Table 6.4). However, upon reduction with NaBH4 to the corresponding alcohols, compounds 6.35 and 6.38 (Scheme 6.7) showed marked correction activity and selectivity for inhibition of PARP-3 and -16 (Table 6.4). The reaction unfortunately gave 6.34 and 6.37 (Scheme 6.7) in poor yields (<10% overall), and as such, offered little material for further elaboration and testing.  As a result, this methodology was abandoned for alternative methodology presented below to access derivatives of 6.35 and 6.38.  Scheme 6.7 Syntheses of 6.32–6.36 NHOO R1 R2 NHOO R1 R2h!NaOH, ACN NOO R1 R2 H+NOO h ! ACN/H 2O NHOONHOOSiMe 3 H+6.33 6.34NOO Si h ! NHOOacidic work up NaBH 4MeOH NHOHO6.33 6.34 6.35NOO Si h! NHOOacidic work up NaBH4MeOH NHOHOClCl ClCl ClCl6.36 6.37 6.38  254 6.5.4 Phthalimide Photoreactions with Styrene Derivatives Next, we chose to explore the excited state phthalimide photochemistry shown in Scheme 6.6 Type I.243, 245 For these transformations we exclusively followed methodology originally developed in Professor Rafael Suau’s lab. 243, 245 From this, we were able to produce [2]-benzazepine-1, 5-diones in low but consistent yields from various vinyl aryl derivatives (Figure 6.9 and Table 6.2).243 As before, the [2]-benzazepine-1, 5-diones 6.39–6.42 (Table 6.2) showed minimal correction activity (Table 6.4). However, NaBH4 reduction to the corresponding alcohols 6.43–6.46 (Figure 6.9 and Table 6.2), failed to produce a single compound with trafficking correction activity (Table 6.4).  When tested directly for in vitro PARP-1, -3 or -16 inhibition, several intriguing observations were made. Encouragingly, 6.39 (Table 6.2) proved a strikingly potent and selective inhibitor of PARP-3. Contrastingly, pyridyl substituted 6.44 and 6.45 (Table 6.2) showed pronounced and selective inhibition activity towards PARP-16.  This substitution pattern proved a serendipitous discovery for selective PARP inhibition, and 6.44 and 6.45 represent the first selective PARP-16 scaffolds reported to date.  Several vinyl-substituted heterocycles were subsequently attempted as reaction partners, only to give intractable mixtures with no evidence of the desired products (Table 6.2).  Future work on these compounds should focus on 6.39 and 6.45 (Table 6.2) as a core scaffold for elaboration to improve potency against PARP-3 and -16.  Figure 6.10 General reaction sequence for styrene derivatives tested .243 Table 6.2 Photoreaction products and reduction products Cmpd # R1 R2 Alcohol # 6.39 phenyl H 6.43 6.40 2-pyridyl H 6.44 6.41 2-pyridyl Cl 6.45 6.42 3, 4-dimethoxy phenyl H 6.46  4-pyridyl H NA  R-N imidazole H NA  pyrazine H NA  N HOO R 1+ h ! N HOO R 1NaOH, ACNR 2R 2 NaBH 4MeOH NHOHO R 1R 2R 2 R 2R 2  255 6.5.5 Other Alkene Phthalimide Photoreactions Styrene derivatives were not the only alkenes used with the methodology shown above.245 We also employed the protected allylic alcohols TMS-allyl ether and allyl acetate as reaction partners for photocyclization with phthalimide to ultimately provide 6.47 (Scheme 6.8) as the product. This reaction generated the opposite product substitution pattern to that seen above for 6.39–6.42 (Table 6.2), containing a hydroxy methylene substituent preferentially alpha to the amide nitrogen in the lactam 6.47 (Scheme 6.8). The compound produced showed promising trafficking of delF508-CFTR in the HTS assay (Table 6.4), though PARP specificity appears to be reduced compared to latonduine A (6.9, Figure 6.8). Reduction of 6.47 with NaBH4 gave 6.48 (Scheme 6.8) as cis and trans diastereomers, which showed decreased trafficking ability (Table 6.4). Interestingly, only the cis diastereomer 6.48 c (Scheme 6.8) showed CFTR trafficking abilities.  Other substituted alkenes such as methyl vinyl sulfone and ethyl acrylate proved unproductive in this reaction type.  Scheme 6.8 Syntheses of 6.47, 6.48 c, and 6.48 t.  Figure 6.11 General formula for side product NHOO h !NaOH, ACN NHOO NaBH 4MeOH NHOHO OHNHOHO OHOR OHR = Ac      TMS +6.47 6.48 t6.48 cN HOHO RR= CH 2 OH      Phenyl      Pyridyl  256 6.5.6 Schmidt Rearrangement  In order to test the effects of varying phenyl substituents in the benzoazepine scaffold, we decided to utilize Schmidt methodology, combining hydrazoic acid with a substituted α-tetralone to form an azepine.246, 247 For these reactions, we chose only to employ inexpensive commercially available α-tetralones listed below. When using concentrated HCl as the solvent for the reaction, three products can potentially be generated: the tetrazole (A, Figure 6.11), and benzazepinones B and C shown in Figure 6.11.246, 247 Each product (A, B, and C) was tested in the HTS and PARP inhibition assay if they were isolated (Table 6.4).   Figure 6.12 Generalized formula for the Schmidt reaction Table 6.3 Schmidt reaction products and isolated yields. # R1, R2, R3 A B C 6.49 H, H, H 32.6 % 16.4 % 22.1 %  6.50 H, NH2, H 51.3 % NA 23.8 % 6.51 OH, H, H NA 36.8 % 22 % 6.52 H, OMe, OMe 35.4 % 15 % 21.4 % 6.53 H, H, F 33.7 % 19.4 % 14.6 %  6.5.7 First Generation Correctors with Combined Functionality Once we had constructed a small library of basic structural motifs showing trafficking ability, we then set out to design compounds that would combine properties of two simpler analogues.  Initially, we chose to work on 6.52A (Scheme 6.9) as a starting point for further elaboration.  We hoped that benzylic functionalization would allow us to install an alcohol as in 6.35 (Scheme 6.7).  After screening several benzylic oxidation conditions we found that KMnO4 in ACN provided the desired oxidation, giving 6.54 (Scheme 6.9) in the highest yields. Subsequently, NaBH4 reduction produced the desired alcohol 6.55 (Scheme 6.9), however, corrector activity was not increased compared to 6.52A for either 6.54 or 6.55 (Table 6.4).  R1R2R3 H O R1R2R3 H HNR1R2R3 H NN R1R2R3 H NHON N O+ +NaN3 (1.5 eq)[HCl] A B C  257  Scheme 6.9 Synthetic route to 6.54 and 6.55 A second combination we explored was the fluorine substitution seen in 6.53C (Scheme 6.9) with azepine functionality seen in 6.47 (Scheme 6.8).  Using the methodology described above (section 6.55) an undergraduate, Isabel Barne, carried out this reaction and performed the necessary HPLC separations. We employed the same methodology used in Figure 6.8, only substituting 4-fluorophthalimide as starting material (Scheme 6.10).  This reaction produced both regio-isomers 6.56 and 6.57, in a 2:1 ratio (Scheme 6.10).  These compounds displayed excellent correction activity and offered improvements in selectivity towards PARP-3 and -16 over the parent compound 6.47 (Table 6.4).   Scheme 6.10 Synthetic route to 6.56 and 6.57.  Figure 6.13 Trafficking correction data for 6.56 and 6.57 NN N NMeO HOMeONN N NMeO OMeONN N NMeOMeO KMnO4ACN NaBH4MeOH6.52A 6.54 6.55N HOO OHFNHOOF h!N aOH, ACNOTMS NHOO OHF+6.56 6.57Increase in surface CFTR expression signal measured in the HTS assay (corrector ability)100pM 1nM 10nM 100nM 1µM 10µM-1.5-1.0-0.50.00.51.01.52.02.5Ap123 2AAp123 2BConcentration of compound6.57 6.56   258  Figure 6.14 PARP inhibition data for 6.56 and 6.57 6.6 Combining Selective PARP Inhibitors  To test the hypothesis that both PARP-3 and PARP-16 inhibition are required for trafficking seen by the latonduines, we ran a combination inhibitor experiment.  We selected our most potent and selective PARP-16 or PARP-3 inhibitors at that time and combined them in the halide ion efflux assay.  We found that the combination of selective PARP-16 inhibitors (e.g. 6.44, Table 6.2), with compounds displaying high selectivity towards PARP-3 (e.g. 6.38, Scheme 6.7), often produced significant trafficking correction, implying synergistic action. This study adds further evidence for a dual PARP-16 and -3 inhibition mechanism for trafficking correction.          Ability of compounds Ap123-2A and Ap123-2B to inhibit the activity of various members o thePARP family01020304050607080901001101201302A-PARP-12A-PARP-32A-PARP-162B-PARP-12B-PARP-32B- PARP-16PARP and COMPOUND used6.57    6.56    259 6.7 Assay Results Table 6.4 delF508-CFTR trafficking correction and PARP inhibition. # CFTR PARP-1 PARP-3 PARP-16 6.9 ~3 99.1 3.5 5.3 6.11 1.2 97.0 41.6 36.1 6.12 0.3 98.2 ~100 ~100 6.27 1.5 99.6 14.3 26.0 6.29 0 91.2 ~100 52.2 6.30 0 72.1 69.9 46.7 6.31 0 61.4 ~100 49.3 6.32 1.2 (10 nm) 26.6 43.9 52.1 6.35 1.5 99.7 31.2 7.8 6.37 0 ~100 15.1 ~100 6.38 1.4 100 3.7 4.5 6.39 0.2 95.3 82.2 74.8 6.40 0.3 99.1 73.9 65.4 6.41 0.6 ~100 66.7 3.6 6.42 0 91.7 ~100 ~100 6.43 0.03 97.0 9.2 ~100 6.44 0.25 ~100 50.5 1.9 6.45 0 ~100 ~100 ~0 6.46 0 12.6 74.8 8.4 6.47 2.75 44.4 26.7 18.3 6.48 t 0 ~100 ~100 5.1 6.48 c 1.5 ~100 48.9 4.5 6.49A 1.5 ~100 18.6 28.9 6.49B 0.3 ~98.0 75.5 4.8 6.49C 0.3 ~100 ~100 ~100 6.50A 0.5 69.0 ~100 46.6 6.50C 1.5 43.7 26.7 32.9 6.51B 0 67.8 ~100 ~100 6.51C 0.3 56.6 79.8 ~100 6.52A 2.4 39.6 54.5 17.2 6.52B 3.3 45.9 34.9 61.9 6.52C 0 ~100 74.3 ~100 6.53A 0.5 90.9 ~100 ~100 6.53B 2.5 (100 nm) ~100 ~100 6.5 6.53C 5.0 (100 nm) 85.0 36.5 26.1 6.54 0 35.1 ~100 ~100 6.55 1.1 (100 nm) ~100 ~100 0.2 6.56 ~2 ~100 15 17 6.57 ~2 92 35 35 6.38/6.44 ~2.5 NA NA NA Maximal trafficking result (in Standard Deviations above control, significant trafficking >1 SD) in the HTS assay. Trafficking result recorded at 1µM. When not recorded at 1µM, concentration is given. PARP inhibition values represent the amount of PARP activity relative to no inhibitor present. Single concentration used that gave the highest recorded correction value. When no correction was observed, 10 µM inhibitor used.  6.8 Conclusions In collaboration with Prof. Thomas and Dr. Carlisle from McGill University, we were able to show that latonduine A (6.9, Figure 6.8) is an inhibitor of the PARP/ARTD family of   260 enzymes, and exerts its trafficking correction of delF508-CFTR through this mechanism.  This was accomplished in part by using the activity-based profiling probe, which I constructed.  The probe design incorporated a benzophenone as a “photo-affinity element” and an alkyne as a “Click element” for enzyme isolation experiments. The subsequent purification and characterization experiments identified the PARP/ARTD family of enzymes as the biological target for latonduine A (6.9, Scheme 6.1).  Once the enzymatic target family had been identified, the enzymes directly affected by latonduine for trafficking correction were found to be PARP-3 and subsequently PARP-16. Trafficking corrector design, based upon PARP-3 and PARP-16 inhibition, has produced a suite of molecules inspired by the core carbon skeleton of the latonduines.  Of these, several compounds (e.g. 6.52A, 6.52B, 6.47, and 6.56) have shown trafficking ability, matching and surpassing levels seen with the latonduine natural products.  Also of note is the discovery of selective PARP-16 inhibitors (e. g. 6.44, 6.45, 6.48 t and 6.55), of which there have been no previous reports in the literature.  Recently, implications of PARP-16 possibly playing a role in certain cancers may propel the discovery of these potent and selective PARP-16 inhibitors into other applications beyond CF.248, 249 Because latonduine A (6.9, Scheme 6.1) and other trafficking correctors are not CFTR trafficking-specific, their action on the protein quality control pathway may have broad applications in other diseases as well.223 Therefore, the possibility exists of using these PARP-16 inhibitors for the treatment of other diseases resulting from malfunctions in protein trafficking.  Many of these are orphan diseases much rarer than CF (e. g. Hermansky-Pudlak syndrome).224, 249   In conclusion, the discovery of the latonduines and their subsequent target identification has led to several serendipitous discoveries.  These findings have not only led to the creation of new chemical biology tools, which will likely aid in the understanding of protein quality control mechanisms, but they may also offer real clinical value for fatal human diseases. 6.9 Experimental 6.9.1 General Experimental All non-aqueous reactions were carried out in oven dried Pyrex® glassware under an Ar atmosphere unless otherwise noted. Air and moisture sensitive reagents were manipulated using airtight dry syringes.  Anhydrous solvents were all obtained from commercial sources and all   261 reagents were obtained from commercial sources without further purification.  All 1H and 13C NMR spectra were recorded at 600 and 150 MHz, respectively, as indicated and referenced to the internal residual solvent peak denoted in the experimental description.  Flash chromatography was performed using silica gel (230–400 mesh) with the solvent system indicated.  All UV reactions were performed in a photo-reactor with a water-cooled Pyrex® filtered 450 W medium pressure mercury lamp.  Preparation of 6.18:  To a round bottom flask containing a solution of 6.17 (0.5 g, 2.65 mmol) in AcOH (10 mL), Br2 (4.77 mmol, 0.25 mL) was slowly added dropwise over 5 minutes.  The reaction was stirred for 3 hours after which point (~1 mL) 10% Na2S2O3 solution was added until the reaction mixture turned a light yellow. The reaction was concentrated under reduced pressure to give a bright yellow solid.  Flash silica gel column chromatography (eluting with a step gradient of DCM, (1:20) acetone/DCM, (1:10) acetone/DCM, and (1:3) acetone/DCM) gave purified 5-bromo-4-(bromomethyl)pyrimidin-2-amine (6.18) as a white solid (0.234 g, 34%). Subjecting the isolated mono-bromo product and unreacted starting material to the above bromination conditions furnished a further 112 mg of the desired product to give an overall yield of 0.346 g (50.6 %). 1H NMR (600 MHz, DMSO-d6) δ 8.36 (s, 1H), 7.04 (bs, 2H), 4.39 (s, 2H); 13C NMR (150 MHz, DMSO-d6) δ 162.5, 162.4, 160.5, 104.4, 132.4, 32.4; positive ion HRESIMS [M+H]+ m/z 265.8932 (calcd. for C5H6N379Br2, 265.8928).   N NN H 2 Br 2 , AcOH2 hrs N NNH 2Br N NN H 2Br Br N NN H 2Br BrBr+ +6.186.17  262   Figure 6.15 1H and 13C NMR spectra of 6.18 in DMSO-d6 at 600 and 150 MHz   263 Preparation of 6.19:   6.18 (0.52 g, 1.9 mmol) was added to 10 mL of a (1:2) H2O and DMF solution in a round bottom flask.  This was stirred open to air and NaN3 (0.253 g, 3.9 mmol) was added into the reaction mixture in one portion.  The mixture was allowed to stir for 1 hour after which time all starting material had been consumed (TLC).  The reaction mixture was added to ~200 mL of H2O and partitioned with (3 X 70 mL) EtOAc.  The organic layers were combined, dried using MgSO4, filtered, and then concentrated under reduced pressure to yield crude product. The crude reaction mixture was then purified by flash silica gel chromatography (eluting with (1:9) acetone/DCM), to give 4-(azidomethyl)-5-bromopyrimidin-2-amine (6.19) as a fluffy white solid (0.437 g, 98%). 1H NMR (600 MHz, DMSO-d6) δ 8.32 (s, 1H), 7.03 (bs, 2H), 4.39 (s, 2H); 13C NMR (150 MHz, DMSO-d6) δ 162.3, 161.9, 159.6, 103.8, 52.8; HREIMS [M]+ m/z 227.97606 (calcd. for C5H5N679Br, 227.97591).       N NNH2Br Br NaN3 N NNH2Br N3DMF 6.196.18  264   Figure 6.16 1H and 13C NMR spectra of 6.19 in DMSO-d6 at 600 and 150 MHz.    265 Preparation of 6.14:  To conical flask containing a solution of 6.19 (0.05 g, 0.22 mmol) in THF (5 mL) was added polymer bound PPh3 (0.15 g) under Ar atmosphere.  A west condenser was affixed to the top of the flask and the mixture was heated to reflux for 4 hours, then cooled to room temperature and NH4OH (1mL) was added.  The reaction mixture was then heated again to reflux for 1.5 hours, cooled and filtered through a silica gel frit and washed repeatedly with MeOH.  The combined washes were concentrated using rotoevaporation, dissolved in ACN (5 mL) and 2-trichloroacetylpyrrole (0.046 g, 0.22 mmol) was added with NEt3 (0.1 mL).  The mixture was stirred overnight concentrated and directly purified via silica gel flash chromatography (eluting with EtOAc/Hex (1:1)) to give N-((2-amino-5-bromopyrimidin-4-yl)methyl)-1H-pyrrole-2-carboxamide (6.14) as a white solid (0.019 g, 29.8%). 1H NMR (600 MHz, acetone-d6) δ 10.70 (bs, 1H), 8.23 (s, 1H), 7.61 (bs, 1H), 6.98 (m, 1H), 6.80 (m, 1H), 6.36 (bs, 1H), 6.16 (m, 1H), 4.49 (d, J = 5.9 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 163.1, 161.6, 160.1, 158.5, 128.3, 120.9, 114.5, 108.9, 108.4, 42.3; positive ion HRESIMS [M+H]+ at m/z 296.0149 (calcd. for C10H11N5OBr, 296.0147).    N NNH 2Br N3 1). i. PPh 3, THF, ! ii. NH 4OH, !, 2hrs2). HN CCl3ONEt 3, ACN30% over 2 steps HN NHO N NNH 2Br6.146.19  266   Figure 6.17 1H and 13C NMR spectra of 6.14 in acetone-d6 at 600 and 150 MHz.   267 Preparation of 6.15:  To a solution of pyrrole 2-carboxylic acid  (0.021 g, 0.187 mmol) and K2CO3 (0.52 g, 0.374 mmol) in DMF (2 mL), in a round bottom flask under Ar atmosphere was added 6.18 (0.05 g, 0.187 mmol).  This was stirred overnight at room temperature.  The reaction was then poured into 50 mL of H2O and partitioned with (3 X 25 mL) EtOAc.  The organic layers were combined, washed with brine, dried over MgSO4, filtered and concentrated.  The crude solid was then purified using flash silica gel chromatography (eluting with EtOAc/Hex (4:1)) to give (2-amino-5-bromopyrimidin-4-yl)methyl 1H-pyrrole-2-carboxylate (6.15) as a light brown solid (0.052 g, 93% yield). 1H NMR (600 MHz, acetone-d6) δ 11.02 (bs, 1H), 8.28 (s, 1H), 7.09 (m, 1H), 6.90 (m, 1H), 6.29 (bs, 2H), 6.24 (m, 1H), 5.19 (s, 2H); 13C NMR (150 MHz, acetone-d6) δ 163.6, 163.2, 160.7, 160.4, 124.7, 124.6, 116.5, 110.7, 105.5, 64.9; positive ion HRESIMS [M+H]+ at m/z 296.9991 (calcd. for 296.9987, C10H10N4O2Br).        N NNH 2Br Br6.18 K 2CO 3,DMF, rt ArHN OHO HN OO N NNH 2Br6.15  268   Figure 6.18 1H and 13C NMR spectra of 6.15 in acetone-d6 at 600 and 150 MHz    269 Preparation of 6.22:  To flask containing 6.21 (0.25 g, 1.16 mmol) and K2CO3 (0.24 g, 1.74 mmol), DMF (2 mL) was added under Ar at room temperature and stirred for 30 min.  Next, propargyl bromide (12 mL, 1.3 mmol) was added dropwise over a minute and the reaction mixture was stirred overnight.  The reaction mixture was then poured into 60 mL of H2O and partitioned with (3 X 20 mL) EtOAc.  The organic layers were combined and concentrated, then purified using flash silica gel chromatography (310 mm X 15 mm, (2:5) EtOAc/Hex) yielding 0.187 mg (64%) of (4-hydroxyphenyl)(4-(prop-2-yn-1-yloxy)phenyl) methanone (6.22), as a white powder.  1H NMR (600MHz, DMSO-d6) δ 10.35 (s, 1H), 7.69 (d, J = 8.9 Hz, 2H), 7.63 (d, J = 8.9 Hz, 2H), 7.11 (d, J = 8.9 Hz, 2H), 6.89 (d, J = 8.9 Hz, 2H), 4.91 (d, J = 2.5 Hz, 2H), 3.64 (t, J = 2.5 Hz, 1H); 13C NMR (150 MHz, DMSO-d6) δ 193.1, 161.6, 160.2, 132.2, 131.6, 131.0, 128.4, 115.1, 114.5, 78.7, 55.8; positive ion HRESIMS [M+H]+ 253.0868 (calcd. for C16H13O3, 253.0865).        HO O OH BrDMF, K2CO3rt, 18hrs HO O O6.21 6.22  270   Figure 6.19 1H and 13C NMR spectra of 6.22 in DMSO-d6 at 600 and 150 MHz.   271 Preparation of 6.23:  To an oven dried flask containing 6.22 (0.63 mg, 0.248 mmol) and K2CO3 (0.343 mg, 2.48 mg) under Ar, was added DMF (1 mL) at room temperature.  This was allowed to stir for 30 minutes, at which time 1, 3-dibromopropane (0.25 mL, 2.48 mmol) was added in dropwise over 2 minutes.  The reaction mixture was allowed to stir overnight, after which it was poured into 30 mL of H2O and extracted with (3 X 12 mL) EtOAc.  The organic layers were combined, concentrated, and the product purified using flash silica gel chromatography (300 mm X 15 mm, 30% EtOAc in hexanes) to give (4-(3-bromopropoxy)phenyl)(4-(prop-2-yn-1-yloxy)phenyl)methanone (6.20), as a white solid (0.093 g, 89.5 % yield). 1H NMR (600 MHz, DMSO-d6) δ 7.72 (dd, J = 9.2, 3.5 Hz, 4H), 7.12 (dd, J = 16.0, 8.9 Hz, 4H), 4.92 (d, J = 2.3 Hz, 2H), 4.19 (t, J = 5.7 Hz, 2H), 3.69 (t, J = 7.4 Hz, 2H), 3.65 (t, J = 2.3 Hz, 1H), 2.29 (quin, J = 6.2 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 193.1, 161.7, 160.4, 131.9, 131.7, 130.7, 130.0, 114.6, 114.3, 78.8, 68.4, 65.6, 55.7, 31.7, 31.1; positive ion HRESIMS [M+Na]+ 395.0263 (calcd. for C19H17O3BrNa, 395.0259).      Br BrDMF, K2CO3rt, 18hrs O O OBrHO O O 6.206.22  272   Figure 6.20 1H and 13C NMR spectra of 6.20 in DMSO-d6 at 600 and 150 MHz.   273 Preparation of 6.23:  To a stirred solution of latonduine A (6.9) (0.8 mg, 0.003 mmol) and K2CO3 (4.0 mg, 0.03 mmol) in a 5mL conical flask containing DMF (0.5 mL) at room temperature under Ar, was added a solution of 6.20 (0.01 g, 0.03 mmol) in DMF (0.5 mL).  This was allowed to stir at room temperature overnight, after which time, the reaction mixture was diluted in 30 mL of H2O and extracted with EtOAc (3 X 15 mL).  The EtOAc was concentrated under reduced pressure and separated directly using reversed phase C8 HPLC (7:10 ACN/H2O, retention time 9 min), yielding 3-amino-9, 10-dibromo-8-(3-(4-(4-(prop-2-yn-1-yloxy) benzoyl) phenoxy) propyl)-5, 6-dihydropyrimido [4, 5-c]pyrrolo[3, 2-e]azepin-7(8H)-one (6.23) as a pure white solid in moderate yield (1.19 mg, 60%). 1H NMR (600 MHz, DMSO-d6) δ 8.71 (s, 1H), 8.33 (t, J = 5.6 Hz), 7.70 (m, 4H), 7.13 (d, J = 9.2 Hz, 2H), 7.03 (d, J = 8.7 Hz, 2H), 6.95 (s, 2 N-H), 4.93 (d, J = 2.6 Hz, 2H), 4.85 (bm, 1H), 4.37 (bm, 1H), 4.07 (bm, 1H), 3.88 (bm, 1H), 3.65 (t, J = 2.6 Hz, 1H), 3.57 (bm, 1H), 2.22 (bm, 2H); 13C NMR (150 MHz, DMSO-d6) δ 193.1, 164.6, 162.2, 161.7, 161.4, 160.4, 156.7, 131.8 (2 X C), 131.7 (2 X C), 130.7, 129.9, 124.9, 121.0, 114.6 (2 X C), 114.2 (2 X C), 112.9, 112.8, 96.7, 78.8, 65.0, 55.7, 45.8, 45.1, 40.0, 29.6; positive ion HRESIMS [M+Na]+ at m/z 686.0020 (calcd. for C29H23N5O4NaBr2, 686.0014).   NH NHBr Br O NN NH2 K 2 CO 3 ,  DMFrt, 1 dayO O OBr + NHN O BrBrNNH2 N O O O6.236.20 6.9  274   Figure 6.21 1H and 13C NMR spectra of 6.23 in DMSO-d6 at 600 and 150 MHz.   275 Preparation of 6.27:  To a dried round bottom flask under Ar atmosphere was added 6.11 (1.54 g, 5.03 mmol) and 30 mL of dry THF.  To this LiBH4 (0.180 g, 7.47 mmol) was added and then 5 mL of 2M BACH-EI (10.0 mmol) was added to the solution dropwise over 10 min.  The reaction was stirred overnight and then quenched using 2-3 mL of 1M NaOH and stirred for 30 min.  After this time, 1.5 mL of a 30% H2O2 solution was added and stirred for an additional 30 min.  The reaction mixture was then concentrated under reduced pressure, dissolved in 300 mL of H2O, and extracted (3 X 100 mL) with EtOAc. Flash silica gel chromatography (eluting with 5% MeOH/EtOAc) gave 6.12 (0.684 g, 42.3%) and 2, 3-dibromo-4-hydroxy-4, 5, 6, 7-tetrahydropyrrolo[2, 3-c]azepin-8(1H)-one (6.27), as a white solid (0.193 g, 12%). Characterization of 6.27: 1H NMR (600 MHz, DMSO-d6) δ 12.47 (s, 1H), 7.95 (bd, J = 6.8 Hz, 1H), 5.07 (d, J = 5.8 Hz, 1H), 4.65 (m, 1H), 3.46 (t, J = 13.5 Hz, 1H), 3.04 (quin, J = 7.2 Hz, 1H), 2.05 (m, 1H), 1.79 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 161.3, 127.3, 124.0, 105.9, 101.2, 63.4, 35.5, 34.7; negative ion HRESIMS [M-H]- m/z 320.8875 (calcd. for C8H7N2O2Br2, 320.8875).     NH NHBrBr O NH NHBrBr O OHi. BACH-EI, LiBH4,THFii. NaOH, H2O2 NH NHBrBr OHO+6.11 6.12 6.27  276   Figure 6.22 1H and 13C NMR spectra of 6.27 in DMSO-d6 at 600 and 150 MHz.   277 Preparation of 6.29 and 6.30:  6.28 (0.2 g, 1.48 mmol) and NaN3 (2.96 mmol) were stirred together in a round bottom flask (25 mL) at 0 °C for 2 hours in conc. HCl (5 mL). The reaction mixture was then warmed to room temperature and stirred for another hour.  The reaction mixture was then poured over ice (100 mL), and treated with a 10% ceric ammonium nitrate solution (CAN). Once bubbling had ceased and the ice had melted, the reaction mixture was extracted with EtOAc (3 X 20 mL). The organic layers were combined and washed with a saturated brine solution, dried over MgSO4, filtered, and concentrated under reduced pressure. The mixture was then purified using silica gel flash chromatography (2.5 X 7 cm, eluting with a step gradient, 1:1, 2:1, 3:1, 4:1, and 1:0 EtOAc/Hex), giving: 6.28 (0.07 g), 5, 6, 7, 8-tetrahydropyrrolo[3, 2-c]tetrazolo[1, 5-a]azepine (6.30) 25.2 mg (15 %, BRSM), and 5, 6, 7, 8-tetrahydropyrrolo[3, 2-c]azepin-4(1H)-one (6.29) (22.3 mg, 13.5 %, BRSM).  5, 6, 7, 8-tetrahydropyrrolo[3, 2-c]tetrazolo[1, 5-a]azepine. 1H NMR (600 MHz, DMSO-d6) δ 11.29 (bs, 1H), 6.82 (t, J = 2.8 Hz, 1H), 6.53 (d, J = 2.8 Hz, 1H), 4.63 (m, 2H), 3.06 (dd, J = 7.4, 6.0 Hz, 2H), 2.14 (pent, J = 6.3 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 150.6, 133.2, 118.6, 107.2, 103.4, 48.8, 26.8, 22.0; positive ion HRESIMS [M+H]+ m/z 176.0937 (calcd. for C8H10N5, 176.0936). N a N 3  (2 eq)HClNH O NH NNH HN N N NO +6.28 6.29 6.30NH NN N N6.30  278    Figure 6.23 1H and 13C NMR spectra of 6.30 in DMSO-d6 at 600 and 150 MHz.    279 Characterization of 6.29:  5, 6, 7, 8-tetrahydropyrrolo[3, 2-c]azepin-4(1H)-one. 1H NMR (600 MHz, DMSO-d6) δ 10.92 (bs, 1H), 7.27 (bt, J = 4.2 Hz, 1H), 6.61 (t, J = 2.6 Hz, 1H), 6.29 (1H, J = 2.6 Hz, 1H), 3.14 (m, 2H), 2.86 (t, J = 7.3 Hz, 2H), 1.88 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 167.0, 133.0, 116.7, 115.4, 110.3, 41.3, 27.6, 26.3; positive ion HRESIMS [M+Na]+ m/z 173.0693 (calcd. for C8H10N2ONa, 173.0691).               NH HNO 6.29  280   Figure 6.24 1H and 13C NMR spectra of 6.29 in DMSO-d6 at 600 and 150 MHz.   281 Preparation of 6.31:  In a round bottom flask, a solution of 6.29 (0.02 g, 0.13 mmol) in DCM (5 mL) at 0 °C was stirred, while N-bromosuccinimide (0.048 g, 0.27 mmol)(NBS) was added in 3 small additions.  Once all of the NBS was added (~5 minutes), the reaction mixture was allowed to warm to room temperature and stir for 16 hours.  Once all starting material had been consumed (TLC), 1–2 mL of a saturated NaHCO3 solution was added and the reaction mixture was concentrated under reduced pressure.  The crude was then dissolved in 30 mL of H2O, and partitioned with EtOAc (3 X 10 mL).  The organic layers were combined, dried with MgSO4, filtered, and concentrated to give a crude solid.  The crude mixture was purified using silica gel chromatography (eluting with EtOAc) to give 2, 3-dibromo-5, 6, 7, 8-tetrahydropyrrolo[3, 2-c]azepin-4(1H)-one (6.31) as a white solid (0.033 g, 83%).  1H NMR (600 MHz, DMSO-d6) δ 12.09 (bs, 1H), 7.53 (s, 1H), 3.06 (m, 2H), 2.82 (t, J = 9.4 Hz, 2H), 1.86 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 135.3, 114.1, 100.5, 97.9, 40.1, 27.4, 25.8; positive ion HRESIMS [M+H]+ m/z 306.9081 (calcd. for C8H8N2OBr2, 306.9082).      HN NHO HN NHOBr BrNBS, DCM 6.31  282   Figure 6.25 1H and 13C NMR spectra of 6.31 in DMSO-d6 at 600 and 150 MHz.   283 Preparation of 6.30:  To a stirred solution of 6.30 (0.02 g, 0.11 mmol) in a round bottom flask with DCM (5 mL) at 0 °C was slowly added NBS (0.041 g, 0.22 mmol) in 3 small additions.  Once all of the NBS was added (~5 min) the reaction mixture was allowed too warm to room temperature and stir for 16 hours.  Once all starting material had been consumed, 1–2 mL of a saturated NaHCO3 solution was added and the reaction mixture was concentrated under reduced pressure.  The crude was then dissolved in 30 mL of H2O, and partitioned with EtOAc (3 X 10 mL).  The organic layers were combined, dried with MgSO4, filtered, and concentrated to give a crude solid.  Silica gel flash chromatography (6 X 2.5 cm, eluting with EtOAc/Hex (1:3)) afforded 9, 10-dibromo-5, 6, 7, 8-tetrahydropyrrolo[3, 2-c]tetrazolo[1, 5-a]azepine (6.32), as a white solid (0.022 g, 60 %). 1H NMR (600 MHz, DMSO-d6) δ 12.51 (bs, 1H), 4.65 (m, 2H), 3.07 (t, J = 6.2 Hz, 2H), 2.11 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 148.3, 135.6, 104.1, 102.4, 96.3, 48.6, 29.5, 27.5, 21.9; positive ion HRESIMS [M+H]+ m/z 331.9149 (calcd. for C8H8N5Br2, 331.9146).      HN NNN N NBS, DCM HN NN NNBr Br 6.32  284   Figure 6.26 1H and 13C NMR spectra of 6.32 in DMSO-d6 at 600 and 150 MHz.    285 Preparation of 6.33:  To a round bottom flask was added a solution of phthalimide (1 g, 6.8 mmol), Cs2CO3 (2.215 g, 6.8 mmol) and DMF (19 mL) under Ar atmosphere and stirred for 1 hour at room temperature.  The mixture was then cooled to 0 °C and bromoethyltrimethylsilane (0.903 mL, 5.7 mmol) was added dropwise over 5 min.  The reaction mixture was then heated to 80 °C for 18 hours.  The reaction was then poured into (~200 mL) H2O and extracted with EtOAc (2 X 75 mL).  The organic layers were combined, washed with brine, dried over MgSO4, filtered, and concentrated under reduced pressure to give the crude product.  Purification was accomplished with flash silica gel chromatography (eluting with EtOAc/Hex (1:1 then 2:1)) to give 2-(2-(trimethylsilyl)ethyl)isoindoline-1,3-dione (6.33), as a white solid (0.210 g, 15%). 1H NMR (600 MHz, CDCl3) δ 7.81 (dd, J = 5.3, 2.8 Hz, 2H), 7.67 (dd, J = 5.3, 2.8 Hz, 2H), 3.70 (m, 2H), 1.00 (m, 2H), 0.05 (s, 9H); 13C NMR (150 MHz, CDCl3) δ 169.5, 134.0, 132.5, 123.3, 34.7, 17.3, -1.6; positive ion LRESIMS [M+Na]+ m/z 270.0 (calcd. for C13H17NO2SiNa, 270.0).244 Preparation of 6.34:  To a Pyrex® test tube was added 6.33 (0.05 g, 0.2 mmol), and 22.4 mL of a 35%(v/v) H2O/ACN solution.  The tube was sealed using a rubber septa and degassed using N2.  The tube was then irradiated with UV light for 2 hours in an ice bath.  The reaction mixture was then concentrated under reduced pressure and partitioned between H2O (30 mL) and EtOAc (3 X 10 mL).  The organic layers were combined, dried with MgSO4, filtered and concentrated.  Subsequent silica gel chromatography (eluting with 10% MeOH/EtOAc) gave starting material (0.017 g) and 3, 4-NOO SiNHOO SiBrD M F, Cs 2CO 3 6.33NOO Si h ! NHOOACN/H 2O6.33 6.34  286 dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.34) (13.5 mg, 58% BRSM). It should be noted that the literature procedure performed the UV irradiation in a Vycor reaction vessel. 245 1H NMR (600 MHz, DMSO-d6) δ 7.85 (d, J = 8.0 Hz, 1H), 7.71 (td, J = 7.1, 1.4 Hz, 1H), 7.65 (m, 2H), 3.56 (q, J = 6.7 Hz, 2H), 2.96 (dd, J = 6.7, 4.8 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 203.4, 170.5, 137.5, 134.2, 133.2, 132.3, 130.8, 128.9, 47.1, 37.7; positive ion TOFHRESIMS, [M+H]+ m/z 176.0708 (calcd. for C10H10NO2, 176.0712).245 Preparation of 6.35:  To a stirred solution of 6.34 (0.01 g, 0.06 mmol) and MeOH (1.5 mL) in a round bottom flask under Ar was added NaBH4 (0.002 g, 0.06 mmol) in one portion. After 2 hours the reaction was stopped by the addition of 1 mL H2O and concentrated under a stream of nitrogen. Crude product purified using silica gel chromatography (eluting with MeOH/EtOAc (1:9)) to give 5-hydroxy-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.35) as a white solid (0.009 g, 89%).1H NMR (600 MHz, DMSO-d6) δ 8.00 (bt, J = 5.7 Hz, 1H), 7.26 (d, J = 7.6 Hz, 1H), 7.48 (td, J = 7.2, 1.3 Hz, 1H), 7.44 (dd, J = 7.5, 1.0 Hz, 1H), 7.32 (td, J = 7.6 Hz, 1H), 5.42 (d, J = 4.8 Hz, 1H), 4.77 (m, 1H), 2.98 (m, 1H), 2.71 (m, 1H), 2.34 (m, 1H), 1.55 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 171.2, 141.9, 133.1, 130.4, 127.6, 126.9, 123.9, 68.3, 39.3, 37.6; positive ion TOFHRESIMS [M+H]+ m/z 178.0870 (calcd. for C10H12NO2, 178.0868). NHOO NaBH4MeOH NHOHO6.34 6.35  287   Figure 6.27 1H and 13C NMR spectra of 6.35 in DMSO-d6 at 600 and 150 MHz.     288 Preparation of 6.36:  A solution of 4, 5-diclorophthalimide (1 g, 4.6 mmol) and Cs2CO3 (1.49 g, 4.6 mmol) under Ar atmosphere was stirred for 1 hour at room temperature in a round bottom flask.  The mixture was then cooled to 0 °C and bromoethyltrimethylsilane (0.61 mL, 3.8 mmol) was added dropwise over 5 minutes.  The reaction mixture was then heated to 80 °C for 18 hours.  After this the reaction was poured into (~200 mL) H2O and extracted with EtOAc (2 X 75 mL).  The organic layers were combined, washed with brine, dried over MgSO4, filtered, and concentrated under reduced pressure to give the crude product. The crude was purified via silica gel chromatography (eluting with EtOAc/Hex (3:20)) to give 5, 6-dichloro-2-(2-(trimethylsilyl)ethyl)isoindoline-1, 3-dione (6.36), as an off-white solid (0.253 g, 21.1%) 1H NMR (600 MHz, CDCl3) δ 7.89 (s, 2H), 3.68 (m, 2H), 0.97 (m, 2H), 0.05 (s, 9H); 13C NMR (150 MHz, CDCl3) δ 166.4, 138.9, 131.7, 125.4, 35.3, 17.2, -1.5; positive ion LRESIMS [M+H]+ m/z 316.2 (calcd. for C13H16NO2SiCl, 316.0).        NOO SiClClNHOO SiBrClCl DMF, Cs 2 CO3 6.36  289   Figure 6.28 1H and 13C NMR spectra of 6.36 in CDCl3 at 600 and 150 MHz.    290 Preparation of 6.37:  To a Pyrex® test tube was added 6.36 (0.221g, 0.702 mmol) in 25 mL of (1:4) H2O/ACN (ACN content increased for 6.36 solubility).  The tube was sealed using a rubber septa and degassed using N2.  The tube was then irradiated with UV light for 2 hours in an ice bath.  The reaction mixture was then concentrated under reduced pressure and partitioned between H2O (30 mL) and EtOAc (3 X 10 mL).  The organic layers were combined, dried with MgSO4, filtered and concentrated.  The crude product was purified using silica gel chromatography (eluting with 5% MeOH/EtOAc) to give 7, 8-dichloro-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.37) as a white solid (0.063 g, 37%). It should be noted that the literature procedure performed the UV irradiation in a Vycor reaction vessel. 245 1H NMR (600 MHz, DMSO-d6) δ 8.74 (bt, J = 5.9 Hz, 1H), 7.94 (s, 1H), 7.77 (s, 1H), 3.37 (q, J = 5.2 Hz, 2H), 2.92 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 200.6, 167.0, 135.6, 135.4, 134.3, 131.8, 129.9, 45.7, 36.1; positive ion HRESIMS [M+H]+ 243.9927 (calcd. for C10H8NO2Cl2, 243.9932).      NOO Si h! NHOOClCl ClClACN/H 2O 6.376.36  291   Figure 6.29 1H and 13C NMR spectra of 6.37 in DMSO-d6 at 600 and 150 MHz.   292 Preparation of 6.38:  A solution of 6.37 (0.02 g, 0.08 mmol), NaBH4 (0.003 g, 0.08 mmol), in 2 mL absolute MeOH was assembled in a small glass vial under Ar atmosphere. After 2 hours the reaction was stopped by the addition of 1 mL H2O and concentrated under a stream of nitrogen. The product was purified using silica gel chromatography (eluting with MeOH/EtOAc (1:9)) to give 7,8-dichloro-5-hydroxy-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.38), as a light tan solid (0.018 g, 90%). 1H NMR (600 MHz, DMSO-d6) δ 8.26 (bt, J = 4.9 Hz, 1H), 7.70 (s, 1H), 7.65 (s, 1H), 5.75 (bs, 1H), 4.78 (t, J = 8.6 Hz, 1H), 3.04 (m, 1H), 2.80 (m, 1H), 2.39 (m, 1H), 1.59 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 168.8, 142.9, 133.5, 133.1, 129.8, 126.3, 67.6, 38.7, 37.4; positive ion HRESIMS [M+H]+ m/z 246.0093 (calcd. for C10H10NO2Cl2, 246.0089).         NHOO NaBH4MeOH NHOHOClCl ClCl6.37 6.38  293   Figure 6.30 1H and 13C NMR spectra of 6.38 in DMSO-d6 at 600 and 150 MHz.   294 General Procedure for reactions of the following type:  Following a procedure from Suau et al.,243 to a Pyrex® Erlenmeyer flask containing 140 mL of ACN with 25 mL of H2O, was added phthalimide (1 g, 6.8 mmol),  and NaOH (1M, 2 mL).  The reaction vessel was capped with a rubber septum, and styrene or substituted styrene derivative (2 eq) was added and the reaction mixture was degassed using a stream of N2.  The reaction was subsequently irradiated for 2 hours in an ice bath after which point the reaction was acidified using 1M HCl.  The mixture was then concentrated under reduced pressure and (~50 mL) H2O was added. The mixture was then extracted with EtOAc (3 X 20 mL).  The organic layers were combined, washed with brine, dried over MgSO4, filtered, and concentrated under reduced pressure. The mixture was purified using flash silica gel chromatography as described below. Preparation of 6.39:  See above general procedure: Product purified eluting with EtOAc/hexanes (1:1) to give 4-phenyl-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.39) as white solid (0.370 g, 22%) 1H NMR (600 MHz, DMSO-d6) δ 8.58 (bt, J = 5.4 Hz, 1H), 7.82 (d, J = 6.8 Hz, 1H), 7.76 (t, J = 6.8 Hz, 1H), 7.70 (t, J = 6.8 Hz, 1H), 7.47 (d, J = 6.8 Hz, 1H), 7.34 (t, J = 7.1 Hz, 2H), 7.29 (m, 1H), 7.18 (d, J = 7.1 Hz, 2H), 4.21 (dd, J = 9.0, 3.9 Hz, 1H), 3.65 (m, 1H), 3.53 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 204.9, 169.2, 138.2, 136.7, 132.5, 131.8, 129.3, 128.7, 128.3, 127.8, 127.3, 125.5, 61.2, 43.2; positive ion TOFHRESIMS [M+H]+ m/z 252.120 (calcd. for C16H14NO2, 252.1025).243   NHOO Ar+ h ! NHOO ArACN/H 2ONHOO + h ! NHOOA CN/H 2 O 6.39  295 Preparation of 6.40:  Using general procedure (p. 294) substituting 2-vinyl pyridine (2.5 eq, 1.44 mL) for styrene. Purified using flash silica gel chromatography (eluting with a step gradient Hex/EtOAc (3:4), EtOAc, and 5 % MeOH/EtOAc) to give 4-(pyridin-2-yl)-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.40) (0.087 g, 5.1 %) as a yellow solid. Thought to exist as the enol tautomer. 1H NMR (600 MHz, DMSO-d6) δ 8.58 (t, J = 6.8 Hz, 1H), 8.50 (d, J = 4.8 Hz, 1H), 7.97 (td, J = 8.7, 1.0 Hz, 1H), 7.86 (d, J = 7.7 Hz, 1H), 7.82 (d, J = 7.7 Hz, 1H), 7.65 (m, 2H), 7.57 (t, J = 7.7 Hz, 1H), 7.26 (dd, J = 7.7, 5.8 Hz, 1H), 4.05 (bs, 1H), 3.61 (bs, 1H); 13C NMR (125 MHz, DMSO-d6) δ 168.7, 163.8, 155.8, 143.9, 138.8, 134.9, 133.9, 130.6, 129.5, 129.4, 127.2, 119.4, 118.6, 107.2, 36.9; positive ion TOFHRESIMS [M+H]+ m/z 253.0975 (calcd. for C15H13N2O2, 253.0977).      NHOO + h! NHOOACN/H 2 ON N6.40  296   Figure 6.31 1H and 13C NMR spectra of 6.40 in DMSO-d6 at 600 and 150 MHz.    297 Preparation of 6.39:  Using general procedure (p. 294) substituting 2-vinyl pyridine (0.96 mL, 2 eq) for styrene and 3, 4-chlorophthalimide (1 g, 4.6 mmol) for phthalimide.  Purified using flash silica gel chromatography (eluting with Hex/EtOAc (3:4), EtOAc, and 5 % MeOH/EtOAc) to give 7, 8-dichloro-4-(pyridin-2-yl)-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.41) (0.032 g, 2.2 %) as a bright yellow solid. 1H NMR (600 MHz, DMSO-d6) δ 8.77 (t, J = 6.3 Hz, 1H), 8.49 (d, J = 4.7 Hz, 1H), 8.00 (s, 1H), 7.99 (obs, 1H), 7.96 (s, 1H), 7.65 (d, J = 8.1 Hz, 1H), 7.27 (dd, J = 7.3, 5.1 Hz, 1H), 3.89 (vbs, 2H); 13C NMR (150 MHz, DMSO-d6) δ 166.7, 163.6, 154.9, 142.9, 139.3, 135.8, 134.0, 133.6, 132.2, 131.3, 129.2, 119.3, 118.8, 106.8, 36.6; negative ion HRESIMS [M-H]- m/z 319.0035 (calcd. for C15H9N2O2Cl2, 319.0041).         NHOO + h ! NHOOACN/H 2ON NClCl ClCl 6.41  298   Figure 6.32 1H and 13C NMR spectra of 6.41 in DMSO-d6 at 600 and 150 MHz.    299 Preparation of 6.42:  Using general procedure (p. 294) substituting 3, 4-dimethoxy styrene (2.0 mL, 2 eq) for styrene.  Purified using flash silica gel chromatography (eluting with 5 % MeOH/EtOAc) to give 4-(3, 4-dimethoxyphenyl)-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.42) (0.017 g, 0.8 %) as a yellow solid. Thought to exist as a rapidly converting quinone methide/enol. 1H NMR (600 MHz, DMSO-d6) δ 8.53 (t, J = 5.6 Hz, 1H), 7.77 (d, J = 9.4 Hz, 1H), 7.71 (t, J = 7.5 Hz, 1H), 7.64 (t, J = 7.5 Hz, 1H), 7.30 (d, J = 7.5 Hz, 1H), 6.93 (d, J = 2.0 Hz, 1H), 6.86 (d, J = 8.7 Hz, 1H), 6.73 (dd, J = 8.4, 2.3 Hz, 1H), 6.52 (bs, 1H), 3.87 (dd, J = 14.9, 5.2 Hz, 1H), 3.69 (s, 3H), 3.66 (s, 3H), 3.22 (dd, J = 15.0, 6.5 Hz, 1H); 13C NMR (150 MHz, DMSO-d6) δ 205.9, 169.2, 148.4 (X 2), 137.0, 132.7, 132.2 (X 2), 131.5, 129.2, 127.7, 117.7, 111.3, 109.4, 83.9, 55.5, 55.3, 49.4; positive ion TOFHRESIMS [M+H]+ m/z 312.1242 (calcd. for C18H18NO4, 312.1236).       NHOO + h! NHOOACN/H2OOO O O+ 6.42  300   Figure 6.33 1H and 13C NMR spectra of 6.42 in DMSO-d6 at 600 and 150 MHz.    301 General transformation scheme for reduction of the following type:  To a stirred solution of azepinedione (1 eq) was added NaBH4 (1.2 eq) under Ar atmosphere in absolute MeOH (~2 mL).  This was stirred while monitoring by TLC with another equivalent of NaBH4 added every 2 hours. Once complete consumption of starting material had been achieved, H2O was added and the reaction mixture was concentrated.  The reaction was then partitioned between H2O (~15 mL) and EtOAc (3 X 7 mL).  The organic layers were combined, concentrated, and subjected to silica gel column chromatography as described below.   Preparation of 6.43:  Using general procedure (p. 301) above: 6.39 (0.1 g, 0.396 mmol) and NaBH4 (0.018 g, 0.47 mmol) were added to a round bottom flask in absolute MeOH.  Crude reaction mixture was purified using silica gel chromatography (eluting with EtOAc/hexanes (9:1)) to give 5-hydroxy-4-phenyl-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.43) as a white solid (0.089 g, 89%).1H NMR (600 MHz, DMSO-d6) δ 8.20 (bs, 1H), 8.54 (d, J = 6.5 Hz, 1H), 7.50 (t, J = 6.5 Hz, 1H), 7.38 (t, J = 6.5 Hz, 1H), 7.36 (obs, 1H), 7.14 (bs, 3H), 6.85 (bd, J = 3.3 Hz, 2H), 5.33 (bs, 1H), 5.15 (bd, J = 6.5 Hz, 1H), 3.64 (m, 1H), 3.13 (d, J = 10.2 Hz, 1H), 3.18 (m, 1H), 2.91 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 171.3, 140.5, 139.2, 133.0, 130.3, 129.0, 128.3, 127.7, 127.6, 127.1, 126.5, 125.3, 70.1, 53.1, 45.2; positive ion HRESIMS [M+H]+ m/z 254.1189 (calcd. for C16H16NO2, 254.1181). NHOO R 1 NaBH 4MeOH NHOHO R 1R 2R 2 R 2R 2NHOO NaBH4MeOH NHOHO 6.436.39  302   Figure 6.34 1H and 13C NMR spectra of 6.43 in DMSO-d6 at 600 and 150 MHz.   303 Preparation of 6.44:  Using general procedure (p. 301) 6.40 (0.01 g, 0.04 mmol), NaBH4 (1.8 mg, 0.05 mmol) and absolute MeOH were added to a small glass vial. The crude product was purified by flash chromatography (eluting with MeOH/EtOAc (1:9)) to give 5-hydroxy-4-(pyridin-2-yl)-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.44) as a white solid (0.009 g, 89 %). 1H NMR (600 MHz, DMSO-d6) δ 8.33 (d, J = 3.4 Hz, 1H), 8.15 (bt, J = 5.6 Hz, 1H), 7.64 (t, J = 6.8 Hz, 1H), 7.53 (d, J = 6.8 Hz, 1H), 7.46 (t, J = 6.8 Hz, 1H), 7.36 (t, J = 7.9 Hz, 1H), 7.28 (d, J = 6.8 Hz, 1H), 7.17 (m, 1H), 7.12 (d, J = 6.8 Hz, 1H), 5.52 (bs, 1H), 5.19 (d, J = 7.1 Hz, 1H), 3.81 (m, 1H), 3.21 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 171.2, 158.6, 148.1, 140.6, 135.6, 132.9, 130.0, 127.5, 126.9, 125.4, 124.6, 121.7, 70.2, 54.3, 43.3; positive ion HRESIMS [M+H]+ 255.1134 (calcd. for C15H15N2O2, 255.1134).         NaBH4MeOH N HOHO NNHOO N6.40 6.44  304   Figure 6.35 1H and 13C NMR spectra of 6.44 in DMSO-d6 at 600 and 150 MHz.    305 Preparation of 6.45:  Using general procedure (p. 301) 6.41 (0.013 g, 0.04 mmol), NaBH4 (1.8 mg, 0.048 mmol), and absolute MeOH were added to a small glass vial.  The crude product was purified using column chromatography (eluting with 5 % MeOH/EtOAc) to give 7, 8-dichloro-5-hydroxy-4-(pyridin-2-yl)-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.45) as a light yellow solid (7 mg, 54%). 1H NMR (600 MHz, MeOD-d4) δ 8.38 (d, J = 5.5 Hz, 1H), 7.76 (s, 1H), 7.70 (td, J = 7.5, 2.0 Hz, 1H), 7.49 (s, 1H), 7.24, (ddd, J = 7.5, 5.0, 1.0 Hz, 1H), 7.16 (d, J = 7.9 Hz, 1H), 5.28 (d, J = 8.3 Hz, 1H), 3.96 (m, 1H), 3.50 (dd, J = 14.0, 11.0 Hz, 1H), 3.39 (dd, J = 14.0, 6.5 Hz, 1H); 13C NMR (150 MHz, MeOD-d4) δ 170.8, 157.2, 147.6, 140.8, 135.9, 134.3, 131.7, 130.5, 129.3, 127.4, 124.3, 121.8, 69.6, 53.7, 42.9; positive ion HRESIMS [M+H]+ m/z 323.0352 (calcd. for C15H13N2O2Cl2, 323.0354).         NHOHO NClClNHOO NClCl NaBH 4MeOH6.41 6.45  306   Figure 6.36 1H and 13C NMR spectra of 6.45 in MeOD-d4 at 600 and 150 MHz.    307 Preparation of 6.46:  Using general procedure (p. 301) 6.42 (0.01 g, 0.03 mmol), NaBH4 (1.4 mg, 0.036 mmol), and absolute MeOH were added to a small glass vial.  The crude product was purified using column chromatography (eluting with MeOH/EtOAc (9:1)) to give 4-(3, 4-dimethoxyphenyl)-5-hydroxy-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.46) product as an off white solid (0.007 g, 75%).  1H NMR (600 MHz, acetone-d6) δ 7.67 (d, J = 7.8 Hz, 1H), 7.55 (m, 1H), 7.43 (m, 1H), 7.33 (bs, 1H), 6.78 (d, J = 8.8 Hz, 1H), 6.53 (dd, J = 8.3, 1.8 Hz, 1H), 6.45 (d, J = 2.0 Hz, 1H), 5.38 (dd, J = 8.2, 5.2 Hz, 1H), 4.07 (d, J = 5.6 Hz, 1H), 3.78 (m, 1H), 3.75 (s, 3H), 3.60 (s, 3H), 3.40 (m, 1H), 3.14 (m, 1H); 13C NMR (150 MHz, acetone-d6) δ 172.4, 149.8, 149.5, 141.5, 134.3, 131.8, 131.2, 129.1, 128.1, 126.5, 122.7, 113.8, 112.4, 71.8, 54.3, 46.7; positive ion HRESIMS [M+Na]+ m/z 336.1217 (calcd. for C18H19NO4Na, 336.1212).        NHOO O O NHOHO O ONaBH4MeOH6.42 6.46  308   Figure 6.37 1H and 13C NMR spectra of 6.46 in acetone-d6 at 600 and 150 MHz.   309 Preparation of 6.47:  Following the procedure from Suau et al.,243 a Pyrex® Erlenmeyer flask containing 140 mL of ACN with 25 mL of H2O, phthalimide (1 g, 6.8 mmol),  and NaOH (1M, 2 mL) was capped using a rubber septum, and allyl trimethylsilyl ether (2.29 mL, 13.6 mmol) was added. The reaction mixture was then degassed using a stream of N2.  The reaction was subsequently irradiated for 2 hours in an ice bath, after which point, the reaction was acidified using 1M HCl.  The mixture was then concentrated under reduced pressure and H2O (~50 mL) was added and the mixture was extracted with EtOAc (3 X 20 mL).  The organic layers were combined, dried over MgSO4, filtered, and concentrated under reduced pressure. The mixture was purified using flash silica gel chromatography (eluting with Hex/EtOAc (3:4), EtOAc, and 5 % MeOH/EtOAc) to give 3-(hydroxymethyl)-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione (6.47) (0.011 g, 0.8 %) as a white solid. 1H NMR (600 MHz, DMSO-d6) δ 8.17 (d, J = 5.4 Hz, 1H), 7.77 (dd, J = 7.7, 1.0 Hz, 1H), 7.71 (td, J = 7.6, 1.0 Hz 2H), 7.66 (td, J = 7.6, 1.0 Hz, 1H), 7.56 (bd, 7.7 Hz, 1H), 4.88 (t, J = 5.8 Hz, 1H), 3.72 (m, 1H), 3.51 (m, 2H), 2.94 (dd, J = 18.8, 2.7 Hz, 2H), 2.79 (dd, J = 18.8, 11.6 Hz, 1H); 13C NMR (150 MHz, DMSO-d6) δ 202.5, 168.5, 136.4, 133.2, 132.4, 131.4, 129.4, 127.5, 61.7, 50.6, 48.8; positive ion TOFHRESIMS [M+Na]+ m/z 228.0644 (calcd. for C11H11NO3Na, 228.0637).    N HOO h !N aOH, ACN NHOOOTMS OH+ 6.47  310   Figure 6.38 1H and 13C NMR spectra of 6.47 in DMSO-d6 at 600 and 150 MHz.   311 Preparation of 6.48 c and 6.48 t:  To a small vial was added 6.47 (0.01 g, 0.076 mmol) under Ar atmosphere in absolute MeOH (1.5 mL).  Then NaBH4 (0.003 g, 0.076 mmol) was added in and the reaction was stirred for 2 hours at which point another aliquot of NaBH4 was added.  After another 2 hour period, the reaction was quenched using 0.5 mL of a saturated ammonium chloride solution, concentrated under a stream of N2, and partitioned between H2O (10 mL) and EtOAc (3 X 3 mL) to give crude alcohol 6.48 in the organic layer. Purification of the crude product using C18 reversed phase HPLC (eluting with 15 % ACN/H2O) gave the cis (6.48 c)(2.5 mg, rt = 11 min) and trans (6.48 t)(5.4 mg, rt = 10 min) diastereomers as individual peaks. Characterization of 6.48 t:  (3S*, 5R*)-5-hydroxy-3-(hydroxymethyl)-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one: 1H NMR (600 MHz, DMSO-d6) δ 7.66 (d, J = 4.6 Hz, 1H), 7.60 (d, J = 7.2 Hz, 1H), 7.51 (td, J = 7.5, 1.2 Hz, 1H), 7.47 (dd, J = 7.7, 1.2 Hz, 1H), 7.35 (t, J = 7.5 Hz, 1H), 5.50 (d, J = 6.0 Hz, 1H), 4.79 (m, 1H), 4.68 (t, J = 5.3 Hz, 1H), 3.42 (m, 2H), 2.96 (hept, J = 5.8 Hz, 1H), 2.18 (m, 1H), 1.58 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 170.4, 142.3, 132.8, 130.5, 127.3, 126.8, 123.2, 67.6, 62.3, 52.3, 42.4; positive ion TOFHRESIMS [M+Na]+ m/z 230.0795 (calcd. for C11H13NO3Na, 230.0793).  NHOO NaBH4MeOH NHOHO OHNHOHO OHOH +6.47 6.48 t 6.48 cNHOHO OH6.48 t  312   Figure 6.39 1H and 13C NMR spectra of 6.48 t in DMSO-d6 at 600 and 150 MHz.   313  Figure 6.40 tROESY NMR spectrum of 6.48 t in DMSO-d6 at 600 MHz Characterization of 6.48 c:  (3S*, 5S*)-5-hydroxy-3-(hydroxymethyl)-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one: 1H NMR (600 MHz, DMSO-d6) δ 7.50 (dd, J = 7.4, 1.2 Hz, 1H), 7.40 (d, J = 6.2 Hz, 1H), 7.39 (td, J = 7.9, 1.2 Hz, 1H), 7.35 (td, J = 7.9, 1.2 Hz, 1H), 7.26 (d, J = 7.3 Hz, 1H), 5.21 (bs, 1H), 4.78 (m, 1H), 4.63 (t, J = 5.6 Hz, 1H), 3.28 (obs, 2H), 3.03 (hept, J = 5.1 Hz, 1H), 2.09 (m, 1H), 1.88 (m, 1H); 13C NMR (150 MHz, DMSO-d6) δ 170.7, 142.3, 132.8, 130.0, 127.5, 126.8, 123.3, 70.2, 62.5, 51.8, 41.3; positive ion TOFHRESIMS [M+Na]+ m/z 230.0795 (calcd. for C11H13NO3Na, 230.0793).  NHOHO OH6.48 c  314   Figure 6.41 1H and 13C NMR spectra of 6.48 c in DMSO-d6 at 600 and 150 MHz.   315  Figure 6.42 tROESY NMR spectra of 6.48 c in DMSO-d6 at 600 MHz General Procedure for Schmidt reaction:  A vigorously stirring flask containing tetralone (0.5 g, 1 eq) in concentrated HCl (0.3 M) was cooled to 0 °C.  To this mixture was added in several small aliquots NaN3 (2 eq) over 5 minutes.  The mixture was left to stir open to air for 2 hours at 0 °C and then allowed to slowly warm to room temp and stir for another 2 hours.  The reaction mixture was then poured over ice and treated with 2 mL of a 10% solution of ceric ammonium nitrate (CAN).  Once bubbling had ceased, the reaction mixture was neutralized with a saturated K2CO3 solution and the ice was allowed to melt.  The reaction mixture was then extracted with (3 X ~10 mL) EtOAc, the organic layers were combined dried over MgSO4, filtered, and reduced via rotoevaporation to give crude products and some remaining starting materials. Purification of the tetrazole, and two regio-isomeric azepines was accomplished using flash silica gel chromatography as described.       316 Preparation of 6.49A, B, and C:  Using the general procedure above (p. 315) to give the crude as described.  Purification using flash silica gel chromatography (eluting with stepwise gradient using 1:3, 1:1, 3:1, and 1:0 (EtOAc/Hex) (200 mL) (4 X 12 cm column)) gave purified starting material (0.134 g), 6, 7-dihydro-5H-benzo[c]tetrazolo[1, 5-a]azepine (6.49A) (0.153 g, 32.6%, BRSM), 4, 5-dihydro-1H-benzo[b]azepin-2(3H)-one (6.49B) (0.066 g, 16.4 %, BRSM), and 2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one (6.49C) (0.089 g, 22.1 %, BRSM).   6, 7-dihydro-5H-benzo[c]tetrazolo[1, 5-a]azepine. 1H NMR (600 MHz, acetone-d6) δ 8.25 (d, J = 7.9 Hz, 1H), 7.49 (td, J = 7.5, 1.2 Hz, 1H), 7.43 (t, J = 7.5, 1H), 7.40 (d, J = 7.9, 1H), 4.70 (dd, J = 8.2, 6.3 Hz, 2H), 3.09 (m, 2H), 2.42 (pent, J = 5.0 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 154.9, 141.6, 132.2, 131.3, 130.6, 127.9, 124.4, 50.0, 33.6, 26.7; positive ion LRESIMS [M+H]+ m/z 187.0 (calcd. for C10H11N4, 187.1).247  4, 5-dihydro-1H-benzo[b]azepin-2(3H)-one. 1H NMR (600 MHz, acetone-d6) δ 8.57 (bs, 1H), 7.26 (d, J = 7.5 Hz, 1H), 7.23 (t, J = 7.5 Hz, 1H), 7.10 (t, J = 7.6 Hz, 1H), 7.04 (d, J = 7.6 Hz, 1H), 2.77 (t, J = 7.6 Hz, 2H), 2.22 (t, J = 7.4 Hz, 2H), 2.16 (pent, J = 7.3 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 173.8, 140.1, 135.1, 130.6, 128.2, 125.8, 122.5, 33.6, 31.1, 29.1; positive ion LRESIMS [M+H]+ m/z 162.2 (calcd. for C10H12NO, 162.1).247 NHOHN ONN NNO NaN 3 (2 eq)[HCl] + +6.49A 6.49B 6.49CNN NN6.49A HN O6.49B  317   2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one. 1H NMR (600 MHz, acetone-d6) δ 8.00 (bs, 1H), 7.48 (td, J = 7.4, 1.5 Hz, 1H), 7.41 (t, J = 7.4, 1H), 7.32 (t, J = 7.4 Hz, 1H), 7.25 (dd, J = 7.2, 1.5 Hz, 1H), 2.89 (q, J = 6.7 Hz, 2H), 2.72 (m, 2H), 1.86 (m, 2H); positive ion LRESIMS [M+H]+ m/z 162.1 (calcd. for C10H12NO, 162.1).247 Preparation of 6.50A and 6.50C:   Using general procedure (p. 315) with 6-amino-3, 4-dihydro-1(2H)-naphthalenone (0.5 g, 3.1 mmol). The crude reaction mixture was obtained as described above.  Purification of the crude using flash silica gel chromatography (eluting with 1:3, 1:1, 4:1 and 1:0 (EtOAc/Hex) (200 mL each) (4 X 10 cm column) gave purified starting material (0.291 g), 6.50A (0.152 g, 51.3 % BRSM), and 6.50C (0.062 g, 23.8 % BRSM). Characterization of 6.50A:  6, 7-dihydro-5H-benzo[c]tetrazolo[1, 5-a]azepin-9-amine: 1H NMR (600 MHz, acetone-d6) δ 8.01 (d, J = 8.8 Hz, 1H), 6.68 (dd, J = 8.5, 2.2 Hz, 1H), 6.61 (d, J = 1.8 Hz, 1H), 5.26 (bs, 2H), 4.61 (t, J = 6.3 Hz, 2H), 2.95 (m, 2H), 2.32 (m, 2H); 13C NMR (150 MHz, acetone-d6) δ 155.2, 152.2, 142.7, 132.2, 115.4, 113.3, 111.7, 50.0, 34.3, 26.1; positive ion TOFHRESIMS [M+H]+ m/z 202.1091 (calcd. for C10H12N5, 202.1093).   NHO6.49CNHONN NNO NaN 3 (2 eq)[HCl] +H2N H2N H2N6.50A 6.50CNN NNH2 N 6.50A  318   Figure 6.43 1H and 13C NMR spectra of 6.50A in acetone-d6 at 600 and 150 MHz.   319 Characterization of 6.50C:  7-amino-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one: 1H NMR (600 MHz, acetone-d6) δ 7.34 (d, J = 6.5 Hz, 1H), 6.83 (bs, 1H), 6.56 (dd, J = 8.2, 2.3 Hz, 1H), 6.46 (s, 1H), 4.99 (bs, 2H), 3.06 (q, J = 5.3 Hz, 2H), 2.67 (t, J = 7.4 Hz, 2H), 1.89 (pent, J = 7.4 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 173.8, 151.9, 140.8, 131.1, 124.9, 114.3, 112.6, 40.0, 31.5, 31.1; positive ion TOFHRESIMS [M+H]+ m/z 177.1023 (calcd. for C10H13N2O, 177.1028).             NHOH2 N 6.50C  320   Figure 6.44 1H and 13C NMR spectra of 6.50C in acetone-d6 at 600 and 150 MHz.   321 Preparation of 6.51B and 6.51C:  Using general methodology (p. 315) with 5-hydroxy-1-tetralone (0.5 g, 3.1 mmol). Purification of the crude reaction mixture by flash silica gel chromatography (eluting with 1:3, 1:1, 4:1 and 1:0 EtOAc/Hex, 200 mL each, 4 X 10 cm column) gave purified starting material (0.216 g), 6.51B (0.114 g, 36.8 % BRSM), and 6.51C (0.078 g, 22 % BRSM). Characterization of 6.51B:  6-hydroxy-4, 5-dihydro-1H-benzo[b]azepin-2(3H)-one: 1H NMR (600 MHz, DMSO-d6) δ 9.44 (s, 1H), δ 9.41 (s, 1H), 6.96 (t, J = 7.9 Hz, 1H), 6.61 (d, J = 7.9 Hz, 1H), 6.43 (d, J = 7.9 Hz, 1H), 2.66 (t, J = 7.4 Hz, 2H), 2.12 (t, J = 7.4 Hz, 2H), 2.02 (pent, J = 7.4 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 173.4, 155.1, 140.3, 126.7, 119.8, 112.6, 111.4, 33.2, 27.3, 21.7; positive ion TOFHRESIMS [M+H]+ m/z 178.0869 (calcd. for C10H12NO2, 178.0868).   NHOHN OO NaN 3 (2 eq)[HCl] +OH OH OH6.51B 6.51CHN OOH6.51B  322   Figure 6.45 1H and 13C NMR spectra of 6.51B in DMSO-d6 at 600 and 150 MHz.    323 Characterization of 6.51C:  6-hydroxy-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one: 1H NMR (600 MHz, DMSO-d6) δ 9.55 (s, 1H), 7.91 (bs, 1H), 7.09 (t, J = 8.0 Hz, 1H), 6.92 (d, J = 8.0 Hz, 2H), 2.86 (q, J = 7.7 Hz, 2H), 2.73 (t, J = 6.8 Hz, 2H), 1.78 (pent, J = 5.8 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 171.9, 153.8, 137.7, 126.9, 123.7, 118.6, 117.0, 38.5, 29.2, 20.8; positive ion TOFHRESIMS [M+H]+ m/z 178.0865 (calcd. for C10H12NO2, 178.0868).              NHOOH6.51C  324   Figure 6.46 1H and 13C NMR spectra of 6.51C in DMSO-d6 at 600 and 150 MHz.   325 Preparation of 6.52A, B and C:  Using the general procedure (p. 315) with 6, 7-dimethoxytetralone (0.5 g, 2.4 mmol). Purification of the crude reaction mixture by flash silica gel (eluting with 1:3, 1:1, 3:1 and 1:0 EtOAc/Hex, 200 mL each, 4 X 10 cm column) gave purified starting material (0.164 g), 6.52A (0.143 g, 35.4 %, BRSM), 6.52B (0.054 g, 15 %, BRSM), and 6.52C (0.077 g, 21.4 %, BRSM).  Characterization of 6.52A:  9, 10-dimethoxy-6, 7-dihydro-5H-benzo[c]tetrazolo[1, 5-a]azepine: 1H NMR (600 MHz, DMSO-d6) δ 7.74 (s, 1H), 7.00 (s, 1H), 4.64 (t, J = 6.6 Hz, 2H), 3.84 (s, 3H), 3.82 (s, 3H), 3.01 (m, 2H), 2.24 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 153.6, 150.9, 147.3, 134.4, 114.2, 113.6, 112.0, 55.64, 55.62, 49.4, 32.4, 24.8; positive ion TOFHRESIMS [M+H]+ m/z 247.1199 (calcd. for C12H15N4O2, 247.1195).     NHOHNMeOMeO MeOMeOONNMeOMeO NNOMeOMeO NaN 3 (2 eq)[HCl] + +6.52A 6.52B 6.52CNNMeOMeO NN6.52A  326   Figure 6.47 1H and 13C NMR spectra of 6.52A in DMSO-d6 at 600 and 150 MHz.    327 Characterization of 6.52B:  7, 8-dimethoxy-4, 5-dihydro-1H-benzo[b]azepin-2(3H)-one: 1H NMR (600 MHz, DMSO-d6) δ 9.24 (s, 1H), 6.86 (s, 1H), 6.58 (s, 1H), 3.73 (s, 3H), 3.70 (s, 3H), 2.60 (t, J = 7.2 Hz, 2H), 2.10 (obs, 2H), 2.07 (q, J = 6.7 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 173.3, 147.4, 145.6, 131.4, 125.5, 113.2, 106.5, 55.7, 55.6, 32.8, 29.2, 28.3; positive ion TOFHRESIMS [M+H]+ m/z 222.1131 (calcd. for C12H16NO3, 222.1130).               HNMeOMeO O6.52B  328   Figure 6.48 1H and 13C NMR spectra of 6.52B in DMSO-d6 at 600 and 150 MHz.   329 Characterization of 6.52C:  7, 8-dimethoxy-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one: 1H NMR (600 MHz, DMSO-d6) δ 7.89 (bt, J = 5.5 Hz, 1H), 7.04 (s, 1H), 6.86 (s, 1H), 3.79 (s, 3H), 3.75 (s, 3H), 2.91 (q, J = 6.1 Hz, 2H), 2.68 (t, J = 6.1 Hz, 2H), 1.86 (pent, J = 6.1 Hz, 2H); 13C NMR (150 MHz, DMSO-d6) δ 171.9, 150.3, 147.1, 131.3, 127.6, 112.0, 111.5, 55.5, 55.4, 38.6, 30.2, 29.4; positive ion TOFHRESIMS [M+H]+ m/z 222.1131 (calcd. for C12H16NO3, 222.1130).              NHOMeOMeO 6.52C  330   Figure 6.49 1H and 13C NMR spectra of 6.52C in DMSO-d6 at 600 and 150 MHz.   331 Preparation of 6.53A, B and C:  Using the general procedure (p. 315) with 7-fluoro-tetralone (0.5 g, 3.0 mmol) as starting material. Purification of the crude by flash silica gel chromatography (eluting with 1:4, 1:2, 1:1, 4:1, 1:0, EtOAc/Hex, 200 mL each, 4 X 10 cm column) gave purified starting material (0.2 g), 6.53A (0.126 g, 33.7 %, BRSM), 6.53B  (0.063 g, 19.4 %, BRSM), and 6.53C (0.047 g, 14.6 %, BRSM).  Characterization of 6.53A:  10-fluoro-6, 7-dihydro-5H-benzo[c]tetrazolo[1, 5-a]azepine: 1H NMR (600 MHz, acetone-d6) δ 7.97 (dd, J = 10.0, 3.4 Hz, 1H), 7.45 (d, J = 6.1 Hz, 1H), 7.27 (td, J = 8.7, 3.4 Hz, 1H), 4.74 (dd, J = 7.8, 6.1 Hz, 2H), 3.12 (m, 2H), 2.42 (pent, J = 6.6 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 163.0 (d, J = 240.5 Hz), 154.0 (d, J = 2.6 Hz), 137.9 (d, J = 3.1 Hz), 133.5 (d, J = 7.3 Hz), 126.1 (d, J = 9.9 Hz), 118.9 (d, J = 20.0 Hz), 116.7 (d, J = 24.8), 50.5, 33.1, 26.2; positive ion TOFHRESIMS [M+H]+ m/z 205.0891 (calcd. for C10H10N4F, 205.0891).   N HOHN ONN NNO NaN 3 (2 eq)[HCl] + +F F F F6.53A 6.53B 6.53CNN NNF 6.53A  332   Figure 6.50 1H and 13C NMR spectra of 6.53A in acetone-d6 at 600 and 150 MHz.    333 Characterization of 6.53B:  8-fluoro-4, 5-dihydro-1H-benzo[b]azepin-2(3H)-one: 1H NMR (600 MHz, acetone-d6) δ 8.74 (bs, 1H), 7.28 (bt, J = 7.0 Hz, 1H), 6.88 (td, J = 8.6, 2.4 Hz, 1H), 6.84 (dd, J = 8.6, 2.6 Hz, 1H), 2.76 (t, J = 7.0 Hz, 2H), 2.26 (t, J = 7.0 Hz, 2H), 2.15 (pent, J = 7.0 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 173.9, 162.7 (d, J = 251.7 Hz), 141.6 (d, J = 12.6 Hz), 131.9 (d, J = 10.1 Hz), 131.0, 112.0 (d, J = 21.4 Hz), 109.4 (d, J = 21.4 Hz), 33.7, 30.4, 29.0; positive ion TOFHRESIMS [M+H]+ m/z 180.0826 (calcd. for C10H11NOF, 180.0825).             HN OF 6.53B  334   Figure 6.51 1H and 13C NMR spectra of 6.53B in acetone-d6 at 600 and 150 MHz.   335 Characterization of 6.53C:  8-fluoro-2, 3, 4, 5-tetrahydro-1H-benzo[c]azepin-1-one: 1H NMR (600 MHz, acetone-d6) δ 7.38 (bs, 1H), 7.29 (m, 2H), 7.18 (td, J = 8.6, 2.8 Hz, 1H), 3.09 (q, J = 6.9 Hz, 2H), 2.82 (t, J = 7.4 Hz, 2H), 1.96 (pent, J = 7.4 Hz, 2H); 13C NMR (150 MHz, acetone-d6) δ 171.8, 162.5 (d, J = 247.3 Hz), 139.2 (d, J = 8.0 Hz), 135.2 (d, J = 3.7 Hz), 131.5 (d, J = 7.5 Hz), 118.1 (d, J = 21.0 Hz), 115.8 (d, J = 23.0 Hz), 39.7, 31.1, 30.1; positive ion TOFHRESIMS [M+H]+ m/z 180.0824 (calcd. for C10H11NOF, 180.0825).             NHOF 6.53C  336   Figure 6.52 1H and 13C NMR spectra of 6.53C in acetone-d6 at 600 and 150 MHz.    337 Preparation of 6.54:  To a stirred solution of 6.52A (0.02 g, 0.08 mmol) and ACN (5 mL) in a round bottom flask at 0 °C, was added KMnO4 (0.038 g, 0.24 mmol) in three portions over a 1 hour period.  The reaction mixture was stirred overnight then filtered and washed with 10 mL of acetone.  The excess KMnO4 was reacted with a 10% metapersulphate solution to give MnO2 as a precipitate, which was subsequently filtered away.  The filtrate was then concentrated and partitioned between 30 mL of H2O and EtOAc (3 X 10 mL).  The combined organic layers were washed with brine, dried over MgSO4, filtered, and concentrated.  The crude concentrate was purified using silica gel column chromatography (eluting with EtOAc/hexanes (1:1)) to yield 9, 10-dimethoxy-5H-benzo[c]tetrazolo[1, 5-a]azepin-7(6H)-one (6.54) as a white solid (0.016 g, 77%) 1H NMR (600 MHz, DMSO-d6) δ 7.76 (s, 1H), 7.64 (s, 1H), 4.83 (m, 2H), 3.95 (s, 3H), 3.90 (s, 3H), 3.81 (m, 1H), 3.24 (m, 2H); 13C NMR (150 MHz, DMSO-d6) δ 195.1, 153.1, 152.7, 150.7, 127.9, 116.6, 112.2, 111.4, 55.8, 41.6; positive ion HRESIMS [M+H]+ m/z 261.0992 (calcd. for C12H13N4O3, 261.0988).     NN N NMeO OMeONN N NMeOMeO KMnO4ACN6.52A 6.54  338   Figure 6.53 1H and 13C NMR spectra of 6.54 in DMSO-d6 at 600 and 150 MHz.   339 Preparation of 6.55:  To a small vial dried and under Ar atmosphere was added 6.54 (0.005 g, 0.02 mmol) in absolute MeOH (2 mL).  NaBH4 (1 mg, 0.02 mmol) was then added to the reaction. Once all starting material had been consumed (monitored by TLC), the reactions was concentrated and partitioned between H2O (10 mL) and EtOAc (3 X 5 mL).  The organic layers were combined, concentrated, and directly purified via silica gel column chromatography (eluting with EtOAc/Hex (7:3)) to give 9, 10-dimethoxy-6, 7-dihydro-5H-benzo[c]tetrazolo[1, 5-a]azepin-7-ol (6.55) as a white solid (4.2 mg, 84%). 1H NMR (600 MHz, DMSO-d6) δ 7.80 (s, 1H), 7.17 (s, 1H), 5.70 (s, 1H), 4.91 (bm, 1H), 4.67 (bm, 2H), 3.84 (s, 3H), 3.83 (s, 3H), 2.32 (bm, 2H); 13C NMR (150 MHz, DMSO-d6) δ 153.2, 150.7, 147.9, 137.1, 112.8, 112.1, 68.9, 55.7, 55.6, 44.0, 31.6; positive ion HRESIMS [M+Na]+ m/z 285.0965 (calcd. for C12H14N4O3Na, 285.0964).         NN N NMeO HOMeONN N NMeO OMeO NaBH4MeOH6.54 6.55  340   Figure 6.54 1H and 13C NMR spectra of 6.55 in DMSO-d6 at 600 and 150 MHz.   341 Preparation of 6.56 and 6.57:  Following a procedure from Suau et al.,243 to an Erlenmeyer flask containing 70 mL of ACN with 12.5 mL of H2O, was added 4-fluorophthalimide (0.5 g, 3.0 mmol)  and NaOH (1M, 1 mL).  The reaction vessel was capped using a rubber septum, and allyl trimethylsilyl ether (2.29 mL, 13.6 mmol, large excess) was added and the reaction mixture was degassed using a stream of N2.  The reaction was subsequently irradiated for 2 hours in an ice bath after which point the reaction was acidified using 1M HCl.  The mixture was then concentrated under reduced pressure and (~50 mL) H2O was added and the mixture was extracted with EtOAc (3 X 20 mL).  The organic layers were combined, dried over MgSO4, filtered, and concentrated under reduced pressure. Purification of the crude product using C18 reversed phase HPLC (eluting with 25 % ACN/H2O) gave the regio-isomers 6.56 (rt = 7.03 min, 0.003 g) and 6.57 (rt = 8.0 min, 0.0013 g). Characterization of 6.56:  7-fluoro-3-(hydroxymethyl)-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione: 1H NMR (600 MHz, acetone-d6) δ 7.94 (dd, J = 8.9, 5.4 Hz, 1H), 7.48 (td, J = 8.3, 3.0 Hz, 1H), 7.34 (dd, J = 9.5, 2.4 Hz, 1H), 7.22 (bs, 1H), 4.27 (bs, 1H), 3.99 (m, 1H), 3.78 (m, 2H),  3.01 (m, 2H); 13C NMR (150 MHz, acetone-d6)(combination of 13C and gHMBC data) δ 200.0, 167.2 (d, J ~ 208 Hz), 163.9, 139.1, 134.1 (d, J = 8.7 Hz), 129.8, 120.2 (d, J = 23.1 Hz), 115.1 (d, J = 23.6 Hz), 63.4, 51.4, 49.3; negative ion HRESIMS [M-H]- m/z 222.0566 (calcd. for C11H9NO3F, 222.0566).   NHOO OHFNHOOF h !N aOH, ACNOTMS NHOO OHF+6.56 6.57NHOO OHF 6.56  342   Figure 6.55 1H and 13C NMR spectra of 6.56 in acetone-d6 at 600 and 150 MHz.    343  Figure 6.56 gHMBC NMR spectra of 6.56 in acetone-d6 at 600 MHz. Characterization of 6.57:  8-fluoro-3-(hydroxymethyl)-3, 4-dihydro-1H-benzo[c]azepine-1, 5(2H)-dione: 1H NMR (600 MHz, acetone-d6) δ 7.76 (dd, J = 8.7, 5.7 Hz, 1H), 7.55 (dd, J = 9.6, 2.3 Hz, 1H), 7.44 (td, J = 8.4, 3.2 Hz, 1H), 7.35 (bs, 1H), 4.28 (bs, 1H), 3.99 (m, 1H), 3.78 (bd, J = 5.4 Hz, 2H), 2.98 (m, 2H); 13C NMR (150 MHz, acetone-d6) δ 200.9, 166.9 (d, J ~ 212.5 Hz), 166.6, 137.4, 134.3, 132.4 (d, J = 8.7 Hz), 119.4 (d, J  = 21.0 Hz), 117.2 (d, J = 24.1 Hz), 63.4, 51.5 (d, J = 17.0 Hz), 49.3; negative ion HRESIMS [M-H]- m/z 222.0562 (calcd. for C11H9NO3F, 222.0566).  NHOO OHF 6.57  344   Figure 6.57 1H and 13C NMR spectra of 6.57 in acetone-d6 at 600 and 150 MHz.   345 Chapter 7: Conclusion Throughout this work, isolation and development of natural products as tools and drug leads has been the guiding focus.  Initially, all of these compounds were isolated from organisms collected in either terrestrial (chapter 2 and 3), or marine (chapter 4–6) environments, underscoring the diversity of sources employed in this field.  Presented below is a chapter-by-chapter breakdown of the goals achieved and those still aspired to as future work.  In Chapter 2, a novel suite of mushroom metabolites, ramariolide A–D (2.18–2.21) was isolated from a previously unstudied source organism, the coral mushroom Ramaria cystidiophora.72 The compounds isolated were highly functionalized butenolides, which showed promising antimycobacterial activity. However, the number of electrophilic centers and inherent instability of these compounds likely will preclude further drug development. Due to the unusual source for these compounds and the relative abundance of this genus in the Pacific Northwest, it would be very interesting to see if these metabolites are wide spread. It would also be interesting to isolate and identify the enzymes responsible for the biosynthesis of the ramariolides.  The unusual and highly strained ring systems present in these metabolites make them unique and the enzymatic machinery responsible could be of great value to synthetic enzymology applications.250  The dhilirolides A–N (3.9–3.22) presented in Chapter 3, posses a series of novel carbon skeletons and a high degree of functionalization.132 The role of these compounds in nature is unclear, but feeding deterrence and insecticidal assays have hinted at a plausible ecological significance.  A proposed feeding deterrence assay performed with Sri Lankan native herbivorous insects would help answer if the dhilirolides provide the fungus or the host plant with any ecological advantage. Future research on the dhilirolides will focus on their insecticidal activity, and identify which member of the family is the best candidate for pesticide development.  The possibility of isolating and identifying the biosynthetic machinery responsible for the natural products may help to expedite future development.132  Chapter 4 and 5 presented the isolation and partial synthesis of several members of the xestoquinone family of compounds as IDO inhibitors.  One of the newly isolated compounds, xestolactone A (4.19), showed poor activity against hIDO, therefore, it would be interesting to screen this compound in other bioassays to determine if it could provide utility elsewhere.    346 Building on the IDO inhibitory activity seen from several natural products, a series of synthetic inhibitors have been constructed.162 These have the advantage of being easily and quickly synthetically accessible, and this work has led to the discovery of the minimal pharmacophore of this family of compounds in 5.33 (Figure 7.1).  The regio-specificity of the thiazine ring moiety, for maximal IDO inhibition, was also elucidated. It is likely that further development will focus on 5.24 and 5.33 (Figure 7.1) as hIDO inhibitor lead compounds for the treatment of cancer. In the future, the testing of 5.24 and 5.33 in cell and animal models will hopefully provide more information as to the potential these compounds have for preclinical development.   Figure 7.1 Xestosaprol O derivatives 5.24 and 5.33 showing the preferred thiazine ring regiochemistry  In the last chapter, the enzymatic target of latonduine A (6.9), PARP-3 (ARTD-3), and the activity-based protein-profiling probe (ABPP) constructed for its identification, were presented.79 More recent experiments, presented in Chapter 6, identified PARP-16 (ARTD-15) as another target for delF508-CFTR trafficking correction by latonduine.251 Current studies with this project are aimed at identification of the mechanisms by which inhibition of these enzymes induces correction.  The syntheses of novel and selective PARP-3 or PARP-16 inhibitors have further supported this dual inhibition mechanism of trafficking correction.79, 251 Lastly, the construction of an easily accessible corrector molecule 6.57 (Figure 7.2), which displays trafficking correction profiles comparable to latonduine A (6.9), was also accomplished.  Resolution of the enantiomers of 6.57 by chiral derivitization is underway to help determine which enantiomer is responsible for the correction activity observed.  Further development of 6.57 and analogues should focus on higher yielding synthetic methodology to produce enough material for preclinical development.  OOOSNHO OO OHOOSNHO O 5.335.24  347  Figure 7.2 Chiral derivitization and resolution plan for 6.57.  The future of each of these projects is unclear, however, the promising activity of many of these compounds in their respective bioassays will hopefully prompt their continued development. It is my hope that the work presented in the preceding five chapters has aided, in some small way, to the progression of the field of natural products chemistry. A research field, which has provided me with a profound sense of wonder and a near inexhaustible supply of motivation for continued research.              NH NHOF FOO OH O O NHOF O OR RChiral Derivatization +HPLC, characterization,and hydrolysisNHF OO OH NHF OO OH+6.57  348 References (1).     Solecki, R. S. Science 1975, 190, 880–881. (2).     Newman, D. J.; Cragg, G. M.; Snader, K. M. Nat. Prod. Rep. 2000, 17, 215–234. (3).     Newman, D. J.; Cragg, G. M.; Snader, K. M.  J. Nat. Prod. 2003, 66, 1022–1037. (4).     Ji, H-F.; Li, X-J.; Zhang, H-Y. EMBO 2009, 10, 194–200. (5).     Zhong, G. S.; Wan, F. Chin. J. Med. Hist. 1999, 29, 178–182. (6).     Yapijakis, C. In Vivo. 2009, 23, 507–514. (7).     Dev, S. Environ. Health Perspect. 1999, 107, 783–789. (8).     Klockgether-Radke, A. P. 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All measurements were made on a Bruker APEX DUO diffractometer with cross-­‐coupled	  multilayer	  optics	  Cu-Kα radiation.. The data were collected at a temperature of -183.0 + 0.1oC to a maximum 2θ value of 111.6o. Data were collected in a series of φ and ω scans in 2.0o oscillations using 30.0-second exposures. The crystal-to-detector distance was 49.92 mm.  Data Reduction  Of the 10914 reflections that were collected, 3627 were unique (Rint = 0.099); equivalent reflections, excluding Friedel pairs, were merged.  Data were collected and integrated using the Bruker SAINT software package. The linear absorption coefficient, µ, for Cu-Kα radiation is 6.71 cm-1. Data were corrected for absorption effects using the multi-scan technique (SADABS), with minimum and maximum transmission coefficients of 0.778 and 0.980, respectively.  The data were corrected for Lorentz and polarization effects. Structure Solution and Refinement  The structure was solved by direct methods.   The material crystallizes as a multi-component twin, however the results of the twinned integration were significantly less reliable than those obtained by integrating only the major twin domain.  The major twin domain is significantly larger than any subsequent twin components, as shown by the 'CELL_NOW' output using 85%of the harvested reflections to fit the major domain.  Unfortunately, the final data set, while sufficient to establish connectivity, does not result in a particularly satisfactory anisotropic refinement.  Most of the atoms anisotropic displacement parameters become NPD unless significant ISOR restraints are applied.  All non-hydrogen atoms were ultimately refined anisotropically.  All hydrogen atoms were placed in calculated positions, the hydroxyl hydrogen atoms placed using the HFIX 147 command.  The final cycle of full-matrix least-squares refinement on F2 was based on 3627 reflections and 363 variable parameters and converged   365 (largest parameter shift was 0.00 times its esd) with unweighted and weighted agreement factors of:  R1 = Σ ||Fo| - |Fc|| / Σ |Fo| = 0.160 wR2 = [ Σ ( w (Fo2 - Fc2)2 )/ Σ w(Fo2)2]1/2 = 0.352  The standard deviation of an observation of unit weight was 1.18. The weighting scheme was based on counting statistics.  The maximum and minimum peaks on the final difference Fourier map corresponded to 0.63 and –0.48 e-/Å, respectively.   Neutral atom scattering factors were taken from Cromer and Waber. Anomalous dispersion effects were included in Fcalc; the values for Δf' and Δf" were those of Creagh and McAuley. The values for the mass attenuation coefficients are those of Creagh and Hubbell. All refinements were performed using the SHELXL-97 via the WinGX interface. Crystal Data: Empirical Formula    C16H26O4 Formula Weight    282.37 Crystal Colour, Habit    colourless, blade Crystal Dimensions    0.03 x 0.12 x 0.40 mm Crystal System    monoclinic Lattice Type                Primitive Lattice Parameters    a = 13.231(2) Å       b = 5.1050(6) Å       c = 23.571(3) Å       α = 90o       β = 91.346(9)o       γ = 90o       V = 1591.5(3) Å3 Space Group     P 21  (#4) Z value     4 Dcalc      1.178 g/cm3 F000      616.00   366 µ(Cu-Kα)     6.71 cm-1 B. Intensity Measurements  Diffractometer    Bruker APEX DUO  Radiation     Cu-Kα (λ = 1.54178 Å) Data Images     1453 exposures @ 30.0 seconds Detector Position    49.92 mm 2θmax      111.6o No. of Reflections Measured Total:   10914 Unique:      3627 (Rint = 0.099; Friedels not merged) Corrections     Absorption (Tmin = 0.778, Tmax= 0.980)       Lorentz-polarization C. Structure Solution and Refinement  Structure Solution    Direct Methods (SIR97) Refinement     Full-matrix least-squares on F2 Function Minimized    Σ w (Fo2 - Fc2)2  Least Squares Weights   w=1/(σ2(Fo2)+(0.0309P) 2+ 25.5021P) Anomalous Dispersion   All non-hydrogen atoms No. Observations (I>0.00σ(I))  3627 No. Variables     363 Reflection/Parameter Ratio   9.99 Residuals (refined on F2, all data):   R1; wR2 0.160; 0.352 Goodness of Fit Indicator   1.18 No. Observations (I>2.00σ(I))  3282 Residuals (refined on F2):    R1; wR2 0.151; 0.347 Max Shift/Error in Final Cycle  0.00 Maximum peak in Final Diff. Map  0.63 e-/Å3 Minimum peak in Final Diff. Map  -0.48 e-/Å3     367 A.2 Compound 3.9 Data Collection: An irregular colourless crystal of C25H28O9 having approximate dimensions of 0.18 x 0.20 x 0.25 mm was mounted on a glass fiber. All measurements were made on a Bruker APEX DUO diffractometer with cross-coupled multilayer optics Cu-Kα radiation.  The data were collected at a temperature of -183.0 + 0.1°C to a maximum 2θ value of 134.8°. Data were collected in a series of ϕ and ω scans in 1.0° oscillations using 5.0-second exposures. The crystal-to-detector distance was 60.00 mm. Data Reduction: Of the 34345 reflections that were collected, 13627 were unique (Rint = 0.037; Friedels not merged); equivalent reflections were merged. Data were collected and integrated using the Bruker SAINT software package. The linear absorption coefficient, µ, for Cu-Κα radiation is 9.13 cm-1. Data were corrected for absorption effects using the multi-scan technique (SADABS), with minimum and maximum transmission coefficients of 0.780 and 0.848, respectively. The data were corrected for Lorentz and polarization effects. Structure Solution and Refinement: The structure was solved by direct methods. The material crystallizes as a two component twin, with twin domains related by a 180° rotation about the [-1 1 0] reciprocal lattice axis. The material crystallizes with four crystallographically independent molecules in the asymmetric unit. All non- hydrogen atoms were refined anisotropically. All hydrogen atoms were placed in calculated positions. The absolute configuration was determined on the basis of the refined Flack parameter. The final cycle of full-matrix least-squares refinement on F2 was based on 13627 reflections and 1266 variable parameters and converged (largest parameter shift was 0.00 times its esd) with unweighted and weighted agreement factors of: R1 = Σ ||Fo| - |Fc|| / Σ |Fo| = 0.042 wR2 = [ Σ ( w (Fo2 - Fc2)2 )/ Σ w(Fo2)2]1/2 = 0.108 The standard deviation of an observation of unit weight was 1.05. The weighting scheme was based on counting statistics. The maximum and minimum peaks on the final difference Fourier map corresponded to 0.19 and –0.21e-/Å, respectively. Neutral atom scattering factors were taken from Cromer and Waber. Anomalous dispersion effects were included in Fcalc; the values   368 for ∗f' and ∗f" were those of Creagh and McAuley. The values for the mass attenuation coefficients are those of Creagh and Hubbell. All refinements were performed using the SHELXL-97 via the WinGX interface. Experimental Details: Radiation      Cu-Κα (λ = 1.54178 Å)  Data Images      10441 exposures @ 5 seconds  Detector Position     60.00 mm 2θmax       134.8°  No. of Reflections Measured    Total: 34345 Corrections      Unique: 13627 (Rint = 0.037; Friedels not merged Absorption (Tmin = 0.780, Tmax= 0.848) Lorentz polarization Structure Solution Refinement:  Structure solution     Direct Methods (SIR97) Refinement      Full-matrix least-squares on F2 Function Minimized     Σ w (Fo2 - Fc2)2 Anomalous Dispersion    All non-hydrogen atoms No. Observations (I>0.00+(I))   13627 No. Variables      1266 Reflection/Parameter Ratio    10.76 Residuals (refined on F2, all data):    R1; wR2 0.042; 0.108  Goodness of Fit Indicator     1.05 No. Observations (I>2.00+(I))    13323 Residuals (refined on F): R1; wR2   0.040; 0.105 Max Shift/Error in Final Cycle   0.00  Maximum peak in Final Diff. Map   0.19 e-/Å3  Minimum peak in Final Diff. Map   -0.21 e-/Å3 Experimental Details: Empirical Formula      C25H28O9 Formula Weight     472.47   369 Crystal Colour, Habit     colourless, irregular Crystal Dimensions      0.18 X 0.20 X 0.25 mm Crystal System      triclinic Lattice Type       Primitive Lattice Parameters     a = 13.5779(5) Å        b = 13.6410(5) Å         c = 15.0542(6) Å        α = 65.339(2)°         β = 65.488(2)°         γ= 64.504(2)°         V=2191.1(1) Å3 Space Group      P1 (#1) Z value      4 Dcalc        1.432 g/cm3 F000       1000.00        μ(Mo-Kα) 9.13 cm-1 Diffractometer     Bruker APEX DUO                370 A.3 Compound 3.20 Data Collection: A purple, prism-like crystal of C25H28O7, having approximate dimensions of 0.15 x 0.23 x 0.24 mm was mounted on a glass fiber. All measurements were made on an X-Ray diffractometer with cross-coupled multilayer optics Cu-Kα radiation..  The data were collected at a temperature of -183.0 + 0.1oC to a maximum 2q value of 135.22o. Data were collected in a series of φ and w scans in 1o oscillations using 3.0-second exposures. The crystal-to-detector distance was 59.74 mm.  Data Reduction Of the 19567 reflections that were collected, 3836 were unique (Rint = 0.029); equivalent reflections, excluding Friedel pairs, were merged.  Data were collected and integrated using the SAINT software package. The linear absorption coefficient, m, for Cu-Ka radiation is 8.13 cm-1. Data were corrected for absorption effects using the multi-scan technique (SADABS), with minimum and maximum transmission coefficients of 0.918 and 0.981, respectively.  The data were corrected for Lorentz and polarization effects. Structure Solution and Refinement: The structure was solved by direct methods.   All non-hydrogen atoms were refined anisotropically.  All hydrogen atoms were placed in calculated positions.  The absolute configuration was determined on the basis of the refined Flack parameter [0.00(4)].  The final cycle of full-matrix least-squares refinement on F2 was based on 3836 reflections and 296 variable parameters and converged (largest parameter shift was 0.00 times its esd) with unweighted and weighted agreement factors of:  R1 = Σ ||Fo| - |Fc|| / Σ |Fo| = 0.027 wR2 = [ Σ ( w (Fo2 - Fc2)2 )/ Σ w(Fo2)2]1/2 = 0.070 The standard deviation of an observation of unit weight was 1.04. The weighting scheme was based on counting statistics.  The maximum and minimum peaks on the final difference Fourier map corresponded to 0.23 and –0.15 e-/Å, respectively.  Neutral atom scattering factors were taken from Cromer and Waber. Anomalous dispersion effects were included in Fcalc; the values for Df' and Df" were those of Creagh and McAuley.   371 The values for the mass attenuation coefficients are those of Creagh and Hubbell. All refinements were performed using the SHELXL-97 via the WinGX11 interface. Experimental Details: A. Crystal Data  Empirical Formula     C25H28O7 Formula Weight     440.47 Crystal Colour, Habit     purple, prism Crystal Dimensions     0.15 x 0.23 x 0.24 mm Crystal System     monoclinic Lattice Type      primitive Lattice Parameters     a = 11.4640(7) Å        b = 13.4589(8) Å        c = 14.0080(8) Å        α = 90°        β = 90°        γ = 90°        V = 2161.3(2) Å3 Space Group      P 212121  (#19) Z value      4 Dcalc       1.354 g/cm3 F000       936.00 m(Cu-Ka)      8.13 cm-1 B. Intensity Measurements    Radiation      Cu-Ka (l = 1.54178 Å) Data Images      4934 exposures @ 3 seconds Detector Position     59.74 mm 2qmax 135.22o No. of Reflections Measured Total:    19567 Unique:       3836 (Rint = 0.029; Friedels not merged)  Corrections Absorption (Tmin = 0.918, Tmax= 0.981) Lorentz-polarization   372 C. Structure Solution and Refinement  Structure Solution     Direct Methods (SIR97) Refinement      Full-matrix least-squares on F2 Function Minimized     S w (Fo2 - Fc2)2  Least Squares Weights    w=1/(s2(Fo2)+(0.0409P) 2+ 0.5097P) Anomalous Dispersion    All non-hydrogen atoms No. Observations (I>0.00s(I))   3836 No. Variables      296 Reflection/Parameter Ratio    12.96 Residuals (refined on F2, all data):    R1; wR2 0.027; 0.070 Goodness of Fit Indicator    1.04 No. Observations (I>2.00s(I))   3796 Residuals (refined on F2):                  R1; wR2 0.026; 0.070 Max Shift/Error in Final Cycle   0.00 Maximum peak in Final Diff. Map   0.23 e-/Å3 Minimum peak in Final Diff. Map   -0.15 e-/Å3    373 A.4 Compound 4.19 Data Collection:  An orange, irregular-like crystal of C19H16O5, having approximate dimensions of 0.08 x 0.14 x 0.23 mm was mounted on a glass fiber. All measurements were made on a Bruker APEX DUO diffractometer with cross-­‐coupled	  multilayer	  optics	  Cu-Kα radiation.. The data were collected at a temperature of -183.0 + 0.1oC to a maximum 2θ value of 130.90o. Data were collected in a series of φ and ω scans in 1.0o oscillations using 30.0-second exposures. The crystal-to-detector distance was 59.76 mm.  Data Reduction  Of the 7244 reflections that were collected, 2400 were unique (Rint = 0.039); equivalent reflections, excluding Friedel pairs, were merged.  Data were collected and integrated using the Bruker SAINT software package. The linear absorption coefficient, µ, for Cu-Kα radiation is 8.81 cm-1. Data were corrected for absorption effects using the multi-scan technique (SADABS), with minimum and maximum transmission coefficients of 0.780 and 0.932, respectively.  The data were corrected for Lorentz and polarization effects. Structure Solution and Refinement  The structure was solved by direct methods.   The absolute configuration at C17 (S) is based on the known configurations at carbons C3 and C6, and is corroborated by the refined Flack X-parameter [0.0(3)].  All non-hydrogen atoms were refined anisotropically.  Hydroxyl hydrogen H2O was located in a difference map and refined isotropically.  All other hydrogen atoms were placed in calculated positions.  The final cycle of full-matrix least-squares refinement on F2 was based on 2400 reflections and 223 variable parameters and converged (largest parameter shift was 0.00 times its esd) with unweighted and weighted agreement factors of:  R1 = Σ ||Fo| - |Fc|| / Σ |Fo| = 0.052 wR2 = [ Σ ( w (Fo2 - Fc2)2 )/ Σ w(Fo2)2]1/2 = 0.119  The standard deviation of an observation of unit weight was 1.08. The weighting scheme was based on counting statistics.  The maximum and minimum peaks on the final difference Fourier map corresponded to 0.27 and –0.23 e-/Å3, respectively.    374  Neutral atom scattering factors were taken from Cromer and Waber. Anomalous dispersion effects were included in Fcalc; the values for Δf' and Δf" were those of Creagh and McAuley. The values for the mass attenuation coefficients are those of Creagh and Hubbell. All refinements were performed using the SHELXL-97 via the WinGX interface. Experimental Details: A. Crystal Data  Empirical Formula   C19H16O5 Formula Weight   324.32 Crystal Colour, Habit   orange, irregular Crystal Dimensions   0.08 x 0.14 x 0.23 m Crystal System   orthorhombic Lattice Type    primitive Lattice Parameters   a = 8.5692(4) Å      b = 21.534(1) Å      c = 7.9705(4) Å      α = 90o      β = 90o      γ = 90o      V = 1470.8(1) Å3 Space Group    P 21212  (#18) Z value    4 Dcalc     1.465 g/cm3 F000     680.00 µ(Cu-Kα)    8.81 cm-1 B. Intensity Measurements  Diffractometer   Bruker APEX DUO  Radiation    Cu-Kα (λ = 1.54178 Å) Data Images    3215 exposures @ 30 seconds Detector Position   59.76 mm   375 2θmax     130.90o No. of Reflections Measured Total:  7244 Unique:     2400 (Rint = 0.039; Friedels not merged) Corrections Absorption (Tmin = 0.780, Tmax= 0.932) Lorentz-polarization C. Structure Solution and Refinement  Structure Solution   Direct Methods (SIR97) Refinement    Full-matrix least-squares on F2 Function Minimized   Σ w (Fo2 - Fc2)2  Least Squares Weights  w=1/(σ2(Fo2)+(0.0502P) 2+ 1.0927P) Anomalous Dispersion  All non-hydrogen atoms No. Observations (I>0.00σ(I)) 2400 No. Variables    223 Reflection/Parameter Ratio  10.76 Residuals (refined on F2, all data):  R1; wR2 0.052; 0.119 Goodness of Fit Indicator  1.08 No. Observations (I>2.00σ(I)) 2202 Residuals (refined on F2):   R1; wR2 0.048; 0.116 Max Shift/Error in Final Cycle 0.00 Maximum peak in Final Diff. Map 0.27 e-/Å3 Minimum peak in Final Diff. Map -0.23 e-/Å3    376 A.5 Compound 4.23 Data Collection:  A yellow, irregular crystal of C20H20O5.H2O, having approximate dimensions of 0.08 x 0.15 x 0.24 mm was mounted on a glass fiber. All measurements were made on a Bruker APEX DUO diffractometer with cross-­‐coupled	  multilayer	  optics	  Cu-Kα radiation. The data were collected at a temperature of -183.0 + 0.1oC to a maximum 2θ value of 132.52o. Data were collected in a series of φ and ω scans in 2o oscillations using 20.0-second exposures. The crystal-to-detector distance was 49.84 mm.  Data Reduction  Of the 15819 reflections that were collected, 2858 were unique (Rint = 0.026); equivalent reflections were merged.  Data were collected and integrated using the Bruker SAINT software package. The linear absorption coefficient, µ, for Cu-Kα radiation is 8.56 cm-1. Data were corrected for absorption effects using the multi-scan technique (SADABS), with minimum and maximum transmission coefficients of 0.849 and 0.934, respectively.  The data were corrected for Lorentz and polarization effects. Structure Solution and Refinement:  The structure was solved by direct methods.   The material crystallizes with one water molecule in the asymmetric unit.  The chiral centers at C2, C3, C6, and C13 have all been established as having the S-configuration, on the basis of the refined Flack12 x-parameter (0.05(5)).  All non-hydrogen atoms were refined anisotropically.  All O—H hydrogen atoms were located in difference maps and refined isotropically.  All other hydrogen atoms were placed in calculated positions.  The final cycle of full-matrix least-squares refinement4 on F2 was based on 2858 reflections and 252 variable parameters and converged (largest parameter shift was 0.00 times its esd) with unweighted and weighted agreement factors of:  R1 = Σ ||Fo| - |Fc|| / Σ |Fo| = 0.025 wR2 = [ Σ ( w (Fo2 - Fc2)2 )/ Σ w(Fo2)2]1/2 = 0.066  The standard deviation of an observation of unit weight was 1.06. The weighting scheme was based on counting statistics.  The maximum and minimum peaks on the final difference Fourier map corresponded to 0.13 and –0.19 e-/Å3, respectively.    377 Neutral atom scattering factors were taken from Cromer and Waber. Anomalous dispersion effects were included in Fcalc; the values for Δf' and Δf" were those of Creagh and McAuley. The values for the mass attenuation coefficients are those of Creagh and Hubbell. All refinements were performed using the SHELXL-2012 via the Olex2 interface. Experimental Details: Crystal Data Empirical Formula C20H22O6 Formula Weight 358.37 Crystal Colour, Habit yellow, irregular Crystal Dimensions 0.08 x 0.15 x 0.24 mm Crystal System monoclinic Lattice Type Primitive Lattice Parameters a = 6.2640(6) Å  b = 12.310(1) Å  c = 11.064(1) Å  α = 90o  β = 95.209(3)o  γ = 90o  V = 849.6(1) Å3 Space Group P21  (#4) Z value 2 Dcalc 1.401 g/cm3 F000 380.00 µ(Cu-Kα) 8.56 cm-1 Intensity Measurements Diffractometer Bruker APEX DUO  Radiation Cu-Kα (λ = 1.54178 Å Data Images 3640 exposures @ 20.0 seconds Detector Position 49.84 mm   378 2θmax 132.52o No. of Reflections Measured Total: 15819   Unique: 2858 (Rint = 0.026) Corrections Absorption (Tmin = 0.849, Tmax= 0.934)  Lorentz-polarization Structure Solution and Refinement Structure Solution Direct Methods (SIR97) Refinement Full-matrix least-squares on F2 Function Minimized Σ w (Fo2 - Fc2)2  Least Squares Weights w=1/(σ2(Fo2)+(0.036P) 2+ 0.2365P) Anomalous Dispersion All non-hydrogen atoms No. Observations (I>0.00σ(I)) 2858 No. Variables 252 Reflection/Parameter Ratio 11.34 Residuals (refined on F2, all data): R1; wR2 0.025; 0.066 Goodness of Fit Indicator 1.06 No. Observations (I>2.00σ(I)) 2828  Residuals (refined on F2): R1; wR2 0.025; 0.066 Max Shift/Error in Final Cycle 0.00 Maximum peak in Final Diff. Map 0.13 e-/Å3 Minimum peak in Final Diff. Map -0.19 e-/Å3  

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