Structure Elucidation and Synthesis
of New Secondary Metabolites from Liverworts and
Microorganisms and Investigation of their Biogenesis
DISSERTATION
In Fulfillment of the Requirements for the Degree of Dr. rer. nat
at the Institute of Organic Chemistry, University of Hamburg
by
Stephan Heinrich von Reuß
Structure Elucidation and Synthesis
of New Secondary Metabolites from Liverworts and
Microorganisms and Investigation of their Biogenesis
DISSERTATION
In Fulfillment of the Requirements for the Degree of Dr. rer. nat
at the Institute of Organic Chemistry, University of Hamburg
by
Stephan Heinrich von Reuß
1. Gutachter: Prof. Dr. Dr. h.c. mult. W. Francke
Institut für Organische Chemie, Universität Hamburg
2. Gutachter: Prof. Dr. C. Meier
Institut für Organische Chemie, Universiät Hamburg
3. Gutachter: Prof. Dr. W. Boland
Department für Bioorganische Chemie,
Max Plank Institut für Chemische Ökologie, Jena
4. Gutachter: Prof. Dr. L. Wessjohann
Abteilung für Natur- und Wirkstoffchemie
Leibniz Institut für Pflanzenbiochemie, Halle
The present work was performed from January 2004 to December 2008
under the supervision of Prof. Dr. Dr. h.c. mult. W. Francke in the laboratory of
the late Prof. Dr. W. A. König at the Institute of Organic Chemistry, as well as
in the laboratory of Dr. K. von Schwartzenberg at the Biozentrum Klein
Flottbek, University of Hamburg, Germany.
Acknowledgements
I like to express my deepest gratitude to the late Professor Dr. Wilfried A. König for his support and inspiring enthusiasm. He initiated this investigation but passed away on Nov.19. 2004, after this investigation has just been started.
I am most grateful to Professor Dr. Dr. h.c. Wittko Francke for the opportunity to continue the research under his supervision, his invaluable support, inspiring discussions and interesting cooperations.
I am very grateful to Dr. Klaus von Schwartzenberg (Biozentrum Klein Flottbek, University of Hamburg, Germany) for the opportunity to work under his guidance, his support in the establishment of in vitro cultures, and the very pleasant atmosphere.
Support by the following people is gratefully acknowledged: Dr. Volker Sinnwell and his team (Org. Chem.) for 1H and 13C NMR measurements, Dr. Erhard T. K. Haupt (Inorg. Chem.) for 2D NMR measurements and helpful discussions, Dr. Stephan Franke, Gaby Graak, Manfred Preuße and Annegret Meiners (Org. Chem.) for HREIMS, direct inlet EIMS and FAB-MS measurements, Professor Dr. Paul Margaretha (Org. Chem.) for advice with the UV isomerisations, Dr. Dietmar Keyser (Zoologisches Institut) for Scanning Electron Microscopy (SEM), S. Bringe (Biozentrum Klein Flottbek) for technical assistance, and Dr. Hermann Muhle (Abteilung Systematische Botanik und Ökologie, University Ulm, Germany) for collection of requested liverworts and the joined fieldtrip to Portugal (March 2005), Professor Dr. Yoshinori Asakawa (Tokushima, Bunri University, Japan), Professor Dr. Cecília Sérgio (Herbário do Museu, Laboratório et Jardin Botânico de Lisboa, Portugal), Dr. Pavel Pribyl (Culture Collection of Autotrophic Organisms, Trebon, Czech Republic), Dr. Tassilo Feuerer (Herbarium Hamburgense, Hamburg, Germany) and Professor Dr. S. Robbert Gradstein (A-v-H Institute, Göttingen, Germany) for providing plant material of Corsinia coriandrina and
Cronisia weddellii, respectively, Professor Dr. Wilhelm Boland (Max Planck Institute for
Chemical Ecology, Jena, Germany) for providing reference compounds of brown algal pheromones, Dr. Alexey V. Tkachev (Novosibirsk Institute of Organic Chemistry, Novosibirsk, Russia) for providing solid super acid, Dr. Detlef Hochmuth for providing the MassFinder Software, and Dr. J. Rheinheimer (BASF AG, Ludwigshafen, Germany) for biotests with synthetic Corsinia constituents..
Special thanks to all cooperative partners who significantly enriched my research:
Dr. Simon Jones (Department of Chemistry, University of Sheffield, Great Britain) for the enantioselective synthesis of (R)-(–)-corsifuran A.
Professor Gerhard Gries and his group (Department of Biological Sciences, Simon Fraser University, Burnaby, Canada) for providing (+)-spiroaxa-5,7-diene from Ulmus americana infected with Ophiostoma novo-ulmi.
Professor Birgit Piechulla and Marco Kai (Institut für Biowissenschaften, University of Rostock, Germany) for providing headspace samples from Serratia odorifera.
I also like to thank Professor Chia Li-Wu (Department of Organic Chemistry, Tamkang University, Tan Shui, Taiwan), Professor Danuta Kalemba (Technical University of Lodz, Institute of General Food Chemistry, Lodz, Poland), Dr. Claudia Höckelmann (Givaudan AG, Dübendorf, Switzerland), and Dr. Florian Schiestl (Geobotanical Institute, ETH-Zürich, Switzerland) for interesting cooperations.
Thanks to former members of W. A. König„s research group: Dr. Fernando Campos Ziegenbein, Dr. Dennis Lass, Dr. Rita Richter, Dr. Heilemichael Tesso, Dr. Frank Werner, and especially Dr. Adewale Martins Adio, Dr. Simla Basar, Dr. Thomas Hackl and Dr. Peter Hansen; members of W. Francke‟s research group: Dr. Robert Twele and Armin Troeger; and members of K. von Schwartzenberg‟s research group: Dr. Hanna Richter, Dr. Natalya Yevdakova, Hanna Turcinov, and Marta Fernandez Nunez.
Financial Support by the „Deutsche Forschungsgemeinschaft” (DFG) and the „Deutsche Akademische Austauschdienst” (DAAD) is gratefully acknowledged.
I also like to thank Sascha Ludolph for kindly providing technical equipment and Mario Maiworm for computer programming.
Table of
Contents
1. Introduction ... 1
2. Objective of Research ... 2
3. General Part ... 3
3.1. Liverwort Biology ... 3
3.2. Secondary Metabolites of Liverworts ... 5
4. Special Part ... 7
4.1. C11 Hydrocarbons from Fossombronia angulosa ... 7
4.1.1. Fossombronia angulosa ... 7
4.1.2. Volatile Constituents of Fossombronia angulosa ... 8
4.1.3. Identification of Dictyopterene A (29) ... 11
4.1.4. Identification of Dictyotene (32) ... 12
4.1.5. Identification of Ectocarpene (30) ... 13
4.1.6. Identification and Synthesis of n-Pentylbenzene (31) ... 14
4.1.7. Enantioselective GC Analysis of C11 Hydrocarbons ... 15
4.1.8. Discussion of C11 Hydrocarbons from Fossombronia angulosa ... 16
4.2. Indole Alkaloids from Riccardia chamedryfolia ... 17
4.2.1. Riccardia chamedryfolia ... 17
4.2.2. Volatile Constituents of Riccardia chamedryfolia ... 17
4.2.3. Identification and Synthesis of 3-Chloro-6- and 7-prenylindoles (49, 50) ... 20
4.2.4. Identification of Chamedryfolian (51) ... 23
4.2.5. Identification of 7-(3-Methylbutadienyl)indole (52)... 26
4.2.6. Discussion of Indole Alkaloids from Riccardia chamedryfolia... 27
4.3. Secondary Metabolites of Corsinia coriandrina ... 28
4.3.1. Taxonomy of the Corsiniaceae ... 28
4.3.2. Volatile Constituents of Corsinia coriandrina ... 29
4.3.3. Identification and Synthesis of 4-Methoxystyrenes from Corsinia coriandrina ... 32
4.3.3.1. (Z/E)-Coriandrins (65) ... 32 4.3.3.2. (Z/E)-O-Methyltridentatols B & A (72)... 36 4.3.3.3. O-Methyltridentatol C (71) ... 37 4.3.3.4. (Z/E)-Corsinians (63) ... 39 4.3.3.5. (Z/E)-Corsiandrens (67) ... 41 4.3.3.6. (Z/E)-Corsiandrenins (92) ... 43 4.3.3.7. (Z/E)-Tuberines (98) ... 45 4.3.3.8. (Z/E)-Corsicillins (99) ... 46
4.3.3.9. Discussion of 4-Methoxystyrenes from Corsinia coriandrina ... 48
4.3.4. Identification and Synthesis of Stilbenoids from Corsinia coriandrina ... 51
4.3.4.2. Corsifuran B (114) ... 55
4.3.4.3. Corsifuran C (74) ... 56
4.3.4.4. (E/Z)-Corsistilbenes (68) ... 57
4.3.4.5. O,O-Dimethyllunularin (69) ... 60
4.3.4.6. 6-Hydroxy-O,O-dimethyllunularin (70) ... 61
4.3.4.7. Discussion of Stilbenoids from Corsinia coriandrina ... 63
4.4. Synthesis of Deuterium Labelled Precursors ... 67
4.4.1. Synthesis of (E)-[2-D]-Cinnamic Acids ([2-D]-89, [2-D]-93, [2-D]-152) ... 67
4.4.2. Synthesis of [3,3-D2]-3-Phenylpropanoic Acids (153 – 158) ... 71
4.4.3. Synthesis of L-[CD3]-O-Methyltyrosine ([CD3]-155) ... 72
4.4.4. Synthesis of DL-[2-D]-Tyrosines ([2-D]-154, [2-D]-155) ... 73
4.4.5. Synthesis of DL-[2,3-threo-D2]-Tyrosine ([2,3-threo-D2]-154) ... 73
4.4.6. Synthesis of [CD3]-O-Methyltyramine ([CD3]-168) ... 74
4.4.7. Synthesis of (±)-[2,3-threo-D2]-Phloretic Acid ((±)-[2,3-threo-D2]-157) ... 75
4.5. Axenic In Vitro Cultures of Corsinia coriandrina ... 76
4.5.1. Collection of Corsinia coriandrina Spores ... 76
4.5.2. Germination of Corsinia Spores and Propagation of Monoclonal Strains ... 77
4.5.3. Determination of Ploidy Levels ... 77
4.5.4. Optimization of Culture Media and Conditions ... 78
4.5.5. Mixed Photo-Heterotrophic Growth ... 79
4.5.6. Aerated Liquid Submersion Cultures ... 80
4.5.7. Temporarily Immersed Cultures using RITA® ... 82
4.5.8. Chemical Investigation of In Vitro Cultured Corsinia coriandrina ... 82
4.6. Application Experiments ... 84
4.6.1. Biosynthesis of Coriandrin in Corsinia coriandrina ... 86
4.6.1.1. The L-Tyrosine Origin of Coriandrin. ... 87
4.6.1.2. Pulsed Application Experiments using RITA® ... 90
4.6.1.3. Tyrosine Decarboxylase Activity ... 92
4.6.1.4. Tyrosine Ammonia Lyase Activity... 93
4.6.1.5. O-Methyl Transferase Activity ... 93
4.6.1.6. Stereospecific Elimination of the 3-Pro-S-Hydrogen of L-Tyrosine ... 95
4.6.1.7. The Glycine Origin of the O-Methyl Group of (Z)-Coriandrin ... 96
4.6.1.8. Discussion of Coriandrin Biosynthesis in Corsinia coriandrina ... 98
4.6.2. Biosynthesis of Corsifuran A in Corsinia coriandrina ... 102
4.6.2.1. Application Experiments using GC-EIMS Detection ... 102
4.6.2.2. Application Experiments using 2D and 13C NMR Spectroscopy ... 103
4.6.2.3. The Phenylpropenoid Origin of Corsifuran A and Corsistilbene. ... 103
4.6.2.4. The Phenylpropenoid Origin of Aromatic Corsifuran C. ... 106
4.6.2.5. Application of Dihydrocinnamic Acids ... 109
4.6.2.7. Phenylalanine Transaminase Activity ... 114
4.6.2.8. Application of D-3,3-Dideuterophenylalanine... 116
4.6.2.9. The Polymalonate Origin of Corsifuran A ... 118
4.6.2.10. -Carbonyl Group Reduction in Corsifuran A Biosynthesis ... 120
4.6.2.11. Exclusion of Stilbenecarboxylic Acid Intermediates ... 123
4.6.2.12. Discussion of Corsifuran A Biosynthesis in Corsinia coriandrina ... 126
4.6.3. Biosynthesis of Terpenoids in Corsinia coriandrina ... 130
4.6.3.1. Biosynthesis of (R)-(–)-(E)-Nerolidol ... 130
4.6.3.2. Biosynthesis of (4S,6S)-( )- -Pinene ... 132
4.6.3.3. Discussion of Terpenoid Biosynthesis in Corsinia coriandrina... 136
4.7. (1S,2R)-(+)-Spiroaxa-5,7-diene associated with Dutch Elm Disease ... 137
4.7.1. Durch Elm Disease (DED) ... 137
4.7.2. Identification of (1S,2R)-(+)-Spiroaxa-5,7-diene ... 139
4.7.3. Attempted Dehydration of (–)-Axenol ... 141
4.7.4. Acid Catalyzed Rearrangement using Amberlyst ... 142
4.7.5. Solid Super Acid Catalyzed Rearrangement using TiO2/SO4 2– ... 144
4.7.6. Discussion of (1S,2R)-(+)-Spiroaxa-5,7-diene from Ulmus americana ... 148
4.8. Octamethylbicyclo[3.2.1]octadienes from Serratia odorifera ... 149
4.8.1. Serratia odorifera ... 149
4.8.2. Identification of Odorifen ... 149
4.8.3. Synthesis of Octamethylbicyclo[3.2.1]octadienes ... 152
4.8.4. Synthesis of Octamethylbicyclo[3.2.1]octa-2,6-diene and -2(10),6-diene ... 154
4.8.5. Biosynthesis of Odorifen ... 156
4.8.6. Discussion of Octamethylbicyclo[3.2.1]octadienes from Serratia odorifera ... 157
5. Summary ... 158
6. Zusammenfassung ... 160
7. Experimental Part ... 162
7.1. General Experimental Procedures ... 162
7.1.1. Nuclear Magnetic Resonance Spectroscopy (NMR) ... 162
7.1.2. Electron Impact Mass Spectrometry (EIMS) ... 162
7.1.3. High Resolution Electron Impact Mass Spectrometry (HREIMS) ... 162
7.1.4. Fast Atom Bombardment Mass Spectrometry (FAB-MS) ... 162
7.1.5. Gas Chromatography (GC) ... 162
7.1.6. Enantioselective Gas Chromatography (eGC) ... 163
7.1.7. Preparative Gas Chromatography (PGC) ... 163
7.1.8. Semi-preparative Gas Chromatography (SPGC) ... 163
7.1.9. Column Chromatography (CC) ... 163
7.1.10. Preparative Thin Layer Chromatography (TLC) ... 163
7.1.11. Polarimetry ... 164
7.1.13. Ultraviolet Spectroscopy (UV) ... 164
7.1.14. UV Induced (E/Z)-Isomerisations ... 164
7.1.15. Photography ... 164
7.1.16. Microphotography ... 164
7.1.17. Fluorescence Micrsocopy ... 164
7.1.18. Scanning Electron Microscopy (SEM) ... 164
7.1.19. Quantum Mechanical Calculations ... 164
7.2. Plant Materials & Extracts ... 165
7.2.1. Origin of Plant Materials ... 165
7.2.2. Deposition of Plant Material ... 165
7.2.3. Hydrodistillation ... 165
7.2.4. Extraction of Air Dried Plant Material ... 166
7.2.5. Extraction of Fresh Plant Material ... 166
7.3. Axenic In Vitro Cultures of Corsinia coriandrina ... 166
7.3.1. Plant media ... 166
7.3.2. Establishment of Axenic In Vitro Cultures ... 167
7.3.3. Flow Cytometry ... 167
7.3.4. Application Experiments ... 167
7.4. Fossombronia angulosa ... 168
7.4.1. Isolation of C11 Hydrocarbons from Fossombronia angulosa ... 168
7.4.2. Synthesis of (±)-1-Phenylpentan-1-ol (41) ... 168
7.4.3. Synthesis of (E)-1-Phenylpent-1-ene (42) ... 169
7.4.4. Synthesis of n-Pentylbenzene (31) ... 169
7.5. Riccardia chamedryfolia ... 169
7.5.1. Isolation of Prenylindoles (14 and 15) ... 169
7.5.2. Synthesis of 3-Chloro-7-prenylindole (49) ... 170
7.5.3. Synthesis of 3-Chloro-6-prenylindole (50) ... 170
7.5.4. Isolation of Chamedryfolian (51) ... 170
7.5.5. Chemical Correlation of Chamedryfolian (51) ... 171
7.5.6. Chemical Correlation of 7-(3-Methylbutadienyl)indole (52) ... 171
7.6. Corsinia coriandrina ... 171
7.6.1. Isolation of Secondary Metabolites ... 171
7.6.1.1. Isolation of ( )- -Pinene (18) ... 171
7.6.1.2. Isolation of ( )-(E)-Nerolidol (64) ... 172
7.6.1.3. Isolation of (E,E)- -Springene (66) ... 172
7.6.1.4. Isolation of (Z)-Coriandrin ((Z)-65) ... 172
7.6.1.5. Isolation of (Z)-O-Methyltridentatol B ((Z)-72) ... 172
7.6.1.6. Isolation of (R)-( -Corsifuran A (73) ... 172
7.6.1.7. Isolation of Corsifuran C (74) ... 173
7.6.1.9. Isolation of 4-Methoxyphenylethanal (59) ... 173
7.6.2. Synthesis of Secondary metabolites from Corsinia coriandrina ... 173
7.6.2.1. Synthesis of 2-Bromo-1-(4-methoxyphenyl)ethanone (76) ... 173
7.6.2.2. Synthesis of 2-Azido-1-(4-methoxyphenyl)ethanone (77) ... 173
7.6.2.3. Synthesis of (±)-2-Amino-1-(4-methoxyphenyl)ethanol (78) ... 174
7.6.2.4. Synthesis of (±)-2-Hydroxy-2-(4-methoxyphenyl)ethylammonium Chloride (79) ... 174
7.6.2.5. Synthesis of (±)-2-Chloro-2-(4-methoxyphenyl)ethylammonium Chloride (80) ... 174
7.6.2.6. Synthesis of (±)-2-Chloro-2-(4-methoxyphenyl)ethyl Isothiocyanate (81) ... 175
7.6.2.7. Synthesis of (Z)- and (E)-Coriandrins (65) ... 175
7.6.2.8. Synthesis of (Z)- and (E)-O-Methyltridentatols (72) ... 176
7.6.2.9. Synthesis of (±)-5-(4-Methoxyphenyl)-2-S-methylthio-4,5-dihydrothiazole (82) ... 176
7.6.2.10. Synthesis of O-Methyltridentatol C (71) ... 177
7.6.2.11. Synthesis of (±)-2-Chloro-2-(4-methoxyphenyl)ethyl Isocyanate (87) ... 177
7.6.2.12. Synthesis of (Z)- and (E)-Corsinians (63)... 178
7.6.2.13. Synthesis of (E)-4-Methoxycinnamic acid (89) by Knoevenagel ... 178
7.6.2.14. Synthesis of (E)-3-(4-Methoxyphenyl)propenoyl Chloride (90) ... 178
7.6.2.15. Synthesis of (E)-3-(4-Methoxyphenyl)propenoyl Azide (91) ... 178
7.6.2.16. Synthesis of (Z)- and (E)-Corsinians (63) by Curtius Rearrangement ... 179
7.6.2.17. Synthesis of (Z)- and (E)-Corsiandrens (67) ... 179
7.6.2.18. Synthesis of (Z)- and (E)-Corsiandrenins (92) ... 180
7.6.2.19. Synthesis of (E)-Cinnamic Acid (93) ... 181
7.6.2.20. Synthesis of (E)-3-Phenylpropenoyl Chloride (94) ... 181
7.6.2.21. Synthesis of (E)-3-Phenylpropenoyl Azide (95) ... 181
7.6.2.22. Synthesis of (E)-2-Phenylethenyl Isocyanate (96) ... 181
7.6.2.23. Synthesis of Dehydroniranin A (97) ... 182
7.6.2.24. Synthesis of (Z)- and (E)-Tuberines (98) ... 182
7.6.2.25. Synthesis of (Z)- and (E)-Corsicillins (99) ... 183
7.6.2.26. Synthesis of (E)-4-Methoxycinnamonitrile (100) ... 183
7.6.2.27. Synthesis of (±)-Corsifuran B (114) ... 183
7.6.2.28. Synthesis of (±)-Corsifuran A (73) ... 184
7.6.2.29. Synthesis of (±)-[5-OCD3]-Corsifuran A ([5-OCD3]-73) ... 184
7.6.2.30. Synthesis of Corsifuran C (74) ... 185
7.6.2.31. Synthesis of 4-Methoxybenzyl chloride (126) ... 185
7.6.2.32. Synthesis of Diethyl-4-methoxybenzyl phosphonate (127) ... 185
7.6.2.33. Synthesis of (E)-Corsistilbene ((E)-68) ... 186
7.6.2.34. Isomerisation to (Z)-Corsistilbene ((Z)-68) ... 186
7.6.2.35 Synthesis of O,O-Dimethyllunularin (69) ... 187
7.6.2.36. Synthesis of 2-Hydroxy-5,4‟-dimethoxybibenzyl (70) ... 187
7.6.3. Synthesis of Deuterium Labelled Precursors ... 187
7.6.3.2. Synthesis of (E)-[2-D]-Cinnamic Acid ([2-D]-93) by Perkin... 188
7.6.3.3. Synthesis of (E/Z)-Cinnamonitriles (149)... 188
7.6.3.4. Synthesis of (E/Z)-[2-D]-Cinnamonitriles ([2-D]-149) ... 188
7.6.3.5. Attempted Synthesis of (E)-[2-D]-Cinnamic Acid ([2-D]-93) ... 189
7.6.3.6. Attempted 1H/2D Exchange in Potassium (E)-Cinnamate ... 189
7.6.3.7. Synthesis of (E)-[2-D]-Cinnamic Acid ([2-D]-93) by Hydrolysis with KOD ... 189
7.6.3.8. Synthesis of (E)-[2-D]-Cinnamic Acid ([2-D]-93) by Hydrolysis with NaOD ... 189
7.6.3.9. Synthesis of (E)-4-Methoxycinnamic Acid (89) by Hydrolysis ... 190
7.6.3.10. Synthesis of (E)-[2-D]-4-Methoxycinnamic acid ([2-D]-89) by Hydrolysis ... 190
7.6.3.11. Synthesis of (E)-4-Hydroxycinnamonitrile (151) ... 190
7.6.3.12. Synthesis of (E)-4-Coumaric Acid (152) by Hydrolysis ... 191
7.6.3.13. Synthesis of (E)-[2-D]-4-Coumaric Acid ([2-D]-152) by Hydrolysis ... 191
7.6.3.14. Synthesis of [3,5-D2]-4-Hydroxybenzaldehyde ([D2]-150) ... 191
7.6.3.15. Synthesis of [D4]-Malonic Acid ([D4]-88) ... 192
7.6.3.16. Synthesis of (E)-[2,3‟,5‟-D3]-4-Coumaric Acid ([D3]-152) by Knoevenagel... 192
7.6.3.17. Synthesis of L- and D-[3,3-D2]-Phenylalanine (L-[3,3-D2]-153, D-[3,3-D2]-153) ... 192
7.6.3.18. Synthesis of L-[3,3-D2]-Tyrosine (L-[3,3-D2]-154) ... 193
7.6.3.19. Synthesis of L-[3,3-D2]-O-Methyltyrosine (L-[3,3-D2]- 155) ... 193
7.6.3.20. Synthesis of [3,3-D2]-Dihydrocinnamic Acid ([3,3-D2]-156) ... 193
7.6.3.21. Synthesis of [2,2,3,3,3‟-D5]-Phloretic Acid ([D5]-157) ... 194
7.6.3.22. Synthesis of [2,2,3‟-D3]-Tyramine ([2,2,3‟-D3]-158) ... 194
7.6.3.23. Synthesis of L-N-Acetyltyrosine (159) ... 195
7.6.3.24. Synthesis of [CD3]-Methyl L-N-Acetyl-[CD3]-O-methyltyrosinate ([CD3]-160) ... 195
7.6.3.25. Synthesis of L-[CD3]-O-Methyltyrosine Hydrochloride ([CD3]-155) ... 195
7.6.3.26. Conversion to L-[CD3]-O-Methyltyrosine ([CD3]-155) ... 196
7.6.3.27. Synthesis of DL-[2-D]-N,O-Diacetyltyrosine ([2-D]-161) ... 196
7.6.3.28. Synthesis of DL-[2-D]-Tyrosine Hydrochloride ([2-D]-154) ... 196
7.6.3.29. Synthesis of L-N-Acetyl-O-methyltyrosine (162) ... 197
7.6.3.30. Synthesis of DL-[2-D]-N-Acetyl-O-methyltyrosine ([2-D]-162) ... 197
7.6.3.31. Synthesis of DL-[2-D]-O-Methyltyrosine Hydrochloride ([2-D]-155) ... 197
7.6.3.32. Synthesis of (Z)-4-(4-Acetoxybenzylidene)-2-methyloxazol-5(4H)-one (164) ... 198
7.6.3.33. Synthesis of (Z)-2-Acetamido-3-(4-acetoxyphenyl)propenoic Acid (165) ... 198
7.6.3.34. Synthesis of DL-[2,3-threo-D2]-N,O-Diacetyltyrosine ([2,3-threo-D2]-161) ... 198
7.6.3.35. Synthesis of DL-[threo-2,3-D2]-Tyrosine Hydrochloride ([2,3-threo-D2]-154) ... 199
7.6.3.36. Synthesis of [CD3]-4-Methoxybenzaldehyde ([CD3]-58) ... 199
7.6.3.37. Synthesis of [CD3]-4-Methoxy- -nitrostyrene ([CD3]-167) ... 199
7.6.3.38. Synthesis of [CD3]-O-Methyltyramine ([CD3]-168) ... 200
7.6.3.39. Synthesis of (±)-[2,3-threo-D2]-Phloretic Acid ([2,3-threo-D2]-157) ... 200
7.7. (1S,2R)-(+)-spiroaxa-5,7-diene from Ulmus americana ... 201
7.7.2. Amberlyst Catalyzed Rearrangement of (+)-Aromadendrene ((+)-216) ... 201
7.7.3. Amberlyst Catalyzed Rearrangement of (+)-Ledene (218) ... 201
7.7.4. Amberlyst Catalyzed Rearrangement of (–)-allo-Aromadendrene (217) ... 201
7.7.5. Solid Superacid Catalyzed Rearrangement of (+)-Aromadendrene ((+)-216) ... 202
7.7.6. Solid Superacid Catalyzed Rearrangement of (+)-Ledene (218) ... 202
7.7.7. Solid Superacid Catalyzed Rearrangement of (–)-allo-Aromadendrene (217) ... 203
7.7.8. Isolation of (–)-ent-Aromadendrene ((–)-216) from Pellia epiphylla ... 203
7.7.9. Solid Superacid Catalyzed Rearrangement of (–)-ent-Aromadendrene ((–)-216) ... 203
7.8. Serratia odorifera - Synthesis of Octamethylbicyclo[3.2.1]octadienes ... 203
7.8.1. Synthesis of 2,4-Dibromopentan-3-one (232) ... 203 7.8.2. 1,2,4,5,6,7,8-Heptamethylbicyclo[3.2.1]oct-6-en-3-ones (236 - 238) ... 203 7.8.3. bisaxial-1,2,4,5,6,7,8-Heptamethyl-3-methylenebicyclo[3.2.1]oct-6-ene (239) ... 204 7.8.4. bisequatorial-1,2,4,5,6,7,8-Heptamethyl-3-methylenebicyclo[3.2.1]oct-6-ene (241) ... 205 7.8.5. 1,2,3,4,5,6,7,8-Octamethylbicyclo[3.2.1]oct-6-en-3-ol (242) ... 205 7.8.6. 1,2,3,4,5,6,7,8-Octamethylbicyclo[3.2.1]octa-2,6-diene (243) and ... 206 1,3,4,5,6,7,8-Heptamethyl-2-methylenebicyclo[3.2.1]oct-6-ene (244) ... 206 8. Hazardous chemicals ... 207 9. Colour Plates ... 210 10. References ... 218 Poster Presentations ... 240 Oral Presentations ... 241 Publications ... 242 CURRICULUM VITAE ... 244 List of Abbreviations ... 245
1
1. Introduction
»Organic Chemists have often been tempted to leave the security of their own proper
pastures and to graze, albeit speculatively, in the attractive fields of biochemistry. It has seemed that, the structures of so many natural products having been established, it should be possible to perceive some relations between them such as to suggest schemes of biosynthesis.« Sir Robert Robinson, 1955
While primary metabolism refers to the anabolic and catabolic processes required for cell maintenance and proliferation, secondary metabolism covers species specific natural products that are not necessary for the individual cell, but thought to be required for the organisms‟ survival when interacting with its environment. The amazing structural diversity displayed by natural products makes them a rewarding field for structure elucidation and synthesis. Apart from the desire to isolate, identify and synthesize natural products in order to determine their biological activity in an ecological or pharmacological context, our ever increasing knowledge on their distribution among the different species allows for chemosystematic classifications. Furthermore, since the availability of radioactive isotopes in the 1950‟s, the discovery of new natural products has initiated the elucidation of precursor-product relationships by application experiments with labelled compounds. Advances in NMR spectroscopy then enabled application experiments using stable isotopes like 2D, 13C, 15N, and
18
O (Schneider, 2007; Simpson, 1998; Vederas, 1987; Garson & Staunton, 1979). The biosynthetic routes of many secondary pathways were outlined and provided the basis for the enzymatic characterization of biosynthetic pathways in the 1970‟s and 1980‟s, followed by the identification of the corresponding genes beginning in the late 1980‟s (Thomas, 2004; Hartmann, 2007; Mahmud, 2007).
Liverworts (Hepaticae) have recently attracted attention, due to their phylogenetic position basal to all other terrestrial plants (Qiu et al., 2006, 2007). Phytochemical studies on liverworts are however hampered by the difficulties encountered with the acquisition of sufficient quantities of pure plant material, unless axenic in vitro cultures are available (Becker, 1990, 1994, 1995). It has been estimated that only 5 % of known liverwort species have been cultured under in vitro conditions (Duckett et al., 2004; Hohe & Reski, 2005) and less than 15 % have been chemically investigated so far (Asakawa, 1995, 2007, 2008).
2
2. Objective of Research
The purpose of this study was the isolation, structure elucidation and synthesis of secondary metabolites from liverworts and microorganisms, along with the investigation of their biosynthesis. Research involves the acquisition of fresh liverwort plant material and its botanical identification, along with the development of suitable extraction procedures monitored by gaschromatography (GC). Unknown constituents are traced using coupled gaschromatography electron impact mass spectrometry (GC-EIMS), in combination with a mass spectral database (König & Hochmuth, 2004; König et al., 2004). Their isolation is achieved by a combination of column chromatography (CC) and thin layer chromatography (TLC) or preparative gaschromatography (PGC). Molecular formulas are determined by high resolution mass spectrometry (HREIMS) and structures are deduced from mass spectral fragmentation patterns and one- or two-dimensional nuclear magnetic resonance spectroscopy, like 1H NMR, 13C {1H} NMR, 13C PENDANT, COSY 90, HMQC, HMBC and
gp-NOESY. Structure assignments are then confirmed by chemical correlation and partial or
total synthesis. The absolute configuration of chiral compounds is finally determined by enantioselective GC (eGC) using authentic reference samples and modified cyclodextrins as stationary phase.
From more than 20 liverworts, screened for interesting compounds, 3 species were investigated in more detail: Fossombronia angulosa, Fossombroniaceae (Chapter 4.1., page 7), Riccardia chamedryfolia, Aneuraceae (Chapter 4.2., page 17), and Corsinia coriandrina, Corsiniaceae (Chapter 4.3., page 28). A variety of deuterium labelled precursors was prepared (Chapter 4.4., page 67) and axenic in vitro cultures of Corsinia coriandrina were established in cooperation with Dr. K. von Schwartzenberg at the Biozentrum Klein Flottbek, University of Hamburg, Germany (Chapter 4.5., page 76). Application experiments were performed in order to study the biosynthesis of 4-methoxystyrenes (Chapter 4.6.1., page 86), stilbenoids (Chapter 4.6.2., page 102) and terpenoids (Chapter 4.6.3., page 130) in Corsinia coriandrina. Research on microbial metabolites involves structure elucidation and synthesis of a sesquiterpene hydrocarbon emitted by the American elm, Ulmus americana, upon infection with the fungal pathogen Ophiostoma novo-ulmi (Chapter 4.7., page 137), performed in co-operation with Prof. Gerhard Gries (Burnaby, Canada), and a new class of octamethyl-tricyclo[3.2.1]octadienes emitted by the rhizobacterium Serratia odorifera (Chapter 4.8., page 149), which was performed in cooperation with Prof. Birgit Piechulla (Rostock, Germany).
3
3. General Part
»Man darf wohl sagen, daß der Chemie der ätherischen Öle auch bei den Kryptogamen noch ein weites Arbeitsfeld offensteht, ein Gebiet, das dem bei den höheren Pflanzen an Ausdehnung und Mannigfaltigkeit vielleicht nicht nachstehen wird.« Karl Müller, 1905
3.1. Liverwort Biology
The bryophytes are taxonomically placed between green algae (chlorophytes) and ferns (pteridophtes) and regarded as the closest living relatives of the first land plants. It is generally assumed that these organisms developed from Charophytean algae ancestors in Ordovician times approximately 500 million years ago (Kenrick & Crane, 1997; Wellman et al., 2003; Graham et al., 2004). Nevertheless, relationships among the three bryophyte classes (liverworts, mosses, hornworts) and between them and other embryophytes have remained unclear for several times (Beckert et al., 1999; Nickrent et al., 2000; Pruchner et al., 2001, 2002). Recent phylogenetic studies have placed the liverworts (Hepaticae, 6.000 – 8.000 species) as the most basal group of all terrestrial plants (Qiu et al., 2006, 2007). The life cycle of liverworts is dominated by haploid gametophytes which form the liverwort plant and develop gametangia for sexual reproduction. Male antheridia and female archegonia are either located on separate thalli (dioecious) or present on the same thallus (monoecious). Diploid sporophytes are only produced after water mediated fertilization and these short lived organs are nourished by the gametophyte. After meiosis the resulting haploid spores are released to germinate and develop new gametophytes. In addition, asexual propagation via gemma cups is known from species like Lunularia cruciata or Marchantia polymorpha.
On the cellular level one of the most striking differences between the cells of the majority of liverworts and those of all other terrestrial plants and algae is the presence of highly refractive structures commonly known as oil bodies (see Colour Plates 1e,3, and 5,pages 210 – 212). In thalloid Marchantialean liverworts oil bodies are restricted to scattered specialized cells, whereas in the leafy Jungermanniales they are generally present in all cells. Electron microscope studies revealed that liverwort oil bodies are confined by a single lipid membrane and contain lipid globules embedded in a carbohydrate matrix (Duckett & Ligrone, 1995).
4
The first extractions of liverworts performed by H. Lohmann (1903) and Karl Müller (1905) afforded essential oils in up to 1.5 % of the dry weight, which were assumed to consist of mono and sesquiterpenoids. Müller (1939) also concluded that these compounds were contained in the oil body structures, because their number correlated with the amount of oil
obtained. Furthermore, the bluish oil bodies of several Calypogeia species were attributed to 1,4-dimethylazulene (1) as the major volatile constituent (Katoh & Takeda, 1990), and 3-methoxybibenzyl (8) has been identified as the main volatile constituent of isolated oil bodies of Radula complanata (Flegel & Becker, 2000) (Figure 1, page 5). Histochemical localization of enzymes related to terpenoid biosynthesis in oil body membranes of
Marchantia polymorpha have shown that these unique organelles are not merely storage
vesicles but constitute metabolically reactive compartments (Suire et al., 2000).
Liverworts are rich in carbohydrates but almost devoid of any mechanical protection against herbivores, due to the imperfectly developed cuticula and the lack of lignins. Nevertheless, they are only rarely affected by herbivory, which led to the assumption that they should posses a kind of antifeedants. Antifungal, antibacterial, molluscicidal, and insecticidal, as well as plant growth inhibiting or promoting activities of liverwort constituents suggested that these compounds act as a chemical defence to become most efficient upon rupture of the plant tissue (Becker & Wurzel, 1987; Zinsmeister & Mues, 1987; Wurzel et al., 1990; Zinsmeister et al., 1991, 1994; Frahm, 2004; Asakawa, 2007, 2008).
In addition, liverworts form associations with proteobacteria and enterobacteria showing antifungal and antibacterial activity (Opelt & Berg, 2004), as well as symbiotic and free-living cyanobacteria (West & Adams 1997; Adams & Duggan, 2008; Rikkinen & Virtanen, 2008). Epiphytic methylobacteria promote liverwort growth by releasing cytokinins and auxin (Kutschera & Koopmann, 2005; Kutschera 2007). Furthermore, many liverworts form mycorrhiza-like associations with fungal symbionts of the Glomeromycota (Read et al., 2000; Russel & Bulman, 2005; Kottke & Nebel 2005; Ligrone et al., 2007; Kottke et al., 2008). Fossilized fungal hyphae and spores from the Ordovician strongly resemble those of current
Glomus species, suggesting that fungal associations might have facilitated colonization of
terrestrial habitats (Redecker et al., 2000; Heckman et al., 2001). The extent of these associations is best seen in the non-photosynthetic ghostwort, Cryptothallus mirabilis, which forms mycorrhizal associations with Tulasnella to obtain photosynthate from other autotrophs associated with the fungus (Bidartondo, et al., 2002, 2003).
5
3.2. Secondary Metabolites of Liverworts
Figure 1: Collection of secondary metabolites from liverworts (and hornworts (13)).
The liverworts are characterized by lunularic acid (5) an ubiquitous liverwort specific phytohormone first isolated from Lunularia cruciata (Valio et al., 1969), and later shown to be a precursor of liverwort bibenzyls (Pryce, 1972a) and bis(bibenzyls) (Friederich et al., 1999a, 1999b). Lunularic acid (5) and the corresponding decarboxylation product lunularin (6) (Pryce & Linton, 1974), as well as its labile precursor, prelunularic acid (7) (Ohta et al., 1983, 1984; Abe & Ohta, 1984, 1985), have been detected in all liverworts examined, but not in algae, mosses, hornworts, ferns, lichens, pteridophytes, or higher plants (Pryce, 1972b; Gorham 1977a, 1977b). The single exception is the Garden Hortensia, Hydrangea
macro-phylla (Saxifragales), which contains lunularic acid (5) and hydrangenic acid (194, Figure
6
In addition, flavonoids (10) of chalcone synthase (CHS) origin are abundant in liverworts (Asakawa, 1982, 1995, 2004), whereas isoflavonoids are extremely rare (Anhut et al., 1984). Furthermore, liverworts are known as rich sources of mono-, sesqui-, and diterpenoids (Asakawa, 1982, 1995, 2004). Some sesquiterpenoids with unique skeletons like anastreptene (2) (Andersen et al., 1978; von Reuß et al., 2004) and cyclomyltaylane (3) (Wu & Chang, 1992) have exclusively been reported from liverworts. In addition, many liverwort sesquiterpenoids like (–)-longifolene (4) exhibit the opposite absolute configuration in comparison to higher plant compounds (Huneck & Klein, 1967; Asakawa, 1982). These ent-sesquiterpenoids and their derivatives from functional group transformations and rearrangement reactions have become a valuable source of reference compounds required for comparative enantioselective GC analysis using modified cyclodextrins as stationary phase (Fricke et al., 1995; König et al., 1996; Bülow & König, 2000, König & Hochmuth, 2004). In contrast to vascular plants and aquatic algae, nitrogen- or sulfur-containing constituents are virtually unknown from the liverworts (Asakawa, 2004, 1995). Considering the vast diversity, distribution, and ecological importance of terrestrial plant alkaloids (Hesse, 2000), their scarcity in lower plants is remarkable. Only 7- and 6-prenylindoles (14, 15) were described from the Metzgerialean liverworts Riccardia chamedryfolia, R. incurvata, and R. multifida, Aneuraceae (Benesova et al., 1969a, 1969b; Huneck et al., 1972; Nagashima et al., 1993), whereas the alkaloid anthocerodiazonin (13) was reported from a hornwort, Anthoceros
agrestis (Trennheuser et al., 1994). Sulfur containing constituents are equally rare. The
isotachins A – C with -S-methylthio-(E)-acrylate structures (16) were obtained from the Jungermannialean liverworts Isotachis japonica (Asakawa et al., 1985), Balantiopsis rosea (Asakawa et al., 1986) and B. cancellata (Labbé et al., 2005). Some chlorinated bibenzyls (9) and bis(bibenzyls) have recently been described from Lepidozia (Scher et al., 2003),
Plagiochila (Anton et al., 1997), Bazzania (Martini et al., 1998; Speicher et al., 2001), Herbertus, Mastigophora (Hashimoto et al., 2000), and Riccardia species (Baeck et al., 2004;
7
4. Special Part
4.1. C
11Hydrocarbons from Fossombronia angulosa
4.1.1. Fossombronia angulosa
Fossombronia angulosa (Dicks.) Raddi is a member of the Fossombroniaceae, which form
thallose gametophytes with lamina divided into leaf-like lobes and a stem-like costa (Plate 1a, page 210) carrying gametangia and sporophytes (Plate 1b). About 50 species have been described of which 11 are represented in Europe, but some of these are difficult to distinguish in the absence of gametangia and spores (Paton, 1999). Fossombronia angulosa forms thalli up to 2.5 cm long and is restricted to lowland coastal regions with a mild climate. Samples were collected on the Islands of Tenerife and Madeira in April 2003 and March 2007, respectively. While former collection was lacking mature sporogons, the identity of the latter specimen could be unambiguously established by inspection of spore morphology (Plate 1c), which exhibited characteristic lamellae appearing on the spore margin as a continuous pale wing (Plate 1d) as described by Paton (1999). Most thallus cells contain 6 – 10 oil bodies with a diameter of 1 – 6 µm (Plate 1e).
Members of the Fossombroniaceae have previously been investigated for secondary metabolites. Antibacterial diterpene dialdehydes were reported from in vitro cultured
Fossombronia pusilla (Sauerwein & Becker, 1990). Furthermore, biosynthesis of geosmin
(39) in F. pusilla has been shown to proceed via the mevalonate (MVA) pathway, in contrast to its methylerythritol-4-phosphate (MEP) origin in Streptomces species (Spiteller et al., 2002), and leucine origin in Myxobacteria (Dickschat et al., 2005). Hopane-type triterpenoids and epi-neoverrucosane-type diterpenoids have been described from in vitro cultures of the rare F. alaskana (Grammes et al., 1994, 1997) and their biosynthesis via the MVA or MEP pathways, respectively, was established (Hertewich et al., 2001).
8
4.1.2. Volatile Constituents of Fossombronia angulosa
Figure 2: Gas chromatograms of hydrodistillation products of Fossombronia angulosa collected
on the Islands of Tenerife and Madeira (30 m, CpSil-5 CB, 50 °C, + 3 °C /min, to 250 °C).
Hydrodistillation of the carefully cleaned fresh plant material afforded an essential oil, which was analyzed by GC (Figure 2) and GC-EIMS. Undecan-2-one (34) or tridecan-2-one (35) was identified as major constituent by comparison of mass spectra and GC retention indices with a spectral library established under identical experimental conditions (König et al., 2004; König & Hochmuth, 2004). Small amounts of the corresponding alcohols (37 and 38) were also detected, along with pentadecan2one (36). Monoterpene hydrocarbons like (–) -sabinene (20) and -phellandrene (26) were identified as major constituents of the specimen from Tenerife only.
Madeira Tenerife 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 37 39 35 29 30 31 32 39 34 38 36 35 10 20 30 40 RT [min] 20 21 26 29 dictyopterene A 30 ectocarpene 31 pentylbenzene 32 dictyotene 33 terpinen-4-ol 34 undecan-2-one 35 tridecan-2-one 36 pentadecan-2-one 37 undecan-2-ol 38 tridecan-2-ol 39 geosmine 17 -thujene 18 -pinene 19 camphene 20 -sabinene 21 -pinene 22 myrcene 23 3-carene 24 -terpinene 25 p-cymene 26 -phellandrene 27 -terpinene 28 terpinolene
9
Three unidentified hydrocarbons with molecular ion signals at m/z 148 [M] (C11H16, 30) or
m/z 150 [M] (C11H18, 29 and 32) were isolated by a combination of column chromatography
on silica gel and semi-preparative GC. The identification of dictyopterene A (29), ectocarpene (30), dictyotene (32), and n-pentylbenzene (31) is described in the following sections.
Figure 3: Volatile constituents identified in the hydrodistillates of Fossombronia angulosa
10
Figure 4: Mass spectra (EI, 70 eV) of dictyopterene A (29), dictyotene (32),
ectocarpene (30), and n-pentylbenzene (31) from Fossombronia angulosa.
41 53 67 79 91 107 121 150 40 60 80 100 120 140 160 20 40 60 80 100 41 53 67 79 93 107 121 150 40 60 80 100 120 140 160 20 40 60 80 100 39 51 66 79 91 105 119 133 148 40 60 80 100 120 140 160 20 40 60 80 100 41 51 57 65 78 91 105 119 148 40 60 80 100 120 140 160 20 40 60 80 100 C11H18 C11H18 C11H16 C11H16 C7H7
11
4.1.3. Identification of Dictyopterene A (29)
The mass spectrum of 29 (Figure 4) exhibited a molecular ion signal at m/z 150 [M] indicating the molecular formula C11H18 with 3 units of unsaturation. The 1H NMR spectrum
(Figure 5) exhibited signals for one vinyl moiety at H 4.88 (1H, dd, 3JZ = 10.4 Hz, 2J = 1.6 Hz), 5.05 (1H, dd, 3JE = 17.0 Hz, 2J = 1.6 Hz) and 5.31 (1H, ddd, 3JE = 17.0 Hz, 3JZ = 10.4 Hz, 3J = 8.2 Hz), one trans-configured ethylene bond at H 4.97 (1H, dd, 3JE = 15.5 Hz, 3J = 7.6 Hz) and 5.46 (1H, dt, 3JE = 15.4 Hz, 3J = 6.6 Hz) adjacent to an allylic methylene group at
H 1.96 (2H, dt, 3J = 6.6 Hz), one overlapping signal at H 1.22 - 1.36(6H, m), one methyl
group at H 0.86 (3H, t, 3J = 7.3 Hz), and a methylene group as part of a cyclopropyl moiety
as indicated by the chemical shift of H 0.67 (2H, t, 3J = 6.9 Hz).
Figure 5: 500 MHz 1H NMR spectrum of (+)-dictyopterene A (29, in C6D6) from F. angulosa.
Inspection of the 13C PENDANT (Figure 6) and HMQC spectra confirmed the presence of a vinyl moiety at C 112.0 (t) and 141.0 (d), one internal double bond at C 129.1 (d) and
132.1 (d), one methyl group at C 14.2 (q), one 1,2-disubstituted cyclopropyl-unit at C 14.9
(t), 23.9 (d), and 24.6 (d), and three methylene groups at C 22.6 (t), 32.2 (t), and 32.6 (t). The
connectivities between the partial structures were deduced from two dimensional COSY and HMBC spectra, which indicated a 1-(hex-1-enyl)-2-vinylcyclopropane skeleton, previously described as dictyopterene A (29) from marine brown algae (Pohnert & Boland, 2002). The
trans-configuration of the cyclopropyl moiety in 29 was assigned according to the gp-NOESY
spectrum, which exhibited NOE-correlations between 3-H and 6-H, as well as 2-H and 5-H. This hypothesis was confirmed by comparison of the mass spectrum and GC retention index
H [ppm] 0.8 1.2 1.6 2.0 2.4 2.8 3.2 3.6 4.0 4.4 4.8 5.2 11-CH3 9,10-CH2 8-CH2 2-CH 3,5-CH 1-CHZ 1-CHE 6-CH 4-CH2 7-CH
12
with those of an authentic sample of (3R,5R)-(+)-dictyopterene A (29). The enantiomeric composition of 29 was determined upon enantioselective GC (Section 4.1.7., page 15).
Figure 6: 100 MHz 13C PENDANT spectrum of (+)-dictyopterene A (29, in C6D6) from F. angulosa.
4.1.4. Identification of Dictyotene (32)
The molecular ion signal at m/z 150 [M] in the mass spectrum of compound 32 (Figure 4, page 10) suggested the molecular formula of C11H18 with 3 units of unsaturation. The 1H
NMR spectrum (Figure 7, page 13) indicated four partially overlapping signals for olefinic protons at H 5.73 (1H, m) and 5.61 (3H, m), thus, suggesting a monocyclic structure.
Furthermore, one anisochoric methylene group at H 2.58 (1H, d.br, 2J = 19.5 Hz) and 2.86
(1H, d.br, 2J = 19.2 Hz), a chiral methine group at H 2.43 (1H, s.br.), one allylic methylene
group at H 2.13 (2H, m, 6-H), one propylene bridge at H 1.24 (6H, s.br), and one methyl
group at H 0.87 (3H, t, J = 7.3 Hz) were identified. Inspection of the COSY spectrum
allowed the assignment of a consecutive 7-butylcyclohepta-1,4-diene structure, previously described as dictyotene (32) from marine brown algae (Pohnert & Boland, 2002). This hypothesis was confirmed by comparison of the mass spectra and GC retention indices with corresponding data of an authentic reference sample, whilst the enantiomeric composition of dictyotene (32) was determined upon enantioselective GC (Section 4.1.7., page 15).
TMS C[ppm] 20 30 40 50 60 70 80 90 100 110 120 130 140 1-CH2 4-CH2 11-CH3 2-CH 7-CH 6-CH 9,8-CH2 10-CH2 3,5-CH solvent TMS C[ppm] 20 30 40 50 60 70 80 90 100 110 120 130 140 1-CH2 4-CH2 11-CH3 2-CH 7-CH 6-CH 9,8-CH2 10-CH2 3,5-CH solvent
13
Figure 7: 500 MHz 1H NMR spectra of dictyotene (32) and ectocarpene (30, in C6D6)
from Fossombronia angulosa.
4.1.5. Identification of Ectocarpene (30)
The molecular ion signal at m/z 148 [M] in the mass spectrum of compound 30 suggested the molecular formula of C11H16 with 4 units of unsaturation (Figure 4, page 10). The 1H NMR
spectrum of 30 was similar to that of dictyotene (32) (Figure 7), exhibiting signals for a
1.2 1.6 2.0 2.4 2.8 3.2 3.6 4.0 4.4 4.8 5.2 5.6 5-CH 1,4-CH 2-CH 8-CH 9-CH 3-CH 7-CH 3-CH’ 6-CH2 10-CH2 11-CH3 [ppm] H [ppm] 1.2 1.6 2.0 2.4 2.8 3.2 3.6 4.0 4.4 4.8 5.2 5.6 5-CH 3-CH’ H 3-CH 7-CH 8-CH2 1,2,4-CH 6-CH2 9,10-CH2 11-CH3
14
cyclohepta-1,4-diene-7-yl skeleton. Nevertheless, signals for the alkyl chain of dictyotene (32) were replaced by those corresponding to a (Z)-configured internal double bond at H 5.48
(1H, dd, J = 10.7 Hz, JZ = 9.5 Hz) and 5.33 (1H, dt, JZ = 9.8, J = 7.3 Hz), adjacent to an allylic methylene group at H 1.97 (2H, dq, J = 7.3 Hz, J = 7.6 Hz). The location of the double
bond in 30 was deduced from the chemical shift and triplet multiplicity of the methyl signal at
H 0.87 (3H, t, J = 7.6 Hz) and the fact that the chiral methine group at H 3.52 (1H, s.br) was
significantly shifted to lower field in comparison to the saturated dictyotene (32). The structure of ectocarpene (30) was established by comparison of the mass spectra and GC retention indices with an authentic sample while the enantiomeric composition was determined upon enantioselective GC (Section 4.1.7., page 15).
4.1.6. Identification and Synthesis of n-Pentylbenzene (31)
GC-EIMS analysis of the hydrocarbon fraction of F. angulosa suggested the presence of trace amounts (< 0.1 % of the total volatiles) of another C11H16 compound (31), as indicated by the
molecular ion signal at m/z 148 [M] (Figure 4, page 10). The base peak at m/z 91 (100) [M – C4H9] for a tropylium ion (C7H7+) suggested a pentylbenzene structure. Considering the
co-occurring C11 hydrocarbons (29, 30 and 32) an unbranched alkyl residue was assumed.
This hypothesis was confirmed by comparison of the mass spectra and GC retention indices with n-pentylbenzene (31) prepared as shown in figure 8. Racemic (±)-1-phenylpentanol (41) was obtained in 79 % yield from benzaldehyde (40) upon reaction with n-butyl lithium. Acid catalyzed dehydration was carried out using ion exchange resin Amberlyst® 15 in dichloro-methane to give (E)-1-phenyl-1-pentene (42) in 66 % yield, which was hydrogenated using palladium on charcoal to give n-pentylbenzene (31) in 90 % yield identical to the natural product from Fossombronia angulosa.
15
4.1.7. Enantioselective GC Analysis of C11 Hydrocarbons
The determination of absolute configurations and enantiomeric excess of dictyopterene A (29) using gaschromatography with modified cyclodextrins as chiral selectors has previously been described, but no suitable cyclodextrin derivative for the resolution of (±)-ectocarpene (30) or (±)-dictyotene (32) was available at that time (Boland et al., 1989; Wirth et al., 1992). Of all the different phases developed and tested till 2004 only heptakis(2,3-di-O-acetyl-6-O-tert-butyldimethylsilyl)- -cyclodextrin (2,3-Ac-6-TBDMS- -CD) separated (±)-30 and (±)-32 (König, 2004). Using 2,3-Ac-6-TBDMS- -CD as the stationary phase at 60 °C isothermally, the following -values were obtained; ectocarpene (30): (S)-(+)/(R)-(–) = 1.026, dictyotene (32):
(R)-(–)/(S)-(+) = 1.029, and dictyopterene A (29): (3S,5S)-(–)/(3R,5R)-(+) = 1.030 (Figure 9).
Figure 9: Enantioselective GC analysis of C11 hydrocarbons from Fossombronia angulosa
(29, 30 and 32) using 2,3-Ac-6-TBDMS- -CD at 60 °C isothermally.
(3R,5R)-(+)-dictyopterene A (29) from F. angulosa exhibited an enantiomeric excess of
ee = 72 and 74 %. The enantiomeric purities of (R)-(–)-dictyotene (32, ee = 87 %) and
(S)-(+)-ectocarpene (30, ee = 64 %) were high in samples from Tenerife, whereas almost racemic mixtures were observed in samples from Madeira with (R)-(–)-dictyotene (32,
ee = 32 %) and (R)-(–)-ectocarpene (30, ee = 14 %) predominating.
RT [min] 20 18 16 (–) (+) (–) (+) (–) (+) (+)-30 ( )-32 racemic standards (–)-30 Tenerife Madeira ( )-32 (+)-29 (+)-29
16
4.1.8. Discussion of C11 Hydrocarbons from Fossombronia angulosa
Relying on GC-EIMS measurements and mass spectral databases Asakawa et al. have recently reported the identification of dictyopterene (29), dictyotene (32), and (Z)-multifidene (43) from F. angulosa (Ludwiczuk et al., 2008). The presence of 29 and 32 is now confirmed by NMR spectroscopy. Because mass spectra and GC retention indices of (Z)-multifidene (43, RI 1140) and (Z)-ectocarpene (30, RI 1136) are very similar (König et al., 2004) identification of multifidene (43) might be in error as indicated by the characterization of ectocarpene (30) using 1H NMR techniques. In addition, n-pentylbenzene (31) was identified as a trace constituent by comparison with a synthetic sample. Furthermore, enantiomeric compositions of ectocarpene (30) and dictyotene (32) are reported for the first time and shown to be highly variable.
Figure 10: C11 hydrocarbons identified in Fossombronia angulosa (29– 32).
Compounds 29, 30 and 32 as well as related C11 hydrocarbons are well known from marine
brown algae (Phaeophyceae), which release enantiomeric mixtures as pheromones of female gametes and in chemical defence (Pohnert & Boland, 2002). Ectocarpene (30) has also been reported from Senecio isatideus, Asteraceae (Bohlmann et al., 1979). Dodecatrienoic acid was recognized as its biogenetic precursor in this higher plant (Boland & Mertes, 1985; Neumann & Boland, 1990), whereas C11-hydrocarbons of brown algae originate from eicosapentaenoic
acid (C11H16) or arachidonic acid (C11H18) (Stratmann et al., 1993; Pohnert & Boland, 2002).
While the biological function and biogenetic origin of C11 hydrocarbons in Fossombronia
angulosa remains to be clarified, the occurrence of arachidonic acid and eicosapentaenoic
acid in some liverworts (Asakawa, 1995) suggests a biogenetic pathway similar to those in marine brown algae, although there are no close phylogenetical relationships between these taxa.
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4.2. Indole Alkaloids from Riccardia chamedryfolia
4.2.1. Riccardia chamedryfolia
The genus Riccardia S. Gray 1821 belongs to the family Aneuraceae, Metzgeriales. Riccardia
chamedryfolia (With.) Grolle 1969 forms dark green, 1 – 2 mm wide and 10 – 30 mm long
thalli consisting of 5 – 8 cell layers (Plate 2, page 211). Almost all epidermal cells contain 1 – 2 spherical, ellipsoidal, brownish oil bodies which range in size from 7 – 15 x 9 – 25 µm, each composed of numerous small spherules (Plate 3,page 211).
As early as 1969 Šorm and co-workers reported on the isolation of 7-prenylindole (14) and 6-prenylindole (15) from Riccardia incurvata and R. sinuata (Hook.) Trev., a synonym for
R. chamedryfolia (Benesova et al., 1969a, 1969b). These results were confirmed by Huneck,
who reported local anesthetic properties of 15 and mentioned the presence of another unidentified polar alkaloid in R. chamedryfolia (Huneck et al., 1972). 6- and 7-prenylindoles (15 and 14) were also detected in R. multifida (L.) S. Gray subsp. decrescens (Steph.) Furuki, (Asakawa et al., 1983; Nagashima et al., 1993). Furthermore, prenylindoles 14 and 15, as well as the corresponding 3- and 5-isomers have been described from higher plants, such as Annonaceae (Nwaji et al., 1972; Achenbach & Renner, 1985; Achenbach, 1986; Muhammad et al., 1986; Nkunya et al., 2004; Boti et al., 2005) or Rutaceae (Delle Monache et al., 1989; Kinoshita et al., 1989). Antifungal activities of 6-prenylindole (15), which has also been described from Streptomyces sp. TP-A0595, Actinomycetes, suggested its role in chemical defence (Sasaki et al., 2002).
4.2.2. Volatile Constituents of Riccardia chamedryfolia
Plant material of Riccardia chamedryfolia (With.) Grolle was collected on the Island of La Palma in January 2006 and in Great Britain in 2007. The carefully cleaned fresh plant material was frozen with liquid nitrogen, crushed and extracted with organic solvents using sodium sulfate as drying reagent, or hydrodistilled, or air dried and solvent extracted. Comparative GC-EIMS analysis indicated significant amounts of oxygenated artifacts in the hydrodistillation products and diethyl ether extracts of air dried material, which were much less pronounced in pentane extracts of fresh plants. GC and GC-EIMS investigations indicated 7-prenylindole (14, 56 % of the total volatiles) and 6-prenylindole (15, 19 %) as major constituents (Figure 11). In addition, sesquiterpene hydrocarbons like -maaliene (44, 13 %), calarene (45, 1 %), -acoradiene (46, 1 %), bicyclogermacrene (47, 5 %), and
18
-cuprenene (48, 1 %) (Figure 12) were identified by comparison of mass spectra and GC retention indices with a spectral library established under identical experimental conditions (König et al., 2004). Extracts were fractionated by column chromatography on silica gel using a hexane – diethyl ether gradient. Prenylindoles 14 and 15 were isolated upon repeated column chromatography on silica gel 60 using a hexane – dichloromethane gradient.
Figure 11: TIC chromatogram of pentane / NaSO4 extract of fresh Riccardia chamedryfolia
from La Palma (30 m CpSil-5 CB, 80 °C, 2 min, + 10 °C/min, to 270 °C).
Figure 12: Sesquiterpene hydrocarbons (44 – 48) and prenylindoles (14, 15, 49 – 52)
from Riccardia chamedryfolia.
10 12 14 16 RT [min] 15 49 50 51 44 47 48 45 46 Sesquiterpenes Prenylindoles 50 TIC [%] 14 10 20 30 40
19
Figure 13: Mass spectra (EI, 70 eV) of 7-prenylindole (14), 6-prenylindole (15),
3-chloro-7-prenylindole (49) and 3-chloro-6-prenylindole (50). 39 51 63 77 83 89 103 117 130 143 155 170 185 40 60 80 100 120 140 160 180 200 220 20 40 60 80 100 39 51 63 77 84 89 103 117 130 143 155 170 185 40 60 80 100 120 140 160 180 200 220 20 40 60 80 100 41 51 63 77 84 101 115 128 141 154 164 169 188 204 219 40 60 80 100 120 140 160 180 200 220 20 40 60 80 100 41 55 63 69 77 84 95 102 115 128 141 154 169 177 184 204 219 40 60 80 100 120 140 160 180 200 220 20 40 60 80 100 C13H15N●+ C13H15N●+ C9H8N+ C12H11N●+ C13H14NCl●+ C13H14NCl●+ C12H11NCl+ C12H11N●+ C12H11NCl+ C9H7NCl+ C12H12N●+ C12H12N●+
20
4.2.3. Identification and Synthesis of 3-Chloro-6- and 7-prenylindoles (49, 50)
Two chlorine containing indole alkaloids with molecular ion signals at m/z 219 [M] were detected as minor (49) or trace constituents (50). The relative intensities of the [M + 2] signals at m/z 221 indicated chlorine as a substituent (Figure 13). The molecular formula C13H14NCl
with seven units of unsaturation was established upon HREIMS. The mass spectrometric fragmentation pattern of 49 was similar to those of 7-prenylindole (14) and exhibited dominant chloroazaazulenium ion signals at m/z 204 [M – CH3] (100) for C12H11NCl+ and
m/z 164 [M – C4H7] (90) for C9H7NCl+, thus, indicating a prenylindole structure with an
aromatic chlorine substitution. The corresponding signals at m/z 204 [M – CH3] (60) and
m/z 164 [M – C4H7] (15) were less intense for the later eluting trace constituent 50, assumed
to represent the 6-prenyl derivative. Its mass spectrum was dominated by a base peak signal at
m/z 169 [M – CH3 – Cl] (100) for C12H11N●+, which was significantly less pronounced for the
7-prenyl indole derivative (49) showing m/z 169 (50). Although rearranged chloroaza-azulenium ions are almost identical for 6- and 7-prenylindoles, the observation that m/z 169 [M – CH3 – Cl] is predominating in the 6-prenyl isomer (50) only, suggested that different
charge distribution in the initial [M – CH3] fragment ion facilitates loss of the chlorine radical.
Quantum mechanical calculations (PM3, RHF) and Mulliken population analysis indicated that the 6-butenylindol fragment ion [M – CH3] exhibits characteristic partial positive charges
located at 2-C, 3a-C, and 5-C of the indole nucleus, thus, suggesting the adjacent 3- (or 4-) position for the chloro-substitution. This hypothesis was unambiguously established by comparison with authentic samples of 7- and 6-prenyl-3-chloroindoles (49 and 50) obtained by partial synthesis (Figure 14). Regioselective chlorination of indole (53) to 3-chloroindole (54) has previously been described (De Rosa & Alonso, 1978). Reaction of 7- or 6-prenyl-indole (14 and 15), isolated from Riccardia chamedryfolia, with N-chlorosuccinimide in dichloromethane afforded the corresponding 3-chloro derivatives (49 and 50) in 85 and 80 % yield, respectively.
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Figure 15: 500 MHz 1H NMR spectra of 7-prenylindole (14) and 3-chloro-7-prenylindole (49).
The 3-chloro substitution was established by 1H NMR spectroscopy (Figures 15 and 16). Comparison of the mass spectra and GC retention indices confirmed the identity of 49 and 50 with the natural products from Riccardia chamedryfolia.
2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 7.5 H [ppm] 12-CH3 11-CH3 8-CH2 9-CH 2-CH NH 6-CH 5-CH solvent 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 H [ppm] solvent 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 11-CH3 12-CH3 8-CH2 9-CH 3-CH 2-CH 6-CH NH 5-CH 4-CH X 4-CH
22
Figure 16: 500 MHz 1H NMR spectra of 6-prenylindole (15) and 3-chloro-6-prenylindole (50).
2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 7.5 H [ppm] 11-CH3 12-CH3 8-CH2 9-CH 2-CH 6-CH NH 5-CH 4-CH solvent X 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 7.5 H [ppm] 11-CH3 12-CH3 8-CH2 9-CH 3-CH 2-CH 7-CH NH 5-CH 4-CH solvent X X X
23
4.2.4. Identification of Chamedryfolian (51)
One new oxazinoindole alkaloid (51) named chamedryfolian, with a molecular ion signal at
m/z 199 [M] (Figure 17) was isolated as a minor constituent by a combination of column
chromatography and thin layer chromatography on silica gel. The molecular formula C13H13NO with eight units of unsaturation was established by GC-HREIMS. Keeping
structures of already identified indoles in mind and due to the base peak for an azaazulenium ion C9H8N+ at m/z 130 [M – C4H5O] (100) the structure of a prenylindole carrying an oxygen
in the side chain was postulated.
Figure 17: Mass spectrum (EI, 70 eV) of chamedryfolian (51) from Riccardia chamedryfolia.
The 1H NMR and COSY spectra exhibited signals for a 7-substituted indole nucleus, along with an oxygenated isopentenyl residue (Table 1, page 24), which, together with eight units of unsaturation for the molecular formula C13H13NO implied a tricyclic indole alkaloid. 1H NMR
signals assigned to the sidechain pointed to one isopropenyl moiety and one anisochoric benzylic methylene group at H 2.73 (1H, dd, 2J = 14.8 Hz, 3J = 3.8 Hz) and H 2.92 (1H, dd,
2
J = 14.8 Hz, 3J = 7.9 Hz), adjacent to an oxygen linked methine group at H 4.38 (1H, dd,
3
J = 7.9 Hz, 3J = 3.8 Hz). The large difference in vicinal coupling constants of the anisochoric
methylene hydrogens (3J = 7.9 Hz vs. 3.8 Hz) indicated their inclusion in a rigid structure and
suggested an oxacyclic bridge between the chiral methine group and the indole nitrogen (Figure 18). 41 51 69 77 103 130 156 171 184 199 40 60 80 100 120 140 160 180 200 20 40 60 80 100 [M] C13H13NO C9H8N+
24
Figure 18: Important H,H-coupling constants, H,H-COSY and H,C-HMBC correlations
in chamedryfolian (51) from Riccardia chamedryfolia.
1 H NMR 14 15 49 50 51 1-NH 7.25 1H s.br 6.77 1H s.br 6.91 1H s.br 6.38 1H s.br. 2 6.65 1H s.br 6.59 1H s.br 6.53 1H s.br 6.44 1H s.br. 6.72 1H s.br 3 6.56 1H s.br 6.49 1H s.br 6.55 1H s.br 4 7.63 1H d 7.2 7.64 1H d 8.2 7.71 1H d 8.2 7.72 1H d 7.9 7.64 1H d 7.9 5 7.19 1H dd 7.2 7.3 7.09 1H d 7.9 7.12 1H dd 7.9 7.3 7.03 1H d 7.6 7.13 1H dd 7.9 7.3 6 7.08 1H d 7.3 7.01 1H d 7.3 6.92 1H d 7.3 7 7.00 1H s 6.83 1H s 8 3.32 2H d 6.9 3.53 2H d 7.3 3.18 2H d 6.9 3.43 2H d 7.3 2.73 1H dd 14.8 3.8 2.92 1H dd 14.8 7.9 9 5.34 1H m 5.55 1H m 5.24 1H t 6.9 5.46 1H t 7.2 4.38 1H dd 7.9 3.8 11 1.57 3H s 1.67 3H s 1.53 3H s 1.63 3H s 4.79 1H s 4.82 1H s 12 1.61 3H s 1.72 3H s 1.61 3H s 1.70 3H s 1.58 3H s
Table 1: 1H NMR data of 7-prenylindole (14), 6-prenylindole (15), 3-chloro-7-prenylindole (49), 3-chloro-6-prenylindole (50), and chamedryfolian (51) from Riccardia chamedryfolia (chemical shift
H [ppm] in C6D6, integral, multiplicity, and coupling constants [Hz]; assignments derived from 1
