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The quinonoid pigments of the lichens nephroma laevigatum and heterodermia obscurata Cohen, Peter Altman 1995

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THE QUINONOID PIGMENTS OF THE LICHENS NEPHROMA LAEVIGATUMAND HETERODERMIA OBSCURATAbyPETER ALTMAN COHENB.G.S., The University of Maryland (College Park), 1984M.Sc., The University of Washington (Seattle), 1989A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinTHE FACULTY OF GRADUATE STUDIES(Department of Botany)We accept this thesis as conformingto the required standardNTHE UNIVERSITY OF BRITISH COLUMBIAJune 1995© Peter Altman Cohen, 1995In presenting this thesis in partial fulfilment of the requirements for an advanceddegree at the University of British Columbia, I agree that the Library shall make itfreely available for reference and study. I further agree that permission for extensivecopying of this thesis for scholarly purposes may be granted by the head of mydepartment or by his or her representatives. It is understood that copying orpublication of this thesis for financial gain shall not be allowed without my writtenpermission.(Signature)__________________________Department of )O74A) VThe University of British ColumbiaVancouver, CanadaDate___________DE-6 (2/88)ABSTRACTField samples of the foliose lichens Nephroma Iaevigatum Ach. and Heterodermia obscurata (NyL) Trevis. were analyzed for anthraquinone and anthraquinonelike pigments. Both lichens were found to contain emodin, 7-chioroemodin and 7,7’-dichiorohypericin. In addition, the N. Iaevigatum specimen contained 7-chloro-1 -0-methylemodin, 7-chloro- 1- 0-methyl-o-hydroxyemodin (7-ch lorocarviolin) and 2,2’,7,7’-tetrachlorohypericin, while the H. obscurata sample contained 5 ,7-dichloroemo-din, flavoobscurin A and flavoobscurin B. Laboratory incubation studies with N. Iaevigatum (in the presence of sodium [1-3C]acetate) also revealed the formation of 5-chloroemodin, 5-chloro- 1- 0-methylemodin, 5-chloro-o-hydroxyemodin and 5-chlo-ro- 1- 0-methyl-o-hydroxyemodin (5-chiorocarviolin). These compounds had notbeen identified in any previous examination of field-collected lichen, and 5-chloro-1 -0-methylemodin, although known from a fungus, has not been reported to occur inany lichen. The structures of the compounds were determined by a combination ofUV, MS, 1H NMR and ‘3C NMR spectral methods.Feeding experiments with sodium [2-14C]acetate and sodium [1-13C]acetatedemonstrated that anthraquinones are biosynthesized in Nephroma Iaevigatumthrough the polyketide pathway, analogous to the pathways operating in fungi andhigher plants. The lichen was also capable of chlorinating endogenous anthraquinones during incubation with sodium36chloride.IIIBiohalogenation experiments with a partially purified lichen chioroperoxidase and a commercially available fungal chloroperoxidase demonstrated enzyme-substrate specificity with respect to the production of different chlorinated anthraquinones. 5,7-Dichloroemodin was produced, in excellent yield, from 7-chioroemodinby the commercial enzyme; the lichen chloroperoxidase, however, yielded only 7-chloroemodin from emodin and did not further chlorinate 7-chloroemodin. 5-Chloro-emodin was a product from both the control and commercial enzyme reactions.Twelve lichen anthraquinones, bianthrones, hypericin derivatives, and synthetic hypericin were tested for their virucidal activity in end-point CPE (viral cytopathic effects) and plaque assays with herpes simplex virus type 1 (HSV-1). Emodin, 7-chloroemodin, 7-chloro- 1-O-methylemodin, 5,7-dichloroemodin, hypericinand 7,7’-dichlorohypericin exhibited fair to good antiviral activity in the presenceof light. In the plaque assay, 5,7-dichioroemodin, hypericin and 7,7’-dichlorohy-pericin completely inhibited the virus at a concentration of 1.0 pg/mI. Only hypericin was active at 0.01 pg/mI. The other anthraquinones and bianthroneswere inactive at a concentration of 5.0 pg/mI.ivTABLE OF CONTENTSABSTRACT iiTABLE OF CONTENTS ivLIST OF TABLES viiLIST OF FIGURES viiiACKNOWLEDGEMENTS ixFOREWORD XCHAPTER 1: OVERVIEW1.1 LICHEN CHEMISTRY AND CHEMOTAXONOMY 11.2 PHYTOCHEMISTRY OF ANTHRAQUINONES 191.3 CHEMISTRY OF NEPHROMA AND HETERODERMIA 27CHAPTER 2: CHEMICAL PROPERTIES OF LICHEN QUINONOID PIGMENTS2.1 ISOLATION OF NEPHROMA LAEVIGATUM NATURALPRODUCTS2.1.1 INTRODUCTION 362.1.2 MATERIALS AND METHODS 362.1.3 RESULTSANDDISCUSSION 412.1.4 SYNTHESES OF 1-O-METHYLEMODIN, 7,7’-DI-CHLOROHYPERICIN AND 2,2’,7,7’-TETRACHLOROHY-PERICIN 492.1.5 PHYSICAL AND CHEMICAL DATA FOR NATURALPRODUCTS 502.2 ISOLATION OF HETERODERMIA OBSCURATA NATURAL PRODUCTS2.2.1 INTRODUCTION 532.2.2 MATERIALS AND METHODS 552.2.3 RESULTS AND DISCUSSION 582.2.4 SYNTHESES OF 7-CHLOROEMODIN AND 5,7-DI-CHLOROEMODIN 632.2.5 PHYSICAL AND CHEMICAL DATA FOR NATURALPRODUCTS 64VCHAPTER 3: BIOSYNTHETIC STUDIES OF QUINONOID PIGMENTS INN. LAEVIGATUM3.1 INTRODUCTION 663.2 MATERIALS AND METHODS 663.3 RESULTS AND DISCUSSION 713.4 PHYSICAL AND CHEMICAL DATA FOR NATURALPRODUCTS 763.4.1 PRODUCTS FROM SODIUM[2-14CjACETATEINCORPORATION 763.4.2 PRODUCTS FROM SODIUM [1-13C]ACETATEINCORPORATION 773.4.3 PRODUCTS FROM SODIUM CHLORIDEINCORPORATION 78CHAPTER 4: BIOHALOGENATION STUDIES OF QUINONOID PIGMENTS4.1 INTRODUCTION 814.2 MATERIALS AND METHODS 824.3 RESULTS AND DISCUSSION 874.4 PHYSICAL AND CHEMICAL DATA FOR REACTIONPRODUCTS 894.4.1 PRODUCTS FROM CONTROL (NO ENZYME)REACTION 894.4.2 PRODUCTS FROM COMMERCIAL CHLOROPEROXIDASE REACTION 894.4.3 PRODUCTS FROM N. LAEVIGATUM CHLOROPEROXIDASE REACTION 90CHAPTER 5: ANTIVIRAL ACTIVITIES OF LICHEN QUINONOID PIGMENTS5.1 INTRODUCTION 925.2 MATERIALS AND METHODS 925.3 RESULTS AND DISCUSSION 98CHAPTER 6: CONCLUSIONS 103BIBLIOGRAPHY 116APPENDIX 1: APPROXIMATE PHYTOGEOGRAPHICAL DISTRIBUTIONOF NEPHRQMA LAEVIGATUM IN NORTH AMERICA 140APPENDIX 2: APPROXIMATE PHYTOGEOGRAPHICAL DISTRIBUTIONOF HETERODERMIA OBSCURATA IN NORTH AMERICA 141viAPPENDIX 3: NUMBERING SYSTEMS IN QUINONOID NATURAL PRODUCTS 142APPENDIX 4: CALCULATIONS OF 3C CARBON CHEMICAL SHIFTSUSING THE METHOD OF EWING (1979) 143APPENDIX 5: CALCULATIONS OF PROTEIN CONTENT AND SPECIFICACTIVITY OF SEMI-PURiFIED CHLOROPEROXIDASEFROM NEPHRQMA LAEV!GATUM 144APPENDIX 6: UV SPECTRUM OF SEMI-PURIFIED CHLOROPEROXIDASE FROM NEPHRQMA LAEVIGATUM 145APPENDIX 7: LICHEN FEEDING EXPERIMENTS WITH STABLE ANDRADIOISOTOPES 146APPENDIX 8: HPLC DATA FOR LICHEN ANTHRAQUINONES, BIANTHRONES AND HYPERICIN DERIVATIVES 147viiLIST OF TABLESTable 1. Lichen taxa containing anthraquinones. 21Table 2. Major lichen compounds in Nephroma species. 32Table 3. Anthraquinone and anthrone pigments in Heterodermia speciesas determined by TLC. 35Table 4. ‘H NMR data for emodin (1), 7-chloroemodin (2), 7-chloro-1-O-methylemodin (3) and 1 -O-methylemodin (7). 42Table 5. “C NMR data for emodin (1), 7-chloroemodin (2) and 7-chloro-1-Q-methylemodin (3). 43Table 6. ‘H NMR data for 7-chloro-1-O-methyl-o-hydroxyemodin (4), 7,7’-dichlorohypericih (5) and 2,2’ ,7,7’-tetrachlorohypericin (6). 47Table 7. ‘3C NMR data for 7,7’-dichlorohypericin (5) and hypericin. 52Table 8. ‘H NMR data for emodin (1), 7-chloroemodin (2) and 5,7-dichioro-emodin (3). 59Table 9. 13C NMR data for emodin (1), 7-chloroemodin (2), 5,7-dichloro-emodin (3) and tiavoobscurin B (5). 60Table 10. 1H NMR data for flavoobscurin A (4), flavoobscurin B (5) and7,7’-dichlorohypericin (6). 64Table 11. ‘H NMR data for emodin (1), 7-chloroemodin (2), 7-chloro-1-O-methylemodin (3), 7-chloro- 1-O-methyl-o-hydroxyemodin (4),5-chloro-1-O-methylemodin (5), 5-chloroemodin (6), and 5-chloro- 1- O-methyko-hydroxyemodin (7) from isotope labellingexperiments. 69Table 12. Properties of radiolabelled anthraquinones isolated fromNephroma Iaevigatum. 70Table 13. ‘3C NMR data and isotope enrichments for 7-chloro-1-Q-methyl-emodin (3). 73Table 14. ‘3C NMR data for 7-chloroemodin (2), 7-chloro-1-O-methyl-emodin (3) and 5-chloro-1-O-methylemodin (5). 74Table 15. Reaction products from chlorination experiments with emodin(1) and 7-chloroemodin (2). 85Table 16. Minimum inhibitory concentrations of lichen compounds againstHSV-1 virus. 94Table 17. Inhibition of HSV-1 virus in plaque assay. 96Table 18. Effect of light on HSV-1 inhibition at different substrate concentrations. 97Table 19. HPLC data for lichen anthraquinones, bianthrones and hypericinderivatives. 147VIIIFigure 1.Figure 2.Figure 3.Figure 4.Figure 5.Figure 6.Figure 7.Figure 8.Figure 9.Figure 10.Figure 11.Figure 12.Figure 13.Figure 14.Figure 15.Figure 16.Figure 17.Figure 18.Figure 19.Figure 20.Figure 21.Figure 22.LIST OF FIGURES91225303337395457688083849193106140141142143144145146Lichen secondary metabolites.The biosynthesis of lichen compounds.The biosynthesis of anthraquinones in lichens and Dermocybe(basidiomycetes).The anthraquinones of Nephroma Iaevigatum.The anth raqu i nones of Heterodermia obscurata.Nephroma Iaevigatum from Galiano Island, British Columbia.The anthraquinones of Nephroma Iaeviagtum.Heterodermia obscurata from Cedarville State Forest, Maryland.The anthraquinones of Heterodermia obscurata.Anthraquinones from isotope labelling experiments with N.Iaevigatum.Autoradiograph of 2D TLC of radiolabelled compounds.Dimedon and Bradford assays.Biohalogenation experiments.Substrates and products from chlorination experiments.Structures of lichen compounds used in antiviral assays.The lichen polyketide pathway in Nephroma Iaevigatum.Approximate phytogeographical distribution of NephromaIaevigatum in North America.Approximate phytogeographical distribution of Heterodermiaobscurata in North America.Numbering systems in quinonoid natural products.Calculations of 13C carbon chemical shifts using the methodof Ewing (1979).Calculations of protein content and specific activity of semi-purified ch loroperoxidase from Nephroma Iaevigatum.UV spectrum of semi-purified chloroperoxidase fromNephroma Iaevigatum.Lichen feeding experiments with stable and radioisotopes.Figure 23.ixACKNOWLEDGEMENTSThe author extends sincere thanks to Professor Neil Towers for his kindness,invaluable guidance, advice and criticism in all matters pertaining to the researchperformed, and scientific results published, under his direction, as well as to Dr.James Hudson for his kindness and scientific support in doing all the antiviral workand assisting in its interpretation. The author also thanks Dr. Michael Adam, Professor Raymond Andersen, Professor Peter Hochachka, and Professor Alan Lewisfor their patience, kindness, advice and scientific support during the course of theauthor’s stay in the Department of Botany, and in the production of this thesis. Finally, I thank the external and university examiners for their critical evaluations ofthis thesis.Financial assistance rendered during the course of this work by the NaturalSciences and Engineering Research Council of Canada, and by the Department ofBotany, UBC is gratefully acknowledged.The author wishes to record his appreciation to the Chemistry Department ofUBC and the Laboratory of Chemistry, NIDDK, National Institutes of Health (Bethesda, Maryland, USA) for their assistance in recording NMR and mass spectra.Finally, the author extends his deepest appreciation to his parents for theirlove, constant and generous support, and invaluable guidance, as well as to XiaoqiuTang for her strong spirit, selflessness, constant encouragement and love.xFOREWORDThe objectives of the research performed over the last three years, as described in this thesis, were to: 1) contribute to the extant knowledge of the chemistry of lichens, in particular, pigment-containing lichens such as Nephroma Iaevigatum and Heterodermia obscurata; 2) establish the ubiquity of the polyketidepathway as the primary source of anthraquinones in all major classes of organisms, including lichens; 3) examine the role of chlorination in Nephroma Iaevigatum, by the isolation and purification of a chlorinating enzyme (chloroperoxidase), and the study of the comparative properties of the lichen enzyme and afungal chloroperoxidase, in a series of in vitro chlorination experiments with putative precursors of chlorinated anthraquinones in N. Iaevigatum; and 4) examinethe virucidal properties of several structurally diverse lichen anthraquinonoid natural products in two assays, using HSV-1 (herpes simplex virus type 1).1CHAPTER 1: OVERVIEW1.1 LICHEN CHEMISTRY AND CHEMOTAXONOMYLichens consist of symbiotic associations between algae and fungi, withapproximately 20,000 species distributed throughout the world, in all types of ecosystems (Ahmadjian and Hale, 1973). Traditionally, the study of lichens has relied on morphological and chemical characters as tools in their taxonomic classification; in fact, lichenology can claim one of the earliest uses of chemistry as ataxonomic tool in any of the botanical and mycological subfields ( Elix, 1992).More recent developments in cell biology and molecular biology, which have beenestablished in mycology for some time, are beginning to make themselves felt inlichenology, augmenting traditional taxonom ic and physiological approaches.The use of chemical methods in lichen taxonomy is not without controversy.While there are few detractors with respect to the inclusion of chemical spot tests(and thin-layer chromatography or TLC) in the set of criteria used to classify lichens,there are differing opinions regarding the relative importance of chemical criteriaand how they should be interpreted (Brodo, 1986). The use of chemical criteria inclassifying lichens began with W. Nylander in 1866, who investigated the reactionof lichen compounds with a variety of chemical reagents, producing characteristiccolour changes (Nylander, 1866a, 1866b, 1866c). In 1907, W. Zopf published“Die Flechtenstoffe”, describing his extensive chemical investigations of lichens(Zopf, 1907). Asahina (1937) proposed two principles for the application of chem2ical methods to lichen classification: 1) two morphologically indistinguishable lichens, which are found to contain different metabolites under the same environmental conditions, should be regarded as different species; 2) the concentrationratio of two or more secondary metabolites may vary depending on the lichenhabitat, and thus cannot be regarded as a reliable criterion in lichen taxonomicclassification. In the intervening years, microchemical analyses of lichens haveprogressed to the modern field of lichen chemistry.The debate dealing with the usefulness of chemistry in lichen classificationhas been thoroughly reviewed by S. Shibata (1964, 1973), M. E. Hale (1966, 1983),C.F. and W.L. Culberson (1969, 1970), D.L. Hawksworth (1976), I. Brodo (1986),R.S. Egan (1986), ft Rogers (1989), J. Poelt (1991), and J. Elix (1992). The mostcomprehensive treatment was that of Culberson and Culberson (1970), who systematically analyzed the distribution of 209 secondary metabolites in 2,315 species oflichens in order to discern chemical patterns that might be indicative of phylogeneticcharacteristics. The researchers reached several conclusions: 1) most lichen generaand families that are morphologically well-defined exhibit uniformity in their chemistry;2) genera, which have atypical chemical patterns for the families in which they aremembers, exhibit affinities with other genera or families based on morphological data;3) the highest degree of chemical variation occurs in the order Lecanorales, particularlywith the systematically useful depside and depsidone classes of compounds; and 4)comparative phytochemistry of lichens is the most reliable method available to aug3ment traditional morphological data in evaluating the “naturalness” of present systemsof lichen classification.Hale (1966), in one study, found a high degree of association between lichensecondary metabolites and particular morphological traits. He also suggests that theremay exist a correlation between the degree of O-methylation of lichen depsides anddepsidones and the structural complexity of the lichen genera containing them.According to this approach, he concluded that Cladonia represented the most advanced genus as it had the highest percentage of O-methylation; indeed, Cladonia isacknowledged by most lichenologists to be one of the most advanced lichen genera.Similarly, he cited the Sphaerophoraceae as an advanced family, and Umbilicaria asa “primitive” genus (the latter conclusion is also consistent with the current phylogenetic view). Large genera, such as Parmella, contain sections that are chemicallydiverse (and may be undergoing speciation) and chemically uniform groups that mayrepresent older stabilized lichen populations. Hale does recognize the limitationsinherent in using a single chemical trait in assessing the taxonomic rank of lichens,and points out that such data should be used in conjunction with other chemical andmorphological informationBrodo describes three fundamental problems in the application of chemistry tolichen taxonomy: practical problems, biological problems and philosophical problems.He emphasizes the importance of basic chemical techniques in systematics (spottests and thin-layer chromatography), as well as the increasing recognition of the im4portance of more rigorous methods, such as high performance liquid chromatography(HPLC), mass spectrometry (MS) and nuclear mag netic resonance spectroscopy(NMR). While the accessibility of even the “advanced” chemical instrumental methods (such as NMR spectroscopy and X-ray crystallography) has increased, the biological problems associated with the use of chemical criteria still remain. Brodo pointsout, for example, that some lichen compounds may be compartmentalized within thelichen, or may be produced during different stages of growth (Brodo, 1986). Culberson and Culberson (1958), in examining the genus Lasallia, found no variation inchemical composition during different stages of growth. The anthraquinones ofHeterodermia are known to be restricted to the lower ecorticate surface of the lichens,while Nephroma anthraquinones are strictly medullary pigments. Obviously, chemical examinations that do not take into account the whole gamut of lichen macro andmicrostructures, as well as the varying growth stages, are incomplete. Furthermore,as Poelt (1991) has pointed out, “it is very difficult, and may even be impossible, todiscriminate between homologies and analogies.” Poelt and Leuckert (1993), in discussing the pitfalls of taxonomic classifications based on chemical patterns, providenumerous examples of compound substitution and supplementation in a wide rangeof lichen genera.For example, some subtropical and tropical Caloplaca species contain usnicacid, rather than the usual assembly of anthraquinones: Caloplaca cuyabensis(Malme) Zahlbr. and C. stenospora (Malme) Zahlbr. are two such lichens. Caloplaca5variabilis (Pers.) Mull. Arg. appears not to contain simple anthraquinones at all,but rather greyish-violet amorphous pigments which may bear a biosynthetic relationship to anthraquinones. Krog (1970) reported the occurrence of a sample ofSolorina crocea (L.) Ach., in Finland, whose lower surface was completely devoidof the expected solorinic acid and related anthraquinones. How does one deal withAcroscyphus sphaerophoroides Lév., in which no less than five major structuralgroups of compounds were reported (Shibata et al., 1968)? A. sphaerophoroidesis described as having morphological similarities to another member of the Caliciales, Thelomma, which has a very different chemistry (Tibell, 1984). Yet, Acroscyphus contains chemical products of all the known lichen secondary metabolicpathways. Does this complexity of assembly lines represent a primitive stage oflichen evolution, from which more advanced genera evolved by selectively disCarding unnecessary metabolic machinery, or does such complex chemistry signify thehighest possible state of lichen morphological and physiological development?A clue leading to a partial resolution of this dilemma may be found in the increasing attention being given to the study of lichen genes (DePriest and Gargas,1994; Gargas et al., 1995). The available evidence (from other biological systems)suggests that there are close relationships between gene structure and biochemicalpathways, and alteration, nonexpression or elimination of genes will influence thenature and quantity of chemical products (Rogers, 1989). Culberson, Culberson andJohnson (1988) have examined the relationships between gene expression, lichen6development and chemical production in the Cladonia chlorophaea complex.They found that, in the Appalachian Mountains, two distinct chemotypes belonging to a single interbreeding population were reproductively isolated fromanother chemotype from the Coastal Plain of North Carolina. The chemistryof the offspring appeared to be reflective of the ability of any one particularchemotype to cross with another. The authors conclude that “The probablebiosynthetic relationships of the diagnostic compounds give no clue to thelimits of the interbreeding populations,” and further, “The old morphology!chemistry argument in lichen taxonomy becomes irrelevant in a problem nowanalyzable by experimentation.” Culberson thus proposed that chemical variants in lichens constitute sibling species, and single differences in chemicalcompositions warrant the attribution of distinct species status. Rogers (1989),in a review on chemical variation in lichens, counters that some chemical variation may be “genetically trivial” or environmentally induced. While acknowledging the role genes play in chemotypic variation, he does not believe thatsmall changes in the structures of lichen compounds (i.e., functional groupmodifications) are necessarily indicative of genetic isolation or population discontinuities. Rogers thus comes down on the side of “selection of strains bythe habitat, not of habitats by strains.” An additional point emphasized byRogers is the notion that biochemical pathways, rather than end products perSe, should be considered when making taxonomic assessments; it was sug7gested that the identification of a unique biosynthetic pathway (and its associated enzymes) should determine the suitability of assigning a unique taxonomic status to a particular lichen, along with proven genetic, morphologicalor physiological differences.Egan (1986), in a detailed study of chemical variation and lichen morphology and geography, described three categories based on the literature data: 1)distinct correlations between lichen chemistry and lichen morphology or geography;2) weak correlations between lichen chemistry and morphology or geography; 3)chemical strains in lichen species, where there does not appear to be any correlation between chemistry and morphology or geography. In his own study of Xanthoparmelia from Texas, Egan (1982) found chemosyndromic variation within differentbarbatic acid-producing species. Thus, each species of Xanthoparmelia produced acharacteristic set of biogenetically related depsides in the medulla. In each case,there was at least one major product and at least one minor product, but the primaryconstituent in any given species occurred as a minor component in all the others. Inanalyzing the various viewpoints regarding chemical applications to lichenology, itwould seem there is near unanimity among lichenologists regarding the inclusion, atsome level, of chemical variation in lichens in any evaluation of taxonomic status,even if there is still debate about the manner of interpretation of the data.As early as the 1 860s, when Nylander was analyzing lichens for chemical substances, the uniqueness of lichen compounds was recognized. To date, approxi8mately 5,000 lichen species have been analyzed for lichen substances, constitutingabout one third of all known species (Elix, 1992). Lichen compounds are structurallyunique and are rarely encountered in other organisms. The depsides and depsidones(Figure 1), the largest group of lichen compounds, are almost restricted to lichens, witha few exceptions known from lower fungi. Dibenzofurans (Figure 1) are also unique tolichens. Xanthones are known from lower fungi, lichens and higher plants; chlorinatedxanthones (Figure 1), however, are ubiquitous in lichens and unknown in higher plants.Similarly, anthraquinones are widely distributed among all classes of organisms, yetchlorinated anthraquinones (Figure 1) are practically restricted to lichens (a few examples are known from lower fungi, basidiomycetes and marine echinoderms). Mevalonate-derived terpenes and steroids (Figure 1), prevalent in both lower fungi andhigher plants, are fairly uncommon in lichens. Pulvic acid derivatives (Figure 1) arerestricted to lichens and fungi. Finally, lichens are the only known class of organismtotally devoid of alkaloids (although a few amino acid and peptide derivatives areknown) (Figure 1).One rather obvious, but hardly trivial, observation is that lichen chemistry mustsomehow be a consequence of the symbiotic partnership between the mycobiont(fungus) and phycobiont (alga). While it is generally thought that the mycobiont isthe source of a majority, if not all, of the lichen metabolites, experiments with culturedlichens and individual mycobionts have provided only a few clues as to the physiological bases for metabolite production (Renner and Gerstner, 1 978b, 1980). In a few9CH3 CH3OHCo— COOCH3H0’%,...)OHCHO CH3Atranorin (depside)CH3 C7H1CH3OJ%.._LO-%)OHMelacarpic acid (dibenzofuran)CH37-Chioroemodin (anthraquinone)N(CH)CH3O%.._%.%._LCQQ..HO>OCH3Solorinine (amino acid)CH3 0 OHCIHO%-L O_—’%)OHCIArthothelin (xanthone)Vulpinic Acid (pulvic acid)Pyxinic Acid (triterpene)COOHOHCICHOOHPannarin (depsidone)Figure 1. Lichen secondary metabolites.10reported instances, individually cultured mycobionts produce compounds not normallyseen in the lichen association (Miyagawa et al., 1994). The reason for this differenceis not yet clear, although it may involve either physiological or genetic adaptationsto the culture conditions. Miyagawa et al. (1994), in their work with cultured lichenmycobionts, have suggested that the production of anomalous metabolites might berelated to the osmotic stress induced during culturing of the mycobiont. In mostcases, though, the mycobiont-derived metabolites were of a structural class of compound known from the lichen association (Hamada and Ueno, 1990). In one particular study, Mathey et al. (1980) cultivated the mycobiont from the tropical lichen,Trypethelium eluteriae Sprengel. They isolated several 1 ,2-napthoquinone pigments;these compounds were lacking in the intact lichen and were unexpected, representing as they do a structural type previously known only from a few higher plant genera,such as Streptocarpus (Gesneriaceae).Culberson and Armaleo (1992) reported on the formation of a lichen-specificsecondary metabolic pathway in the cultured mycobiont of Cladonia grayl G.K. Merr.ex Sandst. The cultured mycobiont produced the same set of depsides and depsidones found in the natural lichen. The researchers also noted that the sequence ofbiogenesis of the metabolites was consistent with contemporary theories on depsidetransformations to depsidones, and metabolite productivity was comparable to thatseen in some nonlichen fungi. They ultimately concluded that the phycobiont (alga)is unnecessary for the production of depsides and depsidones in lichens.11Where parallels do exist between lichen and other plant or fungus-derivednatural products, they probably reflect the presence of ubiquitous biosynthetic pathways. Surprisingly, there is very little literature discussion of the comparative phytochemistries of lichens and other organisms; this lack may simply reflect the scarcityof published data on the biogenesis of lichen compounds (Mosbach, 1973). Of themajor types of lichen compounds, only the depsides, depsidones, pulvic acid derivatives, usnic acid, and tyrosine derivatives have been studied biogenetically (Figure2).The earliest detailed reports on the biogenesis of lichen compounds wereMosbach’s studies of gyrophoric acid (depside) and vulpinic acid (pulvic acid derivative) (Mosbach, 1964a, 1964b, 1967). Mosbach firmly established the origin ofgyrophoric acid, in Lasallia papulosa (Ach.) Llano, from malonyl-SCoA, using [1,3-14C] diethyl malonate. He also elucidated the origin of vulpinic acid, in Lethariavulpina (L.) Hue, from phenylalanine, using[1-’4C}-DL-3-phenylalanine. Subsequent research by Yamazaki and Shibata (1965, 1966) established the origin ofthe depsides lecanoric acid (from [1-14C] acetate) and atranorin (from [1-14C] acetate and [14C] formate), using the lichen Parmotrema tinctorum (Nyl.) Hale. Taguchi et al. (1966) and Pentillä and Fales (1966) concurrently demonstrated theformation of usnic acid from acetylmethylphloroglucinol. In a series of experiments,Bloomer et al. (1968, 1969 and 1970) studied the biosynthesis of (+)-protolichesterinic acid in Cetraria islandica (L.) Ach.; the origin of this acetogenin from14C-Iabelled012Biosynthesis of atranorin (depside) from labelled acetate and formate inParmotrema tinctorum.OHH3cOC.T%.TCH3HO.-OHOH+ I IIHOOHCOçH3Biosynthesis of usnic acid (dibenzofuran) from[14CH3-CO]-acetylmethyl-phloroglucinol in Usnea and Cladonia spp.Q_CH2_CH_ OOHBiosynthesis of pulvic acid from [1-4C]-DL-phenylalanine in Pseudocyphellaria crocata.*NH2HO* *CH3—s—(CH2)C-H HO—%jCH2 —CH—COOCH3COOH j’j(CH3) crBiosynthesis of sticticin (tyrosine derivative) from 3,4-DL-dihydroxyphenyl-[314C]alanine and L-[methyl-14C]methionine in Lobaria Iaetevirens.CH3’4OOH (*) +H14COOH (0).OH* 0COOCH3HI*Figure 2. The biosynthesis of lichen compounds.13acetate and succinate was firmly established. The bitter substance portentol, anunusual lactone from Roccella species, was found to be derived from both acetylSCoA and malonyl-SC0A (Aberhart et al., 1969, 1970). The biogenesis of the pulvicacid derivatives, calycin, vulpinic acid, pulvinic acid dilactone, pulvinamide, epanon pinastric acid, leprapinic acid and rhizocarpic acid were established using[1-14C] phenylalanine incorporated into Pseudocyphellaria crocata (L.) Vainio(Maass et al., 1964; Maass and Neish, 1967). In the intervening years, Blancoet al. (1984) demonstrated that [14C]urea was efficiently incorporated into evernicacid, atranorin and chloroatranorin in Evernia prunastri (L.) Ach., and Bernard andGoas (1981) studied the formation of amines from [2-14C]glycine in Stictaceae andlater, the biosynthesis of tyrosine derivatives in Lobaria Iaetevirens (Lightf.) Zahlbr.from L-[U-14Cjtyrosine and DL-[3-14CjDOPA (Bernard et al., 1981).Considering the number of identified lichen compounds (about 550), thestate of our knowledge regarding their biogenesis is still in its infancy. Whencompared to the data available for the biosyntheses of fungal, plant, microbial,insect and animal natural products, the limited information about the origins oflichen substances is clearly inadequate. What may be even more revealing isthe fact that since lichen biosynthetic studies began to ebb around 1970 (aquarter century ago), there have been only six additional papers in this areaof research. In 1969, Mosbach explained the justification for investigating thebiogenesis of lichen compounds as consisting of three factors: the uniqueness14of lichen compounds and their scarcity in other organisms; the uniqueness oflichens as symbiotic forms of life, and thus, the unique interactions the individualsymbionts must adopt in order to produce lichen substances; and the contributions that elucidation of biosynthetic pathways would make towards a more formative chemotaxonomy of lichens.Although the utility of lichen compounds for systematic purposes is generallyfirmly established, the roles they may play in the lichen itself are poorly understood.Lawrey (1986) recognized two categories of functions for lichen substances: 1) antimicrobial, antiherbivore and allelopathic protective roles; and 2) light screening,photobiont regulation and detoxification roles. While the evidence for the first category is fairly good, as a consequence of detailed ecological, physiological and pharmacological data, the experimental evidence for the second category is mixed andinconclusive.In one study, Lawrey (1989) found a correlation between the antimicrobialproperties of several lichen compounds and the ability of some lichens to deter popotential herbivores. He compared two lichens known to be consumed by the slug,Pallifera varia, with two species avoided by the slug. Acetone extracts of the preferred lichens exhibited lower antimicrobial activity than extracts of the avoidedlichens, for all the bacteria tested. In addition, the lichen compounds vulpinic acid,evernic acid and usnic acid were all found to have effective antimicrobial activityagainst the same bacteria. Lawrey thus concluded that lichen compounds can15protect the lichen from attack by either herbivores or microorganisms, but not inany specific manner. Gerson and Seaward (1977) demonstrated that snails (Helixhortensis) avoided certain lichens, but if the depsides had been removed by washingthe lichens with a dilute soda solution, the snails consumed the same lichens. Experiments with insects (Slansky, 1979; Emmerich et al., 1993) have shown that certainlichen compounds are repellent to insects, while others do not deter feeding, or mayalter normal insect growth and development after ingestion. Studies with vertebratesare few, and tend to be anecdotal. While it is well known that reindeer (Rangifertarandus) eat a variety of lichens, the specific lichen compounds that may act asattractants or deterrents to grazing are not known (Richardson and Young, 1977).While human beings are generally not considered major consumers of lichens,several lichen species are known for their use as food or in herbal preparations:Umbilicaria esculenta (Miyoshi) Minks (Richardson and Young, 1977), Bryoria fremonth (Tuck.) Brodo and Hawks. (Turner, 1977), Parmotrema tinctorum (Nyl.) Haleand Cetraria islandica (L.) Ach. (Richardson, 1975).Lichens and lichen compounds have been shown to be inhibitory towards thegrowth of plants, fungi, bacteria and other lichens (Huneck and Schreiber, 1972;Vartia, 1973; Rundel, 1978; Gardner and Mueller, 1981; Whiton and Lawrey, 1982,1984; lngOlfsdottir et al., 1985, 1994; Higuchi et al., 1993; Lawrey et al., 1994). Inparticular, lichens and their constituents have been shown to have strong enzymeinhibitory, antimicrobial and antiviral acitivities. Higuchi et al. (1993) tested forty-six16species of cultured lichen tissues for their inhibitory effect on the enzyme tyrosinase(an enzyme involved in the production of the skin pigment melanin). Three culturedlichens exhibited strong inhibitory activity; however, extracts of the natural lichenshowed only weak inhibition. Interestingly, none of the tested lichens known tohave anthraquinones showed major inhibitory activity. Work by lngOlfsdOttir et al.(1985) on the antimicrobial activity of seventeen lichen species from Iceland corroborated the results of Higuchi et al. (1993) that the primary compounds responsiblefor the antimicrobial activity were simple phenolics (e.g., methyl —-orsellinate), depsides (e.g., atranorin and chloroatranorin) and pulvic acid derivatives (e.g., vulpinicacid). Usnic acid from several Cladonia species has long been used as an antibioticin the treatment of dermatitis, eczema and other skin problems (Richardson, 1975).Lichens have also attained some importance, particularly in Japan, for the reputedantiviral activity of the constituent polysaccharides (Shibata et al., 1968; Nishikawaet aL, 1970; Takahashi et al., 1974; Nishikawa et al., 1979; Gorin and lacomini, 1984).In particular, polysaccharides from the genera Umbilicaria, Lasallia, Lobaria, Stictaand Usnea were found to be active. Although there has been only one report on thebiological activities of anthraquinones isolated from lichens (Cohen et al., 1995),known lichen anthraquinones that are also produced by fungi and higher plants havebeen examined for their enzyme inhibitory and antimicrobial activities. Anke et al.(1 980a, 1 980b) reported on the antimicrobial activities of emodin, parietin, parietinanthrone, catenarin, and several other anthraquinones from the fungus, Aspergillus17glaucus. The first four anthraquinones are known from several lichen genera,including Nephroma, Asahinea, Xanthoria and Caloplaca. Nikaido et al. (1984)tested several higher-plant and synthetic anthraquinones, several of which werealso known from lichens. The researchers found that emodin, 7-chloroemodin,citreorosein and chrysophanol exhibited strong inhibitory activity against cAMPphosphodiesterase. These anthraquinones are known from the lichen generaNephroma, Caloplaca, Lasallia, Heterodermia and Acroscyphus. Several otheranthraquinones, also known from lichens, exhibited little or no activity. Theseincluded skyrin (Pyxine, Phaeophyscia, Acroscyphus), parietin (Nephroma,Xanthoria) and emodin anthrone (Heterodermia).The role of lichen compounds as light-screening agents was originally suggested by ErtI (1951) and supported by Barkman (1958), who cited evidence forthe presence of higher concentrations of cortical substances in exposed lichensthan in lichens growing in the shade. Several other studies have attempted tocorroborate this conclusion (Scott, 1964; Hill and Woolhouse, 1966; Richardson,1967; Rundel, 1969). Most of these studies dealt with the anthraquinone parietin.For example, Richardson (1967) transplanted Xanthoria thalli from a variety ofdifferent habitats (but all containing parietin) to new locations. The surviving lichenthalli showed morphological changes and varying parietin content, probably as aresponse to the new habitat conditions. While there does seem to be a correlationbetween light intensity in a given lichen habitat and the concentrations of cortical18or medullary substances, additional studies are needed in order to corroborate theearlier findings (Lawrey, 1986).Finally, the potential role of lichen secondary metabolites as phycobiontregulators has been examined by Follmann et al. (1960, 1963, 1965, 1966), whodemonstrated that lichen compounds can alter the permeability of cell membranes,and thus cause the excretion of nutrients from the phycobiont (alga) to the mycobiont (fungus). Green (1970), however, attributed this effect to the acidity of theculture medium, and suggested that a buffered medium would considerably reduceany observed permeability changes resulting from lichen secondary metabolites.Regulation of enzyme activity was also indicated as another possible regulatoryrole for the lichen substances. Studies by Brown et al. (1982) showed that ureaseactivity in the blue-green-alga containing Peltigera canina (L.) Wilid. declined in response to the addition of urea. Vicente et al. (1976, 1978, 1979) found that severalendogenous lichen phenolic substances inhibited urease activity, and suggestedthat chelation of Mn2 (a cofactor for photosynthetic enzymes) in algal cells, by depsides like chloroatranorin, might represent a photosynthesis-regulating function forsecondary metabolites. Clearly, much more research needs to be done before thewhole range of biological, chemical and physical properties of lichen substancescan be fully appreciated.191.2 PHYTOCHEMISTRY OF ANTHRAQUINONESAnthraquinones are pigments widely distributed among all classes of organisms, including lichens (Thomson, 1971, 1987). So far, only the bryophytes havebeen found to be consistently lacking in anthraquinones. The anthraquinones represent the largest and best studied of the quinones. At present, about 40 differentanthraquinones have been characterized from lichens (Elix et al., 1984; Huneck,1984; Thomson, 1987; Huneck, 1991), with only eight new anthraquinones (andtwo new napthoquinones) having been identified in the last ten years (Huneck et al.,1991, 1994; Himmelreich et al., 1994; Cohen and Towers, 1995a, 1995c). In contrast, around 80 new anthraquinones from lower fungi and basidiomycetes havebeen discovered in the last decade (Gill, 1994, 1995). Despite the slow progressin the discovery of new lichen anthraquinones, many lichen genera still remain tobe systematically analyzed for new compounds.While anthraquinones are known from a taxonomically and morphologicallydiverse range of lichens, they are more prominent in some families and generathan others. Thus, anthraquinones are particularly prevalent in the lichen familiesTelosch istaceae (Caloplaca, Xanthoria, Teloschistes, Fulgensia), Psoraceae(Psora, Protoblastenia), Physciaceae (Heterodermia, Pyxine), Umbi I icariaceae(Lasallia), Parmel iaceae (Asahinea), Haematommataceae (Haematomma), Nephromataceae (Nephroma), Solorinaceae (Solorina). There are a few families that havenot been thoroughly examined, or for which there are occasional reports of anthra20quinones: Parmeliaceae (Cetraria, Parmella, Nephromopsis, Esslingeriana, Xanthoparmelia), Usneaceae (Usnea, Oropogon), Coccocarpiaceae (Coccocarpia), Physciaceae (Phaeophyscia), Caliciaceae (Acroscyphus), Laureraceae (Laurera), Sphaerophoraceae (Sphaerophorus) and Cladoniaceae (Cladonia) (Table 1).There are several important aspects to the distribution of anthraquinones, andnatural products in general, in lichens. First, there does not appear to be an obviousassociation between the presence of a particular anthraquinone (or other metabolite)and conspicuous morphological characters for a given lichen. Despite the generallyaccepted use of depsides and depsidones as chemical markers, their broad phytochemical distributions, and the lack of knowledge regarding why they are made,make it difficult to arrive at conclusions about supposed chemical affinities in lichens.In their study of chemical evolution in the cetrariold lichens, Kärnefelt and Thell(1993) found chemical representatives of all the major lichen metabolic pathways inthe cetrarioid genera. They ultimately concluded that the only discernible chemicalaffinity was the presence of higher aliphatic compounds among a majority of thecetrarioid genera. Pulvic acid derivatives (shikimate pathway) show some chemicalaftin ities in Pseudocyphellaria (Lobariaceae), Candalariaceae, Thelomma, Cypheliumand Acroscyphus (Caliciaceae). Triterpenes (mevalonate pathway) show chemicalaffinities in Lobaria and Pseudocyphellaria (Lobariaceae), Nephroma (Neph romataceae) and Peltigera (Peltigeraceae). In the case of anthraquinones, however, anypurported chemical affinities with known lichen taxa are even less obvious.21Table 1. Lichen taxa containing anthraguinones.4Family GenusCoccocarpiaceae CoccocarpiaSolorinaceae SolorinaNephromataceae NephromaLecideaceae LecideaLopadiaceae LopadiumMycoblastaceae Mycoblastus2Psoraceae ProtoblasteniaPsoraUmbilicariaceae LasalliaHaematommataceae HaematommaOphioparmaceae Ophioparma1Squamarinaceae SquamarinaPlacolecidaceae PlacolecisParmeliaceae AsahineaCetraria2EsslingerianaNephromopsisParmeliaXanthoparmeliaUsneaceae OropogonUsnea2Physciaceae HeterodermiaPhaeophysciaPyxine2Teloschistaceae CaloplacaFulgenisaTeloschistesXanthoriaCaliciaceae AcroscyphusSphaerophoraceae SphaerophorusArthoniaceae ArthoniaPyrenulaceae PyrenulaAcarosporaceae BiatorellaLaureraceae LaureraChiodectonaceae Chiodecton’Trypetheliaceae Trypethelium3Cladon iaceae Cladonia21OnIy napthoquinones reported.2Anthraquinones and napthoquinones reported.3Napthoquinone reported from cultured mycobiont;anthraquinones in native lichen.4Data from Culberson (1969), Mathey et al. (1980),Huneck et al. (1994) and Himmelreich et al. (1994).22Lichen species have rarely been assigned a taxonomic status on the basis oftheir anthraquinone constituents alone. There are, however, a few instances wherethe presence (or absence) of anthraquinones, along with distinguishing morphological traits, have been sufficient to warrant segregation of individual species intonew genera. For example, Rogers and Hafellner (1988) segregated the speciesHaematomma ventosum (L.) Massal. into the genus Ophioparma Norman (in a newfamily Ophioparmaceae Rogers and Hafellner) based on its non-lecanoroid ascusstructure, arctic-boreal distribution, saxicolous substratum preferences and the presence of the acetone-soluble red napthoquinone, haemoventosin, in the apothecia.In contrast, Haematomma puniceum (Ach.) Massal. and H. ochroleucum (Necker)Laundon were maintained taxonomically, in part, on the basis of having anthraquinone apothecial pigments (e.g., haematommone). The genus Lasallia consists ofabout 15 species, of which seven are known to contain anthraquinones (Posner etal., 1990, 1991). Llano (1950), in his definitive study of the Umbilicariaceae, segregated Lasallia from Umbilicaria based on the pustulate thallus, spore number in theascus and mode of reproduction. Even though the chemical structures of the Lasallia pigments were not fully characterized until 1969-1972 (Bohman, 1969b; Fox etal., 1969; Briggs et al., 1972), the pigmented thalli of several Lasallia species wererecognized by Llano (1950), Culberson and Culberson (1958) and others. Asahineawas segregated from Cetraria by Culberson and Culberson (1965) on the bases ofthe absence of rhizines or marginal ciliae, imperforate apothecial discs, geographical23distribution, absence of aliphatic compounds and the presence of purple pigments,later characterized as anthraquinones (Mischenko et al., 1980). The genera Pyxineand Heterodermia (Physciaceae) contain species having anthraquinones. Poelt(1965) originally proposed a new classification of Physciaceae, in which Heterodermia was segregated from Anaptychia primarily on the basis of morphological characters. Subsequently, Culberson (1966) maintained that Heterodermia should be givenseparate status on the basis of chemistry as well. We now know that approximately16 of the 81 known Heterodermia species contain anthraquinones, while they arecompletely absent from Anaptychia (Yosioka et al., 1 968c; Kurokawa, 1973; Trass,1992). In the case of Pyxine, opinions differ as to the relative merits of quinonoidpigment production as a taxonomic criterion. This appreciably reflects the lack ofchemical information about the exact chemical structures of the compounds, as wellas a lack of diagnostic uniformity in the assigning of taxonomic status. Rogers (1986),in his study of Pyxine in Australia, felt that cortical chemistry was an important taxonomic criterion, but medullary (pigment) chemistry was useful only as a confirmatorycharacter. Swinscow and Krog (1975), in their examination of East African Pyxine,placed a greater emphasis on the presence or absence of medullary pigments in thespecies they examined, but ruled out pigmentation of the internal stipe as a valid criterion. Imshaug (1957) in his systematic study of Pyxine in the New World, was themost enthusiastic of all about using the medullary and stipe pigments as valid markersin taxonomic classification. At present, the chemical structures of many pigments in24Pyxine species (Huneck, 1976) and Heterodermia species (Kurokawa, 1973) remainto be elucidated. This void would seem to support then, in principle, the advocacy ofusing caution in relying too heavily on any purported taxonomic implications of secondary chemistry in lichens, unless that chemistry was very well understood.The biogenesis of anthraquinones in lower fungi (Gatenbeck, 1958, 1960,1962), higher fungi (Steglich et al., 1972; Gill and Giménez, 1990a, 1990b, 1992) andhigher plants (Leistnér, 1971, 1973; Yagi et al., 1978) has been well documented.The majority of anthraquinones are derived from acetate (as acetyl C0A) or malonatethrough the polyketide pathway (Weiss and Edwards, 1980). A few anthraquinonesfrom higher plants are derived, instead, from shikimic acid (Leistner, 1973); but allknown fungal and lichen anthraquinones are believed to be derived solely from acetate or malonate (Culberson, 1969; Mosbach, 1969; Gill and Steglich, 1987; Gill,1994). The anthraquinone emodin (Figure 3) was shown to be formed from acetatein Penicillium islandicum (Gatenbeck, 1958, 1960, 1962), in the higher plant generaRhamnus, Rheum and Polygonum (Leistner, 1971), and in the basidiomycete generaCortinarius and Dermocybe (Gill and Steglich, 1987; Gill, 1994, 1995). Feeding experiments in Dermocybe have established the polyketide origin of emodin, and itstransformation into a variety of structurally modified anthraquinones (Steglich et al.,1972). In addition, several anthrone precursors, leading to a host of anthraquinoneend products in Cortinarius and Dermocybe, have been isolated, and their transformations into anthraquinones studied in vivo using13C-labelled acetate (Gill andGiménez, 1990a, 1990b, 1991, 1992).CH3OSCoA + 7 HOOC-CH2-COSCoA 250CO-S-CoAOH 0 OH OH o OH8a J& 9a JJCIri9Y IXICHHO%CH3 HOoEmodin[Cl] I [OH][CH]i/CH3Emodinanth rone\O]Endocrocin•COOHCH3(Dermocybe only)-ø[CO/7-Ch loroemodinanth rone[Cl]CIHO5,7-Dichloroemodinanthroneci R 0 R5-Ch loroendocroci n[CH3] 4, [OH]OCH 0 OHCOOHCH3CI 0 RR = R’ = = = H; 7-Chloroemodin R = H; 5-ChiorodermoluteinR = R” = R’” = H; R = OH; Papulosin R = OH; 5-ChlorodermorubinR = Cl; R = R” = R” = H; 5,7-DichloroemodinR = R’ = R’” = H; R” = CH3; 7-Chloro-1-Q-methylemodinR = R’ = R” = H; R” = CH3; FragilinR = R’ = H; R” = R’” = CH3; 7-Chloro-1 ,6-di-O-methylemodinFigure 3. The biosynthesis of anthraquinones in lichens and Dermocybe (basidiomycete).26The few biosynthetic studies on lichen compounds have dealt primarily withdepsides and pulvic acid derivatives (Maass et al., 1964, 1967; Yamazaki et al.,1965, 1966; Maass, 1970; Bloomer et aL, 1970). So far, there have been no studies on the biogenesis of anthraquinones in lichens; the assumption has beenmade that they originate from acetate and malonate through the polyketide pathway (Shibata, 1964; Culberson, 1969; Mosbach, 1969; Santesson, 1970a).Santesson (1 970a) proposed a scheme describing the possible biogeneticrelationships between the anthraquinones found in the genus Caloplaca. He suggested that anthrones might be precursors to corresponding anthraquinones, basedon their co-occurrence in some lichens. In addition, Santesson (1970a) was the firstto propose that chlorinated anthraquinones could be formed, in lichens, by directhalogenation of emodin and related compounds (Figure 3). Steglich et at. (1969)had suggested a similar mechanism for the formation of chlorinated anthraquinonesin the basidiomycete genus Dermocybe (Figure 3). Although such proposals haveyet to be tested experimentally for lichens or basidiomycetes, a mechanism for thechlorination of anthraquinones in the lower fungus, Aspergillus fumigatus, has beenexamined (Yamamoto et al., 1968). The authors grew cultures of the fungus withvarying concentrations of chloride ion in the medium. They were able to establishan optimum concentration of chloride ion which resulted in the highest yield of chlorinated anthrones and anthraquinones in the fungus. When bromide ion was substituted for chloride ion in the culture medium, the corresponding brominated anthrone was produced by the fungus.271.3 THE CHEMISTRY OF NEPHROMA AND HETERODERMIAThe genus Nephroma was reviewed extensively by Gyelnick (1931, 1932),later followed by Wetmore (1960) who revised the North American taxa. In theintervening years, Renner et al. (1982) categorized the South American speciesof Nephroma into nine chemical groups. Finally, James and White (1987, 1988)systematically studied the European, Macronesian, northern temperate and southern temperate species. The researchers analyzed the chemistry, by TLC, of atotal of 28 species, in an attempt to formulate phytochemical groupings within thegenus.The earliest studies of the chemistry of Nephroma were those of Bach-mann (1887). Despite some confusion surrounding the identity of the lichen substances, Bachmann was able to recognize the presence of a yellow emodin-likecompound in the medulla of Nephroma Iaevigatum Ach. (as N. lusitanicum (Ach.)Nyl.) which turned purple with KOH. The presence of emodin in higher plants(Frangula alnus) had already been established. Hesse (1 898a) also confirmed thepresence of an hydroxyanthraquinone (nephromin) in N. Iaevigatum, but regardedthe substance as unique to lichens. Zopf (1907) later compared nephromin to theyellow substance studied by Bachmann, and found they were identical. Bendz etal. (1967) separated nephromin into five compounds, and characterized each bymass spectrometry. Bohman (1968), continuing this work, fully established theidentity of the five anthraquinones, utilizing mass spectrometry, IR, UV and 1H NMR28spectroscopy. In addition, Bohman isolated a triterpene (possibly nephrin) and afatty acid. The Nephroma compounds are shown in Figure 4. Several other lichensubstances have been isolated from Nephroma species, including nephroarctin(depsidone from N. arcticum (L.) Torss.; Nuno et al., 1969), phenarctin (depsidefrom N. arcticum; Brüun, 1971), gyrophoric acid and tenuiorin (depsides from N.pseudoparile Räs.; Renner et al., 1982), perlatolic and glornelliferic acids (depsides from N. cellulosum (Sm.) Ach.; Renner and Gerstner, 1978a), ergochromes(dimeric xanthones from N. analogicum Nyl.; Renner et al., 1982), and a series ofhopane triterpenoids present in almost all Nephroma species (James and White,1987; White and James, 1988).In their phytochemical analyses of 15 South American species of Nephroma,Renner et al. (1982) recognized nine chemical groupings. One species, N. analogicum, was found to contain yellow ergochrome pigments (secalonic acids A andC), which had been known previously only from the genera Parmelia (Yosioka etal., 1 968d) and Nephromopsis (Yosioka et al., 1972; Kärnefelt and Thell, 1993).There was no evidence, however, for the presence of anthraquinones in any ofthe southern species.James and White (1987) reexamined the European and MacronesianNephroma, recognizing seven distinct chemical groupings (Table 2). Anthraquinones were identified in three species, N. Iaevigatum, N. tangeriense (Maheu andA. Gillet) Zahlbr. and N. venosum Degel. The authors report that up to 16 anthra29quinones may be present in some varieties of N. Iaevigatum, according to TLC,but also indicate that nonpigmented or pigment-deficient morphotypes have beenfound in Portugal and the Canary Islands. N. tangeriense was said to bear a similarity to N. Iaevigatum chemically, although it is restricted to southern Europe andNorth Africa and generally lacks apothecia. N. venosum is endemic to the Azoresand morphologically distinct from N. Iaevigatum or N. tangeriense. Finally, in theirstudy of the southern temperate Nephroma, White and James (1988) expanded onthe earlier work of Renner et al. (1982). Their chemical results are consistent withthe phytochemical data obtained by Renner et al. (1982), and again confirm the absence of anthraquinones in the southern temperate species (Table 2).The chemical study of Heterodermia was initiated in 1898 with Hesse’s identification of the hydroxyanthraquinone pigment blastenin in Heterodermia obscurata(Nyl.) Trevis. (as Anaptychia heterochroa Vain.) (Hesse, 1 898b, 1901). Asahina andYosioka (1940) isolated the triterpene zeorin and depside atranorin from Heterodermia speciosa (Wulf.) Trevis. (as Anaptychia speciosa (WuIf.) Mass.), H. hypoleuca(Ach.) Trevis. (as A. hypoleuca (Ach.) Mass.), and H. obscurata (as A. heterochroa).They also recognized the presence of a yellow-orange undersurface in H. obscurata,and attributed this colour to an hydroxyanthraquinone (identical with blastenin) whichthey then isolated. Hale (1956) analyzed several North American lichens, includingHeterodermia obscurata (as Anaptychia heterochroa) and observed the presence ofa yellow anthraquinone in the lower surface of this lichen. The first systematic study30OH 0 OHCH37-Chioroemodin7-Chloro-1 -0-methylemodin‘CH37-Chloro-6-O-methylemodin 7-Chloro-1 ,6-di-O-methylemodinEmodin000Figure 4. The anthraquinones of Nephroma Iaevigatum.31of Heterodermia and Anaptychia was that of Kurokawa (1962). In his monograph,the author revised all species classified in the genus Anaptychia. Using chemicaland morphological data, Kurokawa proposed a new infrageneric classification ofAnaptychia, but did not feel the chemical data warranted the need for segregationof taxa into new genera. Poelt (1965), on the other hand, proposed a new classification of the Physciaceae, in which Anaptychia would be segregated into Anaptychia and Heterodermia.According to Poelt’s classification, Anaptychia is primarily characterized bythe presence of thin-walled spores and the absence of atranorin, while Heterodermia has thick-walled spores and contains atranorin. It was later shown by Culberson (1966) and Kurokawa (1973) that, in fact, atranorin is present in some Anaptychia species. This was followed by a major study by Culberson (1966). Theauthor examined 14 species of Anaptychia from North and South Carolina. Following the principles set forth by Poelt (1965), Culberson separated Heterodermiafrom Anaptychia on the basis of spore morphology as well as chemistry. The authordemonstrated that all species of Heterodermia contained, in addition to atranorin,other lichen substances (e.g., zeorin, salazinic acid, norstictic acid and anthraquinones) completely absent from Anaptychia.Yosioka et al. (1968a, 1968b) examined, in some detail, the chemistry ofHeterodermia obscurata. Several anthraquinones and bianthrones were isolatedfrom the lichen growing in Japan (Figure 5). In a survey of several other pigmentedTable2.MajorlichencomDoundsinNeohromasoecies.N.analogicumNyl.N.antarcticum(Jacq.)Nyl.N.arcticum(L.)Torss.N.areolatumJames&WhiteN.australeA.RichRace1N.australeA.RichRace2N.bellum(Sprengel)Tuck.Race1N.bellum(Sprengel)Tuck.Race2N.cellulosum(Ach.)Ach.N.chubutenseLambN.expallidum(Nyl.)Nyl.N.foliolatumJames&WhiteN.helveticumAch.N.hensseniaeJames&WhiteN.isidiosum(Nyl.)GyelnikN.kuehnemannhiLambN.IaevigatumAch.Race1N.laevigatumAch.Race2N.microphyllumHenssenN.occultumWetmoreN.papillosumWhite&JamesN.parile(Ach.)Ach.Race1&2N.parile(Ach.)Ach.Race3N.plumbeum(Mont.)Mont.N.pseudoparile(Räsänen)Zahlbr.N.resupinatum(L.)Ach.N.rufum(Church.Bab.)JamesN.silvae-veterisGoward&GoffinetN.skottsbergllWhite&JamesN.sulcatumJames&WhiteN.tangerienseMaheu&A.GilletN.venosumDeqel.T3T3T3-T6T3T3T1-T6T2/T3/T5T3T6T2/T3/T5T3[Ti]/T4T1IT3IT41T6[Ti]/T4T6T3T3T3T2/T3/T5Ti/T3/T4/T6T3[T2]/T5T3T3/T4/T5/T6[T2]/T3/[T41/T6T3C”C)NephromaspeciesPhenarctin(P)GyrophoricUsnicacidSticticacidHyposalazinicAnthraquinonesHopaneNephoarctin(N)acid(G)(U)(S)acid(HZ)triterpenoids[HZ]HZS/C/HC/HS5HSPR/SP/GL4PNPP[P1/NT 2UUUUU[U][U]UUUUUECAA&AB6AQAQAQ[G]G/MG/MGA31Symbolkey:compoundsdetectedbyTLCareindicatedby“U”(present)or“[U]”(presentinsomesamples).2Containstenuiorin(T).Containsmethylgyrophorate(MG)and4-O-methylgyrophoricacid(MGA).Containsperlatolicacid(PR),stenosporicacid(SP)andglomellifericacid(GL).Containsconsticticacid(C),hypoconsticticacid(HC)andhyposticticacid(HS).ContainsergochromesAAandAB(ECAA&AB).UnknownxanthonedetectedinRace2.8DatafromRenner.etal.(1982),JamesandWhite(1987)andWhiteandJames(1988).33CH3OH 0 OHFlavoobscurin BCH3CH30 OHCH3Emodin 7-ChioroemodinCI 05,7-DichioroemodinOH 0 OHFlavoobscurin AFigure 5. The anthraquinones of Heterodermia obscurata.34Heterodermia (Yosioka et a!., 1968c), anthraquinones and anthrones were detected,by TLC, in eight tropical and semi tropical Heterodermia (Table 3). In addition, several yellow non quinonoid pigments (possibly pulvic acid derivatives) were detectedin two species.The last comprehensive study of Heterodermia (as Anaptychia) was that ofKurokawa (1973), who reviewed all known species, performed TLC analyses, discussed phytochemical variations within the genus, provided geographical distributions, and reclassified Anaptychia into two subgenera: subgenus Anaptychia andsubgenus Heterodérmia. Anaptychia was defined as having subascending or erectlobes, and an upper cortex with varying thickness; Heterodermia, in contrast, hasadnate lobes and a rather uniform upper cortex. As in his earlier work (Kurokawa,1962), chemical variation was not considered as a factor in taxonomic reassessment.Finally, Trass (1992) compiled the known information for the 81 acceptedspecies of Heterodermia in tabular form, providing morphological and geographicaldata, and indicating the presence or absence of pigments in each species.It)CoHeterodermiaRange7-Chioroemodin7-Chloroemodin5,7-Dichioro-Flavoob-FlavoobSpeciesanthroneemodinscurinAscurinBH.cyathiformis(Kurok.)S.Africa++++++H.dendritica(Pers.)PoeltE.,S.E.Asia+++++++++H.fauriel(Kurok.)2Hawaii,ThailandH.firmula(Nyl.)Trevis.E.,S.E.Asia+++++++++H.flabelatta(Fee)AwasthiS.Am.,Asia,E.Afr.++++++++++H.hypocaesia(Yasuda)AwasthiAus.,Asia,S.AfricaH.hypochraea(Vain.)Swinscow&KrogS.Am.,E.Afr.,Asia+÷+++H.loriformis(Kurok.)Swinscow&KrogE.Africa++++H.lutescens(Kurok.)Folim.S.Am.,Africa,Asia+1-H.obesa(Pers.)Trass2HawaiiH.obscurata(Nyl.)Trevis.Aus.,N.Z.,USA,Eur.++++++++++S.Am.,Asia,E.Afr.H.pacifica(Kurok.)Japan,Taiwan++H.pandurata(Kurok.)TrassJapan,Taiwan,Thai.++++H.propaguilfera(Vain.)DayS.Am.,Asia,USA(?)++++++++++H.rugulosa(Kurok.)TrassMexico,S.W.USA++++++++++H.subascendens(Asah.)TrassJapan,Taiwan,China++++H.vulgaris(Vain.)FolIm.&Rédon2S.Am.,E.&S.Africa1 Symbolkey:compoundsdemonstratedbyTLCaremarkedwith++(significant)or+(trace).CompoundsdemonstratedbyTLCinaportionofthesamplesaremarked++(+)or-(absent).2 Uncharacterizedanthraquinonesandanthrones.3 DatafromYosiokaetal.(1968)andKurokawa(1973).Table3.AnthraquinoneandanthronepicimentsinHeterodermiaseciesasdeterminedbyTLC.1 ’ 336CHAPTER 2: CHEMICAL PROPERTIES OF LICHEN QUINONOID PIGMENTS2.1 ISOLATION OF NEPHROMA LAEVIGATUM NATURAL PRODUCTS2.1.1 INTRODUCTIONThe lichen Nephroma Iaevigatum is widely distributed throughout westernEurope and the Mediterranean, where its range extends east to Israel (James &White, 1987). In North America, it can be found on both the Atlantic and Pacificcoasts. In the east, the lichen is known from Massachusetts to Labrador, whilein the west it occurs from northern California to British Columbia (Wetmore, 1960).This particular species is the most “oceanic” of the North American Nephroma,restricted to coastal areas near the Atlantic and Pacific Oceans (Figure 6; Appendix 1).In British Columbia, Nephroma Iaevigatum is abundant on sub-basicshoreline rocks on islands dotting the mainland coast north to Alaska. It can alsooccasionally be found growing on the trunks and branches of Big-Leaf Maple(Acer macrophyllum) nearthe coast.2.1.2 MATERIALS AND METHODSSeveral collections of the lichen were made from different islands in southwestern British Columbia: Bowen Island, Gabriola Island and Galiano Island.Attempts were made to locate the lichen on the adjacent mainland, but withoutsuccess. Nephroma Iaevigatum, in British Columbia, would appear to be highlylocalized in humid relic coastal forests on the numerous islands adjacent to the37Figure 6. Nephroma Iaevigatum from Gabriola Island, British Columbia.38mainland.The different collections were cleaned of soil, moss and other debris andair dried. Two reference samples were deposited in the UBC Botany DepartmentHerbarium. Small quantities of lichen were extracted with ice-cold diethyl etherfor 10 minutes, and the extracts examined by TLC. The patterns of chemical products were identical for all the samples examined; this suggests that there is little(or no) chemotypic variation within the lichen communities sampled. This uniformity does not exclude, however, the possibility that N. Iaevigatum chemotypesexist in other localities within British Columbia. In fact, James & White (1987) reported on the existence of two distinct chemical races of N. Iaevigatum, differingonly by the substitution of one hopane triterpenoid with another. The rarer Race2 (Table 2) was found in only two collections from the Azores Islands. In addition,rare pigment deficient varieties have been found on the Canary Islands, Madeiraand in Portugal (James & White, 1987). These chemotypes are morphologicallyand chemically identical to the common N. Iaevigatum, the only difference beingthe complete or partial absence of anthraquinones in the medulla.The dried lichen material was combined (0.4 Kg) and extracted, in succession, with cold diethyl ether, acetone and methanol. The extracts were concentrated to small volumes and examined by TLC. The diethyl ether extract showed 12coloured spots, the primary constituent being 7-chloroemodin (2) (R 0.6; 9:1chloroform:methanol). The other prominent compounds present in the extract39H0Emodin (1)RHOCIHO7-Chloroemodin (2)R = CI; 7-Chloro-1-Q-methylemodin (3)R = H; 1-O-methylemodin (7)7-Chlorocarviolin (4) R = H; 7,T-Dichlorohypericin (5)R = CI; 2,2’,7,7’-Tetrachlorohypericin (6)0 0CH30ciFigure 7. The anthraquinones of Nephroma Iaevigatum.40were: emodin (1) (R 0.8); 7-chloro-1-O-methylemodin (3) (R 0.7); 7-chloro-1-Q-methyko-hydroxyemodin (4) (R, 0.3); 7,7-dichlorohypericin (5) (R1 0.2); and2,2’,7,7’-tetrachlorohypericin (6) (R 0.2). These compounds are shown in Figure 7.Minor coloured constituents, evident from the TLC plate, were presumed to beother anthraquinones or anthrones based on their UV fluorescence, but were notpresent in sufficient quantities to permit conclusive identification. Two additionalnon coloured spots that fluoresced under UV light were probably phenolic constituents, such as depsides.Following TLC examination, the orange-red extract was concentrated to abrown-red solid. The solid (10 g)was divided into three equal portions. One portion (3.3 g) was purified by column chromatography on 100 g of Sephadex LH-20,using a gradient of chloroform:methanol (9:1) to methanol. Fractions (20 ml) werecollected and analyzed by TLC. Fractions 2-4 contained emodin (1) and 7-chloro-1-O-methylemodin (3). Fractions 5-7 contained 7-chloro-1 -O-methylemodin (3).Fractions 6-8 contained 7-chloro-1-O-methylemodin (3) and 7-chloroemodin (2).Fractions 9-12 contained 7-chloroemodin (2). 7-Ch loro- 1- Q-methyl-o-hydroxyemodin (4) came from fractions 13-18. Small amounts of 7,7’-dichlorohypericin(5) and 2,2’,7,7’-tetrachlorohypericin (6) were found in fractions 19-22. In order toobtain the bulk of compounds 5 and 6, the stationary purple layer was extrudedfrom the column and then extracted with pyridine for 16 hours. The purple extract41was concentrated to give a mixture of compounds 5 and 6. Separation of the mixture was accomplished by column chromatography on Sephadex LH-20, usingmethanol as the eluant. Prior to spectroscopic analysis, all recrystallized compounds were checked for purity by TLC and reversed-phase HPLC.2.1.3 RESULTS AND DISCUSSIONSpectral analysis of compound 2 supported the structure as 7-chioroemodin.The ElMS (Electron Ionization Mass Spectrometry) shows peaks at 306 and 304and the CIMS (Chemical Ionization Mass Spectrometry) reveals peaks at 307 and305. The 2D COSY (Two-Dimensional Correlated Spectroscopy) 1H NMR spectraldata are shown in Table 4. The assignment of the chlorine to position 7 is based onthe chemical shifts of H-5, H-4 and H-2, which are consistent with those previouslyreported (Bohman, 1968; Yamamoto et al., 1968; Yosioka et al., 1968a). SeveralNOE (Nuclear Overhauser Effect) experiments demonstrated an enhancement ofthe signals of H-2 and H-4 upon irradiation of the methyl protons, as well as an enhancement of H-5 upon irradiation of the hydroxyl proton at C-6. These results areconsistent with structure 2. 13C NMR assignments listed in Table 5 are based onAPT (Attached Proton Test) and HETCOR (Heteronuclear Chemical Shift Correlation) experiments, and calculations of carbon chemical shifts using the methodsdescribed by Ewing (1979) and Silverstein et al. (1991).Compound 3 proved to be the 1 -0-methyl derivative of 2 by analysis of the42Table 4. 1H NMR data for emodin (1), 7-chloroemodin (2), 7-chloro-1-O-methyl-emodin (3) and 1-O-methvlemodin (7).’Proton 12 2 33 722 7.15,s 7.14,s 7.45,s 7.45,s4 7.58, s 7.50, s 7.62, s 7.67, s5 7.25, d (2.5) 7.26, s 7.22, s 7.20, d (2.5)7 6.65, d (2.5) 6.67, d (2.5)1-OH 12.05,s 11.92, s6-OH8-OH 12.15,s 12.78,s1-OMe 3.98, s 4.05, s3-Me 2.47, s 2.48, S 2.52, s 2.50, s1Chemical shifts (ö) are reported in ppm from TMS internal standard. The coupling constants are given in Hz.2The spectra were recorded inMe2CO-d6at 300 MHz.3The spectra were recorded in DMSO-d5at 500 MHz.431Chemical shifts (S) are reported in ppm from TMS internal standard.The spectra were recorded in DMSO-d6at 125 MHz.2Values calculated according to the methods described in Ewing (1979)and Silverstein et al. (1991).3Values for C-8a and C-ga can be interchanged.4Values for C-lOa and C-4a can be interchanged.Table 5. 13C NMR data for emodin (1), 7-chloroemodin (2) and 7-chioro-1 -Q-methylemodin (3)1Carbon 1 12 2 22 3 32123456789104a8a39a31 0a41-OMe3-Me158.7120.1142.4125.2111.8162.8106.7160.2189.3173.9131.5110.0107.1132.8161.3124.0148.1120.3108.7165.5107.8164.4189.5181.1132.6108.8113.1134.921.5158.7120i142.4125.2113.2163.2112.9160.6187.4173.9‘131.5111.3107.1130.9161.4124.0148.5120.4108.6163.1121.3162.8191.0181.7133.2110.1114.2133.021.6163.2118.5142.0124.9113.2163.2112.9160.6187.1173.5131.1111.0107.1129.3160.9120.8148.3120.5106.9162.1118.5161.1187.5182.6134.5111.4114.2132.356.421.7442D COSY 1H NMR (Table 4) and ElMS spectra. The ElMS of 3 shows peaks at 320and 318, while the CIMS shows peaks at 321 and 319. The UV and 2D COSY 1HNMR spectral assignments are consistent with previously reported data (Bohman,1968; Ayer and Trifonov, 1994). A series of NOE experiments confirmed the identityof 3. Upon irradiation of the methyl protons, there was a corresponding increase inthe signal intensities of H-2 and H-4. Irradiation of the methoxy protons resulted inan enhancement of the H-2 signal. Finally, irradiation of the hydroxyl proton at C-6produced an enhanced signal for H-5. The 13C NMR data are shown in Table 5.Assignments were made on the basis of APT and HETCOR experiments, as well ascarbon chemical shifts calculated according to the methods described by Ewing (1979)and Silverstein et al. (1991). Additional structural proof was afforded by reduction of3 with Raney nickel to give the dechlorinated product (Figure 7). The 2D COSY 1HNMR shifts (Table 4) of this product are consistent with those previously reported for1-O-methylemodin (7) (Ayer and Trifonov, 1994). A comparison of TLC Rf values forcompound 7 and published data for 1-Q-methylemodin also helped in confirming theidentity of compound 3 (Ayer and Trifonov, 1994; Cameron and Crossley, 1977).Emodin (1) was characterized by UV, ElMS, CIMS and 2D COSY 1H NMRspectra (Table 4). Several NOE experiments confirmed that compound 1 was emodin.Irradiation of the methyl protons produced a corresponding increase in the signal intensities of H-2 and H-4. Irradiation of the hydroxyl proton at C-8 also produced an increase in the signal intensity of H-7. Table 5 gives the ‘3C NMR data as well. The45shift assignments were made on the basis of APT and HETCOR experiments, aswell as carbon chemical shifts calculated according to the methods described byEwing (1979) and Silverstein et al. (1991). The data are in agreement with thosepreviously reported (Bohman, 1968; Gill et al., 1988).Compound 4 was quite different from the other anthraquinones. It wasapparent from the 2D COSY 1H NMR spectral data that the signal correspondingto the methyl group of emodin had been replaced by a new one at 6 4.73 ppm.This suggested the presence of an hydroxymethyl group at C-3 (Table 6). Furtherconfirmation was obtained from the CIMS and UV spectra, which demonstratedthe compound to be 7-chloro-1 -0-methyko-hydroxyemodin (7-chloro-1 -0-methylcitreorosein or 7-chlorocarviolin) (4). An NOE experiment was performed in whichirradiation of the hydroxymethyl protons resulted in increases in the signal intensities of H-2 and H-4. 7-Chlorocitreorosein had been isolated from Aspergillus fumigatus (Yamamoto et al., 1968), and 1-0-methylcitreorosein (carviolin) has beenobtained from a culture of Penicililum roseo-purpureum (Hind, 1940). Compound4 thus represents a new natural anthraquinone.Compounds 5 and 6 are related to the well-known natural product hypericin,found in Hypericum species (Brockmann and Sanne, 1957; Falk and Schmitzberger, 1992) and in a basidiomycete, Dermocybe austroveneta (Gill et al., 1988).Hypericin is the subject of intensive medical scrutiny because of its antiviral activity46(Lopez-Bazzocchi et aL, 1991). Meruelo and coworkers, in 1988, showed thathypericin inhibited the spread of the Friend and radiation leukemia viruses in vitroand in vivo (Meruelo et al., 1988). The same group also reported that hypericincan inactivate human immunodeficiency virus (HIV), when measured by reversetranscriptase (RT) activity; it would appear, however, that the purified enzyme isnot the main target of hypericin activity (Lavie et al., 1989). Thus, the mode ofaction of hypericin still remains a topic of debate (Kraus et al., 1990).The only known natural halogenated hypericin-like compounds are thegymnochromes, brominated phenanthroperylenequinones in the crinoid, Gymnocrinus richeri (De Riccardis et al., 1991). The identities of compounds 5 and 6became apparent on an examination of the UV spectra, which were similar to thatof hypericin; they were also in very close agreement with the UV spectra of thegymnochromes (De Riccardis et al., 1991), and synthetic brominated derivativesof hypericin (Falk and Schmitzberger, 1992).The negative-ion LSIMS (Liquid Secondary Ion Mass Spectrometry) spectrum of 5 shows three peaks at 575, 573 and 571 (relative intensity 1:4.2:6). Thisis indicative of two chlorine atoms in the molecule. The combination of 2D COSYH NMR (Table 6) and LSIMS data indicate that 5 is a symmetrical dimer. A seriesof NOE experiments confirmed that 5 has the structure shown. Irradiation of themethyl protons produced an enhancement of protons H-2 and H-2’. Irradiation ofthe HO-8 (HO-8’) hydroxyl proton did not, however, produce an NOE enhancement47Table 6. 1H NMR data for 7-chloro-1-O-methyl-co-hydroxyemodin (4), 7,7’-dichlorohy-pericin (5) and 2,2’,7,7’-tetrachlorohypericin (6) .Proton - 42 - 53 54 62,2’ 6.78, s 7.42, s 7.20, s4,4’ 7.78, s5,5’ 7.43, s7,7’1,1’-OH 13.79,s 13.95,s6,6’-OH 18.28, s8,8’-OH 15.55, s 15.65, s3-OMe 3.95, s3-CH2OH 4.63, d (5.0)3-Me, Me’ 2.65, s 2.70, s 2.80, s1Chemical shifts () are reported in ppm from TMS internal standard. The couplingconstants in parentheses are given in Hz.2The spectrum was recorded in DMF-d6 at 300 MHz.3The spectra were recorded in DMSO-d at 400 MHz.4The spectrum was recorded in MeOH-d4at 400 MHz.5The hydroxymethyl protons were coupled to a broad hydroxyl proton. When the hydroxyl proton was exchanged by addition of a small amount of MeOH-d4, this signalbecame a singlet.48at positions C-7 and C-7’.The UV spectrum of 6 is very similar to that of 5. The LSIMS (negative ion)spectrum shows four peaks at 645, 643, 641 and 639 (relative intensity 1:3:5:3.5).This is indicative of four chlorine atoms in the molecule. The 2D COSY H NMRspectral data are shown in Table 6. Compound 6, like 5, must also be a symmetrical dimer based on the spectroscopic evidence.The number of free hydroxyl groups in each assigned structure was confirmed by peracetylation with acetic anhydride in pyridine. Mass spectra weredetermined by direct injection of the acetylation mixtures. Observation of the progressive loss of ketene was especially helpful in confirming the dimeric natureof 5 and 6.The biogenesis of the anthraquinones and the perylenequinones presumably takes place through the polyketide pathway (Figure 3). Detailed studies onthe biosynthesis of emodin and related compounds in fungi and higher plantshave been carried out (Gatenbeck, 1962; Zenk and Leistner, 1968; Leistner,1971; Steglich et al, 1972; Leistner, 1973; Shibata, 1974; Casey et al., 1978;Weiss and Edwards, 1980; Steyn et al., 1981; Turner and Aldridge, 1983; Gilland Steglich, 1987; Gill, 1994). Hypericin is believed to be formed by the linkage of two emodinanthrone units, with subsequent oxidation leading to theperylenequinone structure (Brockmann and Sanne, 1953). The mechanismand stage of formation of the chlorinated anthraquinones and hypericins is,however, uncertain at present (Figure.3).492.1.4 SYNTHESES OF 1-O-METHYLEMODIN (7), 7,7’-DICHLOROHYPERICIN(5) AND 2,2’,7,7’-TETRACHLOROHYPERICIN (6)7-Chloro-1-Q-methylemodin (3) (1.5 mg, 0.005 mmol) was dissolved in 5 mlof 0.015 M NaOH. Raney nickel (3.0 mg) was added, in one portion, to the stirredsolution. The reaction mixture was heated at reflux for 15 minutes, cooled to roomtemperature and filtered. The red filtrate was acidified to pH 7 with 6 N HCI. Theyellow solution was extracted three times with 50 ml portions of diethyl ether. Thecombined organic layers were dried and concentrated to a yellow solid. Recrystallization from 3:1 toluene:chloroform afforded 1.0mg of 1-O-methylemodin (7)(0.004 mmol, 75 % yield). The R1 of the product was 0.3 (5:1 benzene:ethyl acetate), 0.15 (3:3:1 toluene:chloroform:ethyl acetate) and 0.6 (70:20:8:2 chloroform:petroleum ether:ethyl acetate:methanol). The product was characterized by CIMSand 2D COSY 1H NMR spectra (Table 4).Hypericin (7 mg, 0.014 mmol) was dissolved, with stirring, in 10 ml of dryN,N-dimethylformamide. To the stirred, deep purple solution was added N-chlorosuccinimide (4 mg, 0.03 mmol). After four hours at ambient temperature, solventwas removed under reduced pressure, and the residual purple solid was dried,in vacuo, for 24 hours. This material was chromatographed on a column ofSephadex LH-20 with methanol:pyridine (9:1) as the eluant. Compound 6 (1.8mg from acetic acid, 20 % yield) was eluted first and was characterized by UV,LSIMS and 2D COSY 1H NMR spectra. The compound proved to be identicalwith the natural product in all respects. Compound 5 (4 mg from acetic acid, 5050% yield) was characterized by UV, LSIMS, 2D COSY 1H NMR and 3C NMRspectra. The 13C NMR shifts for compound 5 (Table 7) were determined usingAPT and HETCOR techniques. Several NOE experiments confirmed the presence of protons at C-2 and C-2’. Irradiation of the methyl protons resulted inan increase in each proton signal at C-2 and C-2’. Thus, the location of thechlorine atoms was at C-7 and C-7’. The compound proved to be identical withthe natural product in all respects.2.1.5 PHYSICAL AND CHEMICAL DATA FOR NATURAL PRODUCTSEmodin (1) (3 mg, 0.002 % yield, based on dry tissue weight) was obtainedas orange crystals from ethyl acetate: MP 256-257° C; UV (ethanol) ?. max (log E)253 (4.31), 265 (4.29), 289 (4.36), 438 (4.18); CIMS m/z [MH} 271 (94), 197 (100);CIMS mlz[MH] 270 (100). For 1H and 13C NMR data respectively, see Tables 4and 5.7-Chloroemodin (2) (78 mg, 0.06 % yield) was obtained as orange crystalsfrom ethyl acetate: MP 281-283° C; UV (ethanol) ? max (logE) 257 (4.24), 315(4.22), 325 (4.08), 437 (3.88), 504 (3.70); CIMS m/z [MH]307 (26), 305 (100);ElMS (70 eV) mlz [M] 306 (35), 304 (100). For 1H and 13C NMR data respectively,see Tables 4 and 5.7-Chloro-1-O-methylemodin (3) (82 mg, 0.06 % yield) was obtained asorange crystals from ethyl acetate: MP 289-291° C; UV (ethanol) 2. max (log E) 256(4.24), 286 (4.23), 423 (3.78); CIMS m/z[MH]321 (30), 319 (100); ElMS (70 eV)mIz[M] 320 (35), 318 (100). For 1H and ‘3C NMR data respectively, see Tables 4and 5.517-Chloro-1-O-methyl-o-hydroxyemodin (4) (1 mg, 0.008 % yield) was obtained as pink crystals from ethanol: MP > 2900 C; UV (ethanol) max (log E) 222(4.50), 250 (4.20), 300 (4.23), 434 (3.95), 452 (3.90), 490 (3.78), 525 (3.60); CIMSmIz[MH] 337 (3), 335 (12); (4) triacetate: CIMS m/z[MH] 480 (49), 478 (100), 463(35), 461 (79). For 1H NMR data, see Table 6.7,7’-Dichlorohypericin (5) (1.4 mg, 0.001 % yield) was obtained as purplecrystals from acetic acid: MP > 3500 C; UV (dimethylsulfoxide) 2 max (log E) 251(4.70), 294 (4.60), 332 (4.50), 388 (3.93), 485 (4.08), 553 (4.29), 597 (4.58); LSIMSm/z[M-H] 575 (9), 573 (38), 571 (54). For 1H NMR data, see Table 6.2,2’,7,7’-Tetrachlorohypericin (6) (0.8 mg, 0.0006 % yield) was obtained aspurple crystals from acetic acid: MP > 350° C; UV (dimethylsultoxide) max (log e)251 (4.70), 295 (4.60), 332 (4.50), 390 (3.90), 484 (4.06), 554 (4.27), 598 (4.56);LSIMS m/z[M-H] 645 (2), 643 (6), 641 (10), 639 (7). For 1H NMR data, seeTable 6.52Table 7. 3C NMR data for 7,7’-dichlorohvpericin (5) and hvericin.1Carbon 52 -. Hypericin2 Hypericin31,1’ 162.8 161.4 161.22,2’ 117.2 118.9 118.63,3’ 144.3 143.4 143.56,6’ 175.2 174.2 174.77,7’ 107.7 105.5 105.48,8’ 167.5 168.1 166.09,10 184.3 183.6 183.3la,9a 101.6 102.0 101.9ib,9b4 120.3 120.6 119.23a,3b4 120.4 120.7 120.76a,6b5 125.2 127.0 126.68a,lOa 109.6 108.3 108.38b,10b5 124.3 126.0 126.09c,lOc 120.8 121.2 121.23-Me, 3’-Me 23.8 23.6 23.61Chemical shifts (ö) are reported in ppm from TMS internal standard.2The spectra were recorded in DMSO-d6at 125 MHz.3The spectrum was recorded in DMSO-d at 90 MHz (Falk and Schmitzberger, 1992).4Values for C-i b/C-9b and C-3a/C-3b can be interchanged.5Values for C-6a/C-6b and C-8b/C-1 Ob can be interchanged.53CHAPTER 2: CHEMICAL PROPERTIES OF LICHEN QUINONOID PIGMENTS2.2 ISOLATION OF HETERODERMIA OBSCURATA NATURAL PRODUCTS2.2.1 INTRODUCTIONThe lichen Heterodermia obscurata can be found in temperate and tropicalareas of the world. It has been reported from East Asia, Southeast Asia, India,Nepal, Australia, New Zealand, Polynesia, Hawaii, Central and South America,Western Europe, Russia, East Africa and North America (Kurokawa, 1962; Culberson, 1966; Kurokawa, 1973; Swinscow and Krog, 1976, Hale, 1979; Swinscowand Krog, 1988; Trass, 1992). Of the 16 Heterodermia species known to containanthraquinones, Heterodermia obscurata is the only one to be found in temperateregions (Table 3).Heterodermia obscurata is abundant in deciduous forests of the easternUnited States (Figure 8; Appendix 2). It can be found from Florida to Pennsylvania and west to Texas and Wisconsin (Kurokawa, 1962; Hale, 1979). Isolatedpopulations of the lichen have also been reported from Arizona (Weber, 1963),southern Ontario (Brodo, 1988), New Brunswick (Gowan and Brodo, 1988), theBlack Hills of South Dakota (Wetmore, 1968) and Hawaii (Kurokawa, 1962).In Maryland, the lichen is fairly common in forested areas; in particular, itcan be found growing on the trunks of Black Oak (Quercus velutina), Ash (Fraxinus spp.), Black Walnut (Juglans nigra), Sweet Gum (Liquidambar styraciflua),Hickory (Caiya spp.) and occasionally on acidic rocks in hilly areas.I—54AAFigure 8. Heterodermia obscurata from Cedarville State Forest, Maryland.552.2.2 MATERIALS AND METHODSSeveral collections of Heterodermia obscurata were made from differentareas within Cedarville State Forest in southern Maryland. Two reference samples were deposited in the UBC Botany Department Herbarium. The lichen wascleaned of debris and air-dried.Immersion of the dried lichen in ice-cold diethyl ether for ten minutes produced a yellow extract which was examined by TLC (9:1 chloroform:methanol).The TLC patterns for all the individual collections were indistinguishable from oneanother, suggesting a chemical uniformity in H. obscurata populations within thepark. In fact, the TLC pattern was practically identical to the one reported byYosioka et al. (1 968c) for H. obscurata collected in central Japan.The lichen collections were combined and extracted, in succession, withice-cold diethyl ether, acetone and methanol. All the extracts were concentratedto minimal volumes and examined by TLC. 7-Chloroemodin (2) was the main constituent of the three extracts. The ether extract indicated the presence of severalyellow pigments, believed to be anthrones. This was supported by their UV fluorescence colors (dark red) and similarities in R, values with anthrones reportedearlier in H. obscurata (Yosioka et al., 1968c). The structural identities of the anthrones, however, could not be confirmed, as they were present in the extract inonly minute amounts. Six compounds were ultimately identified in the three extracts (TLC: 8:2 chloroform:methanol): emodin (1) (Rf 0.88); 7-chloroemodin (2)56(R1 0.72); 5,7-dichloroemodin (3) (R 0.32); flavoobscurin A (4) (R, 0.42); flavoobscurin B (5) (R, 0.20); 7,7’-dichlorohypericin (6) (Rf 0.30); and atranorin (R0.55). Compounds 1-5 were reported to be present in H. obscurata from Japan(Yosioka et al., 1968a, 1968b) (Figure 5); 7,7’-dichlorohypericin (6), also isolatedfrom Nephroma Iaevigatum, is a novel derivative of hypericin (Cohen and Towers,1 995a). The structures of the lichen compounds are shown in Figure 9.Following TLC examination, the yellow-orange extract from lOg of lichenwas concentrated to 25 ml, and filtered. The material remaining on the filterproved to be atranorin (180 mg, 1.8 % yield) based on a TLC comparison withcommercial atranorin (RfO.85for both substances in 12:12:1 chloroform:petroleum ether:methanol) and mixed melting point (198-200°C; no depression). Themixture of pigments (40 mg) was chromatographed on a column of SephadexLH-20 (100 g). A gradient of chloroform:methanol (8:2) to methanol was usedfor elution of the lichen compounds. The order of elution was: 1) emodin; 2) 7-chloroemodin; 3) 5,7-dichloroemodin; 4) flavoobscurin A; 5) flavoobscurin B;and, finally 6) 7,7’-dichlorohypericin. 7-Chlóroemodin was purified by recrystallization from hot ethyl acetate; the other compounds were purified by preparative TLC (8:2 chloroform:methanol). The immobile purple layer remaining on thecolumn was extruded and extracted with 100 ml of pyridine for 16 hours. Theextract was concentrated, and dried under vacuum, to give pure 7,7’-dichlorohy-pericin.570Emodin (1)CIH0CIHO7-Chloroemodin (2)5,7-Dichloroemodin (3)R = H; Flavoobscurin A (4)R = CI; Flavoobscurin B (5)7,7’-Dichlorohypericin (6)CH30CI 0CH3H3Figure 9. The anthraquinones of Heterodermia obscurata.582.2.3 RESULTS AND DISCUSSIONThe identity of compound 2 was confirmed by analysis of the CIMS and 2DCOSY 1H NMR spectra. The positive-ion CIMS spectrum shows peaks at 307 and305; the negative-ion CIMS spectrum shows peaks at 306 and 304. The 2D COSY1H NMR spectral data are shown in Table 8. The CIMS and 2D COSY 1H NMR dataare consistent with those previously reported (Bohman, 1968; Yosioka et al., 1968a;Yamamoto et al., 1968). A series of NOE experiments confirmed the identity of 2.Irradiation of the methyl protons resulted in the enhancement of the H-2 and H-4proton signals. Irradiation of the hydroxyl proton at C-6 resulted in a signal intensityenhancement of H-5. 13C NMR assignments were obtained from APT and HETCORexperiments, and are shown in Table 9.Compound 3 proved to be 5,7-dichioroemodin on the basis of the MS and 2DCOSY 1H NMR spectra. The LSIMS spectrum shows peaks at 341, 339 and 337(relative intensity 1:2:3). This is indicative of two chlorine atoms in the molecule.The 2D COSY 1H NMR spectral data are shown in Table 8. The data agree wellwith earlier results (Yosioka et al., 1 968a; Lam et al., 1972). An NOE experiment confirmed the identity of 3. Irradiation of the methyl protons produced a correspondingincrease in the proton signal intensity of H-2 and H-4.Emodin was characterized by UV, CIMS and 2D COSY 1H NMR spectra (Table8). Several NOE experiments confirmed that 1 was emodin. Irradiation of the methylprotons resulted in increases in the proton signal intensities of H-2 and H-4. Irradia59Table 8. 1H NMR data for emodin (1), 7-chloroemodin (2) and5.7-dichioroemodin (3)•12Proton 2 32 7.19,s 7.18,s 6.95,s4 7.57, s 7.61, s 7.43, s5 7.26, d (2.5) 7.43, s7 6.69,d(25)1-OH 11.97,s 11.72,s6-OH 13.66, s8-OH 12.20, s 12.80, s3-Me 2.48, s 2.48, s 2.40, s1Chemical shifts (6) are reported in ppm from TMS internalstandard. The coupling constants are given in Hz.2The spectra were recorded inMe2CO-d6at 400 MHz.60Table 9. 13C NMR data for emodin (1), 7-chloroemodin (2), 5,7-dichloroemodin (3)and flavoobscurin B (5)1Carbon 1 12 2 22 3 32 51 161.9 158.7 161.4 158.7 160.0 158.7 160.41’ 160.42 124.8 120.2 124.2 120.1 124.6 120.1 122.12’ 122.13 149.3 142.4 148.3 142.4 145.4 142.4 146.03’ 146.04 121.2 125.3 120.4 125.3 119.4 125.3 117.24’ 117.25 109.7 111.9 108.5 113.2 119.3 119.4 115.45’ 115.46 167.0 160.4 163.1 160.8 168.5 161.2 166.66’ 166.67 108.6 106.7 121.0 112.9 122.7 114.2 117.37’ 117.38 165.9 160.2 162.8 160.6 160.4 158.7 161.68’ 161.69 191.9 189.3 189.0 187.4 210.9 185.5 188.810 182.2 161.7 180.8 161.7 182.4 163.0 188.8la 116.0lb 139.54a (9b) 134.0 131.5 132.8 131.5 127.0 129.6 139.58a iio.o 110.0 110.8 111.3 1li.0 112.6 112.98b 139.69a 114.4 107.1 113.4 107.1 113.6 107.1 116.09c 30.8lOa 136.6k 132.8 132.6k 130.9 133.6k 131.3 112.9lOb 139.6lOc 30.83-Me 21.8 21.6 21.5 21.63-Me’_______ _______ _______ _______ _______ _______21.61Chemical shifts (8) are reported in ppm from TMS internal standard. The spectrawere recorded in DMSO-d6at 125 MHz.2Values calculated according to the methods described in Silverstein et al. (1991).3Values for C-8a and C-ga can be interchanged.4Values for C-i Oa and C-4a can be interchanged.5Values for C-i b/9b and C-8b/i Ob can be interchanged.61Table 10. 1H NMR data for flavoobscurin A (4), flavoobscurinB (5) and 7.7’-dichlorohvDericin (6).1Proton 42 52 62,2’ 6.54, s 6.68, s 7.42, s4,4’ 5.11,s 5.74,s5,5’ 6.96, s7,7’10,10’ 4.66, d (3.0) 4.95, s410,10’ 4.72, d (3.0)1,1’-OH6,6’-OH8,8’-OH3-Me, 3’-Me 2.32, s 2.32, s 2.65,s1Chemical shifts (6) are reported in ppm from TMS internalstandard. The coupling constants are given in Hz.2The spectra were recorded inMe2CO-d6at 400 MHz.3The spectrum was recorded in DMSO-d at 400 MHz.4No attempt has been made to assign stereochemistry.62tion of the C-8 hydroxyl proton also produced a proton signal enhancement of H-7.Compounds 4 and 5 are chlorinated bianthrones, originally isolated by Yosiokaet al. (1968b) (Figure 9). Nonhalogenated bianthrones have been found in a crinoid,Lamprometra palmata (Rideout and Sutherland, 1985) and in the fungi Aspergilluschevalieri (Bachmann et al., 1979) and A. wentli (Assante et al., 1980). The structuresof 4 and 5 were proven by LSIMS, CIMS and 2D COSY ‘H NMR spectra. The twocompounds produce characteristic fragmentation patterns in their mass spectra. Thenegative-ion LSIMS spectrum of 4 shows peaks corresponding to the parent bianthrone structure, as well as the two nonsymmetrical monomers. The negative-ionLSIMS spectrum of 5 also shows the parent peak, along with a single set of monomeric fragments. Only the monomeric fragments can be seen in the CIMS spectra ofthe two compounds; they do display, however, the same overall patterns seen in thetwo LSIMS spectra. The 2D COSY ‘H NMR chemical shifts for 4 and 5 are shown inTable 10. Table 9 lists the 13C NMR shifts for flavoobscurin B, which were not reportedby Yosioka et al. (1968b).7,7’-Dichlorohypericin (Figure 9) gave a negative-ion LSIMS spectrum with parent peaks at 575, 573 and 571 (relative intensity 1:2:3). This is indicative of two chlorine atoms in the compound. This conclusion was supported by the 2D COSY ‘H NMRdata (Table 10). An NOE experiment also confirmed the identity of compound 6. Irradiation of the methyl protons produced an increase in the proton signal intensity of H-2(H-2’).63Many lichen genera produce anthraquinones, and anthrones have been detectedin several species by TLC (Bohman, 1968; Yosioka et al., 1968c; Kurokawa, 1973;Steiner et al., 1974). Bianthrones are probably formed by oxidative coupling ofanthrones. A TLC, performed a few hours after sample collection, clearly showedthe presence of bianthrones and 7,7’-dichlorohypericin. Thus, it seems unlikelythat these natural products are exclusively artifacts generated during extractionand sample manipulation.2.2.4 SYNTHESES OF 7-CHLOROEMODIN (2) AND 5,7-DICHLOROEMODIN (3)Emodin (40 mg, 0.15 mmol) was added to 50 ml of dry N,N-dimethylformamide. The red solution was allowed to stir at ambient temperature until all theemodin had dissolved (40 minutes). N-Chlorosuccinimide (40 mg, 0.30 mmol) wasadded, in one portion, to the stirred solution. The reaction mixture was maintainedat room temperature for 24 hours. The solvent was removed, under reduced pressure, to give an orange solid. The solid was dried, in vacuo, for 24 hours. Columnchromatography of the crude mixture on Sephadex LH-20 (8:2 chloroform:methanol) afforded (in order of elution) emodin (1) (8 mg from ethyl acetate, 20 % recovery), 7-chloroemodin (2) (22 mg from ethyl acetate, 48 % yield) and 5,7-dichloro-emodin (3) (12 mg from methanol, 24 % yield). All compounds were characterizedby UV, CIMS, ElMS, 1H NMR and ‘3C NMR spectra. They were found to be identical with the natural products in all respects.642.2.5 PHYSICAL AND CHEMICAL DATA FOR NATURAL PRODUCTSEmodin (1) (3 mg, 0.03 % yield, based on dry tissue weight) was obtainedas orange crystals from ethyl acetate: MP 255-256°C; UV (ethanol) ? max (log E)262 (4.30), 288 (4.34), 434 (4.20); CIMS mlz [MH] 270 (100). For 1H and 13C NMRdata respectively, see Tables 8 and 9.7-Chloroemodin (2) (12 mg, 0.12 % yield) was obtained as orange crystalsfrom ethyl acetate: MP 280-282°C; UV (ethanol) 2i max (log E) 257 (4.24), 315(4.22), 325 (4.08), 437 (3.88), 504 (3.70); CIMS m/z[MH] 307 (38), .305 (100);CIMS m/z[MH] 306 (33), 304 (100). For 1H and 13C NMR data respectively, seeTables 8 and 9.5,7-Dichloroemodin (3) (1 mg, 0.01 % yield) was obtained as red crystalsfrom methanol: MP 268-270°C; UV (ethanol) max (log ) 262 (4.43), 320 (4.20),457 (4.00), 524 (3.90); LSIMS m/z[M-H] 341 (12), 339 (25), 337 (35); CIMS mlz[MH] 343 (7), 341 (38), 339 (55); CIMS m/z [MH] 342 (12), 340 (68), 338 (100).For ‘H and 13C NMR data respectively, see Tables 8 and 9.Flavoobscurin A (4) (2 mg, 0.02 % yield) was obtained as lemon-yellowcrystals from acetic acid: MP > 350°C; UV (ethanol) 2. max (log E) 273 (4.25), 410(4.30); LSIMS mlz[M-H] 615 (2), 613 (4), 611(5), 326 (19), 324 (75), 322 (100),290(9), 288 (18); CIMS m/z[MH]329 (17), 327 (80), 325 (100), 293 (47), 291(86); CIMS rnlz[MH] 327 (7), 325 (27), 323 (33), 290 (40), 288 (66). For ‘H NMRdata, see Table 10.65Flavoobscurin B (5) (3 mg, 0.03 % yield) was obtained as lemon-yellowcrystals from acetic acid: MP> 350°C; UV (ethanol) 2. max (log E) 276 (4.20), 403(4.26); LSIMS m/z[M-H] 651 (2), 649 (6), 647 (12), 645 (9), 327 (7), 325 (26), 323(32); CIMS m/z[MH]329 (11), 327 (54), 325 (79); CIMS m/z [MHJ 327 (8), 325(31), 323 (40). For 1H and 13C NMR data respectively, see Tables 10 and 9.7,7’-Dichlorohypericin (6) (0.8 mg, 0.008 % yield) was obtained as purplecrystals from acetic acid: MP > 350°C; UV (ethanol) 2 max (log E) 259 (4.70), 292(4.60), 332 (4.51), 485 (4.08), 552 (4.26), 594 (4.56); LSIMS m/z[M-H] 575 (19),573 (39), 571 (50). For 1H NMR data, see Table 10.66CHAPTER 3: BIOSYNTHETIC STUDIES OF QUINONOID PIGMENTS IN N.LAEVIGATUM3.1 INTRODUCTIONIn order to determine if the lichen anthraquinones are derived from acetate through the polyketide pathway, sodium [2-14C]acetate and sodium [1-13Cjace-tate were administered to Nephroma Iaevigatum, maintained in aqueous culture, intwo separate experiments. In a third experiment, sodium chIoride was fed to thelichen, in the hope of observing in situ chlorination of endogenous anthraquinones,or their precursors.3.2 MATERIALS AND METHODSLichen thalli of Nephroma Iaevigatum were collected from shoreline rockson Gabriola Island, British Columbia in October, 1994. The lichen (100 g) wascarefully cleaned of moss, soil, infected or damaged lichen thalli and other debris,and washed several times with sterile distilled water. The lichen (20 g) was placedin each of three plastic dishes (245 x 245 x 20 cm). In three different experiments,the lichen was incubated with aqueous solutions of sodium [2-14C]acetate (50 microcuries, 99.9 atom % ‘C), sodium [1-13C}acetate (0.12 M solution, 99 atom %13C) and sodium36chloride (25 microcuries, 99 atom % 36C1). Each isotope wasdissolved in a sufficient amount of sterile distilled water to thoroughly moisten thelichen, but not submerge it. The total liquid (100 ml) was absorbed within a fewminutes. The incubation experiments were maintained in an indoor greenhouse at27°C under constant conditions of light/dark cycles (16 hours light/8 hours dark)67and temperature. After five days, the lichen was harvested, dried and weighed.The isolation and characterization of the labelled anthraquinones followthe same procedures utilized in the initial identification of compounds in Nephroma Iaevigatum (Chapter 2). The primary constituent in the mixture of pigmentsobtained from the sodium [2-14C]acetate incorporation experiment was 7-chioro-emodin (2) (TLC; R, 0.5; 9:1 chloroform:methanol). The TLC pattern revealedthe presence of two additional compounds, which were subsequently isolatedand characterized as: emodin (1) (Rf 0.8) and 7-chloro-1-Q-methylemodin (3)(Rf 0.7). Five products were identified in the pigment mixture obtained from thesodium [1-3C}acetate incorporation experiment: 7-chloro- 1-O-methylemodin (3)(R,0.7), 5-chloro-1-Q-methylemodin (5) (R, 0.65), 5-chloroemodin (6) (R0.55),5-chloro-1 -Q-methyl-a-hydroxyemodin (7) (Rf 0.3) and 5-chloro-w-hydroxyemo-din (8) (R, 0.25). Finally, the pigment mixture from the sodium36chloride incorporation experiment contained 7-chloroemodin (2) (R 0.5), 7-chloro-1 -0-methyl-emodin (3) (R 0.7) and 7-chloro-1 -0-methyko-hydroxyemodin (4) (R, 0.35).The structures of these compounds are shown in Figure 10. The lichen was extracted successively with acetone and methanol. After TLC examination, thecombined extracts were concentrated to a brown-red solid. The solid was purified by column chromatography on Sephadex LH-20, using a gradient of chloroform:methanol (9:1) to methanol. Fractions (20 ml) were collected and analyzed by TLC. All compounds were further purified by preparative TLC (8:268OH 0 OHHO%%•— CH30HOR = Cl, R’ = H; 7-Chloro-1-O-methylemodin (3)R H, R’ = Cl; 5-Chloro-1-O-methylemodin (5)R Cl, R = H, R” = CH3; 7-Chloro-1-O-methyl-o-hydroxyemodin (4)R = H, R’ = Cl, R” = CH3; 5-Chloro-1-O-methyl-0-hydroxyemQdin (7)R = H, R’ = Cl, R” = H; 5-Chloro-o) -hydroxyemodin (8)Emodin (1)R 0CH3R = Cl, R’ = H; 7-Chloroemodin (2)R = H, R’ = Cl; 5-Chloroemodin (6)R = Cl, R’ = CI; 5,7-Dichloroemodin (9)R 0Figure 10. Anthraquinones from isotope labelling experiments with N. Iaevigatum.0)CDTable11.‘HNMRdataforemodin(1),7-chloroemodin(2),7-chloro-1-O-methylemodin(3),7-chloro-1-O-methyl-o-hydroxyemodin(4),5-chloro-1-Q-methylemodin(5),5-chloroemodin(6),5-chloro-1-O-methyl-o)-hydroxyemodin(7,and5-chloro-o-hydroxyemodin(8)fromisotopelabellingexperiments.’Proton1(‘4 C)22(’4C)2(36Cl)3(’4C)23(’C)43(36C1)4(36Cl)25(’3C)46(’3C)47(’3C)48(’3C)427.21,S7.09,s7.20,S7.50,S7.48,S7.45,s6.90,s7.38,s7.08,s7.46,s7.12,s47.57,s7.49,s7.62,s7.68,s7.62,s7.68,s7.80,s7.58,s7.38,s7.73,s7.54,557.24,d(2.5)7.11,s7.48,s7.29,s7.25,s7.35,s7.50,s76.65,d(2.5)6.78,s6.68,s6.78,s6.63,s1-OH6-OH8-OH1-OMe4.05,s4.00,s4.03,s4.00,s3.92,s3.92,s3-CH2OH4.73,d(5.0)4.63,d(5.0)4.54,d(5.O)3-Me2.48,s2.48,s2.50,S2.50,S2.50,52.50,s2.47,s2.40,S‘Chemicalshifts(6)arereportedinppmfromTMSinternalstandard.ThecouplingconstantsaregiveninHz.Theradioactive/stableisotopelabelorisotopeusedinthefeedingstudyisshowninparentheses.2 ThespectrawererecordedinDMF-dat400MHz.3 ThespectrawererecordedinMe2CO-d6at400MHz.4 ThespectrawererecordedinDMSO-d6at400MHz.5 Thehydroxymethylprotonsarecoupledtoabroadhydroxylproton.IfthehydroxylprotonisexchangedbyadditionofasmallamountofMeOH-d4,thissignalbecomesasinglet.01 Yieldsarebasedonlichendryweight.2 Disintegrationsperminute.3 TotaIradioactivity(microcuries).4 Specificactivity.5 Percentage(absolute)incorporation([DPMofisolatedproductlDPMofisotopefedtolichen]%100).Table12.PropertiesofradiolabelledanthrauinonesisolatedfromNehromaIaeviqaturn.CompoundIsotopeYield(%)1DPM2DPM/mg1i,C,3rnC/mM4%Incorp.5114C2mg(0.02)8,18840940.0040.000540.01214C30mg(0.23)112,71637570.050.000510.1236Cl5mg(0.06)18,20036380.010.000610.04314C22mg(0.17)23,80410820.010.000140.02336C15mg(0.06)2,0104020.0010.000060.00471chloroform:methanol), and repeatedly recrystallized from a suitable solvent untilpurity (> 99 %) could be established on the basis of reversed-phase HPLC.Radiolabelled compounds were recrystallized to constant specific activity, andpurity checked by reversed-phase HPLC. All compounds were characterizedby UV, MS, 1H NMR and 13C NMR spectra.Autoradiographs of the radiolabelled compounds were made by exposing2D silica gel TLC plates of anthraquinones ‘and isotope to X-ray film, after solventdevelopment in chloroform:methanol (8:2) and acetone:acetic acid (8:2). After twomonths exposure, the X-ray film was developed and the radioactive “halos”, corresponding to either 14C or36C1-labelled anthraquinones, marked (Figure 1 1). Radioactive “halos” representing sodium36chloride or sodium [2-14C]acetate wereclearly distinguishable from the radiolabelled anthraquinones. Development of the2D TLC plate in either solvent system did not result in the migration of isotopicallylabelled reagent from the original point of application. Furthermore, there wasno evidence for the presence of either isotope at the locations where the samplesof radiolabelled anthraquinones were applied to the TLC plate. It was thereforeconcluded that the purified radiolabelled anthraquinones were free from any contamination by the isotopes used in the lichen tracer studies.3.3 RESULTS AND DISCUSSIONThe identity of 2 was established by 1H NMR (Table 11) and mass spectra.The spectral data are consistent with our previous results (Cohen and Towers,721 995a, 1 995b). Table 12 provides the specific activities and percentage incorporations for 14C and36C1-labelled 7-chloroemodin (2).Emodin (1) was characterized by 1H NMR (Table 11) and mass spectra.The spectral data are indistinguishable from those of authentic material (Cohenand Towers, 1 995a, 1 995b). The specific activity and incorporation levels for14C-Iabelled emodin (1) are shown in Table 12. The amount of emodin producedin the sodium [2-’4C]acetate incubation experiment was insufficient for a 13C NMRspectrum to be taken.7-Chloro-1-Q-methylemodin (3) was identified by 1H NMR (Table 11), 13CNMR (Table 13) and mass spectra. The results of an NOE experiment performedon compound 3 were consistent with the assigned structure. Irradiation of the H-2proton resulted in an enhancement of the methoxy protons at C-i. The percentageincorporation of 14C label into compound 3 was 20 % of the level for 7-chioroemodin,and the specific activity of14C-Iabelled 7-chloro-1-O-methylemodin was also 20 %of the values reported for 7-chioroemodin and emodin (Table 12).The 13C NMR spectrum of 7-chloro-1-O-methylemodin isolated from the feeding experiment shows specific incorporation at carbon atoms 1, 3, 4a, 6, 8, 9 andlOa (Table 13). This is entirely consistent with its formation from an octaketide precursor, itself derived from acetate. The ‘3C isotopic enrichments were measured bycomparing the peak intensities in both the natural abundance and enriched spectraafter normalization (Casey et al.,1978). The enrichments values are consistent with73Table 13. 13C NMR data and isotope enrichments for7-chloro-1 -O-methvlemodin (3)•1Carbon Chemical Shift (6) % Enrichment21 160.0 3.12 120.5 0.73 147.8 3.74 120.0 0.74a 131.8 5.25 118.0 0.66 161.0 4.97 106.4 0.08 160.2 3.18a 112.4 0.09 186.2 6.09a 113.5 0.010 182.0 1.9lOa 135.6 3.8OMe 56.5 0.0Me 21.8 -0.51The lichen was fed 0.12 M sodium [1-13C} acetate. Thespectrum was recorded in DMSO-d6at 125 MHz. Chemical shifts (6) are reported in ppm from TMS internalstandard.2Percentage 13C enrichments (greater than 1.1 % naturalabundance) were calculated from the ratio in peak heightobtained from the 13C NMR spectra of labelled and unlabelled products, respectively.74Table 14. C NMR data for 7-chloroemodin (2),7-chloro-1 -O-methylemodin (3) and 5-chloro-1 -O-methvlemodin (5)1Carbon 2(14C) 3(14C) 3(13C) 5(13C)1 161.4 160.0 160.0 160.22 124.0 120.3 120.5 120.03 148.5 148.1 147.8 145.84 120.4 120.1 120.0 119.85 108.6 106.6 106.4 118.66 163.1 160.7 161.0 161.07 121.3 119.8 118.0 112.68 162.8 160.5 160.2 160.29 191.0 186.4 186.2 191.810 181.7 182.0 182.0 183.64a2 133.2 131.1 131.8 130.68a3 110.1 109.8 112.4 112.69a3 114.2 113.1 113.5 113.410a2 133.0 134.4 135.6 134.8OMe 56.4 56.5 56.5Me 21.6 21.7 21.8 21.81Chemical shifts (ö) are reported in ppm from TMSinternal standard. The spectra were recorded inDMSO-d8at 125 MHz. The isotope label is shownin parentheses.2Values for 4a and 1 Oa can be interchanged3Values for 8a and 9a can be interchanged.75levels obtained from biosynthetic studies of anthraquinones in Penicillium isiandicum (Casey et al., 1978), perylenequinones in Pyrenochaeta terrestris (Kurobaneet al., 1981) and anthraquinones in Dermocybe (Gill and Giménez, 1990a, 1990b).5-Chloro-1-O-methylemodin (5) had a mass spectrum similar to that of the7-chloro isomer; the ‘H NMR spectrum was quite different however. The locationof the chlorine at C-5 was based on the chemical shift for H-7 (6.78 ppm) whichis characteristic for an aromatic proton situated between two aromatic hydroxylgroups (Table 11). The location of the methoxy group was determined by an NOEexperiment; irradiation of the methoxy protons at C-i resulted in an increase inthe signal intensity for the H-2 proton.5-Chloro-1-Q-methylemodin had previously been isolated from the fungusPhialophora alba (Ayer and Trifonov, 1994). Although two other 5-chloro-substi-tuted anthraquinones are known from the fungal genus Dermocybe (Steglich et al.,1969), they have not previously been reported from lichens collected in the field.7-Chloro- 1-O-methyko-hydroxyemodin (4), a new metabolite previously isolated by us (Cohen and Towers, 1 995a), was also obtained from our lichen feedingexperiments. We could not, however, isolate sufficient quantities for accurate measurements of isotope enrichments. The structure of 4 was proven by 1H NMR, NOEexperiments and mass spectra. The location of the methoxy protons at C-i wasconfirmed by their irradiation, which produced an increase in the 1H NMR signalintensity of the H-2 proton.765-Chloroemodin (6) was confirmed by ‘H NMR and mass spectra. TheCIMS mass spectra of 6 and 6 triacetate indicated that it was a monochloroemodin. The ‘H NMR spectrum showed an aromatic signal consistent with an H-7proton (6.70 ppm) (Table 11).5-Chloro-1-O-methyl-c.o-hydroxyemodin (7) was confirmed by 1H NMR,NOE experiments and mass spectra. The CIMS mass spectrum was similarto that of 4; however, the presence of an H-7 proton shift (6.78 ppm) indicatedthat the chlorine must be at C-5 (Table 11). The position of the methoxy protonswas shown by an NOE experiment; irradiation of the methoxy protons resultedin an increase in the ‘H NMR signal intensity of H-2.5-Chloro--hydroxyemodin (8) was confirmed by ‘H NMR and mass spectra. The ElMS mass spectrum gave a parent molecular ion corresponding to themolecular mass of 8. The presence of an H-7 proton shift (6.63 ppm) was readilyapparent from the ‘H NMR spectrum, and indicated that the chlorine must be atC-5 (Table 11).3.4 PHYSICAL AND CHEMICAL DATA FOR NATURAL PRODUCTS3.4.1 PRODUCTS FROM SODIUM [2-’4C]ACETATE INCORPORATIONEmodin (1) (2 mg, 0.02 % yield, based on dry tissue weight) was obtainedas orange crystals from ethyl acetate: MP 256-257°C; UV (ethanol) 2. max (log e)253 (4.31), 265 (4.29), 289 (4.36), 438 (4.18); CIMS mlz [MH] 271 (100), 242(77). For ‘H NMR data, see Table 11.777-Chloroemodin (2) (30 mg, 0.23 % yield) was obtained as orange crystalsfrom ethyl acetate: MP 281 -283°C; UV (ethanol) max (log E) 257 (4.24), 315(4.22), 325 (4.08), 437 (3.88), 504 (3.70); CIMS m/z[MH] 307 (26), 305 (100);ElMS (70 eV) m/z [M] 306 (35), 304 (100). For ‘H and ‘3C NMR data respectively,see Tables 11 and 14.7-Chloro-1-O-methylemodin (3) (22 mg, 0.17% yield) was obtained asorange crystals from ethyl acetate: MP 289-291°C; UV (ethanol) 2. max (log £) 256(4.24), 286 (4.23), 423 (3.78); CIMS m/z[MH]321 (30), 319 (100); ElMS (70 eV)m/z [M] 320 (35), 318 (100). For ‘H and ‘3C NMR data respectively, see Tables 11and 14.3.4.2 PRODUCTS FROM SODIUM [1-’3C]ACETATE INCORPORATION7-Chloro-1 -O-methylemodin (3) (3 mg, 0.02 % yield) was obtained as orangecrystals from ethyl acetate: MP 288-290°C; UV (ethanol) 2 max (log E) 256 (4.24),286 (4.23), 423 (3.82); CIMS m/z[MH] 321 (33), 319 (100). For ‘H and ‘3C NMRdata respectively, see Tables 11, 13 and 14.5-Chloro-1-Q-methylemodin (5) (3 mg, 0.02 % yield) was obtained as orangecrystals from ethyl acetate: MP 252-253° C; UV (ethanol) ? max (log E) 228 (4.30),259 (4.33), 314 (4.25), 440 (4.02); CIMS m/z[MH]321 (33), 319 (100). For1Hand‘3C NMR data respectively, see Tables 11 and 14.5-Chloroemodin (6) (2 mg, 0.01 % yield) was obtained as orange crystals fromethyl acetate: CIMS mlz [MH] 307 (33), 305 (100); ElMS m/z [M] 306 (22), 304(100); 5-chloroemodin triacetate: ElMS mIz[M] 390 (9), 388 (27), 348 (9), 346 (25),306 (51), 304 (100). For 1H NMR data, see Table 11.785-Chloro-1-O-methyl-w-hydroxyemodin (7) (2 mg, 0.01 % yield) was obtainedas salmon crystals from methanol: UV (ethanol) ? max (log E) 218 (4.50), 257 (4.22),322 (4.08), 496 (3.56); CIMS m/z[MH] 337 (33), 335 (100); 5-chloro-1-Q-methyl-(o-hydroxyemodin triacetate: CIMS m/z [MH] 463 (33), 461 (100); ElMS m/z [M] 420(20), 418 (46), 378 (18), 376 (55), 336 (2), 334 (7), 318 (42), 316 (100), 306 (20),304 (51). For ‘H NMR data, see Table 11.5-Chloro-w-hydroxyemodin (8) (2 mg, 0.01 % yield) was obtained as orangecrystals from methanol: ElMS m/z[M] 322 (33), 320 (100). For 1H NMR data, seeTable 11.3.4.3 PRODUCTS FROM SODIUM36CHLORIDE INCORPORATION7-Chloroemodin (2) (5 mg, 0.06 % yield) was obtained as orange crystalsfrom ethyl acetate: MP 281-282°C; UV (ethanol) ? max (log E) 257 (4.24), 315(4.22), 325 (4.10), 437 (3.88), 504 (3.80); CIMS m/z[MH] 307 (30), 305 (100).For1HNMR data, see Table 11.7-Chloro-1-O-methylemodin (3) (5 mg, 0.06 % yield) was obtained asorange crystals from ethyl acetate: MP 289-291°C; UV (ethanol) ? max (log E)256 (4.24), 286 (4.22), 423 (3.80); CIMS mlz [MH] 321 (33), 319 (100). For‘H NMR data, see Table 11.7-Chloro-1-O-methyl-w-hydroxyemodin (4) (1 mg, 0.01 % yield) was obtamed as pink crystals from ethanol: MP > 290°C; UV (ethanol) ?. max (log E)79222 (4.50), 250 (4.20), 300 (4.23), 434 (3.95), 452 (3.90), 490 (3.78), 525 (3.60).CIMS m/z[MH] 337 (3), 335 (12); 7-chloro-1-O-methyl-o-hydroxyemodin tnacetate: CIMS m/z[MH] 480 (49), 478 (100), 463 (35), 461 (79). For ‘H NMRdata, see Table 11.80Rgure 11. Autoradiographs of 2D TLC of radiolabelled compounds.81CHAPTER 4: BIOHALOGENATION STUDIES OF QUINONOID PIGMENTS4.1 INTRODUCTIONSince the discovery of the first halogenating enzymes in the early 1 950s,our knowledge about the physiological roles, molecular structures, and mechanisms of action of haloperoxidases has greatly expanded (Neidleman and Geigert,1986; van Pee, 1990; Franssen and van der Plas, 1992; Butler and Walker, 1993).Three classes of haloperoxidases are now known: chloroperoxidases (which canchlorinate, brominate, and iodinate various substrates); bromoperoxidases (bromination and iodination), and iodoperoxidases (iodination only). The first systematicinvestigation of a haloperoxidase began with the isolation and mechanistic study ofa chloroperoxidase from the fungus Caldariomyces fumago (Morris and Hager,1966; Hager et al., 1966). In the intervening years, additional chloroperoxidaseshave been isolated from bacteria and fungi; bromoperoxidases from bacteria, marine algae, marine worms and sea urchins; and iodoperoxidases from mammals,birds, higher plants and brown algae (Neidleman and Geigert, 1986; Franssen andvan der Plas, 1992). Fungi appear to contaih only chloroperoxidases, although basidiomycetes have yet to be examined in any detail. The prevalence of chlorinatednatural products in fungi (Gribble, 1992) is consistent with the presence of chloroperoxidases; although the same enzymes can also utilize higher halogens, thescarcity of brominated or iodinated compounds in fungi probably reflects the scarcity of these halogens in terrestrial substrates. The single report of a lichen haloperoxidase is that of a bromoperoxidase isolated from Xanthoria parietina (Plat82et al., 1987). The investigators found that the enzyme contained vanadium,essential for catalytic activity, and was remarkably thermostable, maintainingfull enzymatic activity at 50°C. In addition, the bromoperoxidase had a high affinity for bromide ion and was only active at low concentrations. The enzymewas also inhibited by chloride and fluoride ions, and exhibited a pH optimum atpH 5.5. Considering the absence of brominated compounds in lichens, and theubiquity of chlorinated substances, it is surprising that the lichen would producea bromoperoxidase, rather than a chloroperoxidase. Interestingly, a bromoperoxidase isolated from the green alga Penicillus capitatus was later shown to become a chloroperoxidase at a lower pH (Manthey and Hager, 1989; Franssenand van der Plas, 1992). Had the vanadium-containing bromoperoxidase fromXanthoria parietina been studied at lower pH, it might also have shown chloroperoxidase activity (Soedjak and Butler, 1990).4.2 MATERIALS AND METHODSSince Nephroma Iaevigatum produced several chlorinated anthraquinones,and there was evidence from the sodium36chloride incorporation experiment forin situ chlorination, it seemed worthwhile to try to identify a chioroperoxidase inthe lichen,Ultimately, a semi-purified chloroperoxidase preparation was obtained fromthe lichen in the following manner. The lichen (400 g) was homogenized in ablender with 2 liters of 0.1 M potassium phosphate buffer, pH 5.8. The homo83Dimedon AssayCI- chioroperoxidase+ Cl+H20MCDReaction mixture: (2 ml) 0.2 M potassium phosphate buffer, pH 3.0(0.1 ml) 50 mM potassium chloride(0.1 ml) 2 mM MCD(0.1 ml) 10 mM hydrogen peroxide.(10 jil) chioroperoxidaseProcedure:1. Mix all the ingredients, except hydrogen peroxide, in a vial.2. Initiate chlorination reaction by the addition of peroxide.3. Assay mixture continuously in a quartz UV cuvette. Monitorthe decrease in absorbance (292 nm) corresponding to theconsumption of MCD.4. The MCD reaction is the standard assay used to assign theactivity of a haloperoxidase enzyme.Bradford AssayProcedure:1. Dissolve 100 mg Coomassie Blue G-250 in 50 ml 95% ethanol. To this solution add 100 ml 85% (w/v) phosphoric acid.Dilute the resulting solution to a final volume of 1 liter.2. Prepare protein solutions (bovine serum albumin) by dissolving an appropriate amount of BSA in 0.15 M NaCl. Preparestandard solutions containing 10 to 100 ig protein per 0.1ml.3. Pipet 0.1 ml of each protein solution into test tubes and adjust volume, if necessary, to 0.1 ml with 0.15 M NaCI.4. Add 5 ml of Coomassie Blue Reagent, mix contents well andmeasure absorbance (595 nm) after 2 minutes and 45 minutes in a quartz cuvette (against reagent blank containingdye and buffer)5. Plot weight of protein against corresponding absorbances.Determine the protein content in unknown samples fromthe standard curve.DCDFigure 12. Dimedon and Bradford assays.84commercial CPO- no enzyme+ Cl+H20CH3 N. Iaevigatum CPOcommercial CPOno enzyme+ Cr+H20N. Iaevigatum CPC7-Chloroemodin (2)Experimental protocols:1. Control reaction: 3 mg (10 j.imol) of 1 or 2 and 100 mg (1.4 mmol) KCI wasplaced in 34 ml of 6:4 0.2 M potassium phosphate-DMF buffer, pH 3.4. Tothe stirred solution was added 17 tl (280 jimol) of 50 % aq. H20The reaction mixture was maintained at ambient temperature, checked periodically by TLC and stopped after 68 hours by extraction of dried productswith ethyl acetate. The dried extract was fractionated by column chromatography and compounds purified by preparative TLC (8:2 chloroform:methanol).2. Commercial (fungal) chloroperoxidase (CPO) reaction: 3 mg (10 jimol) of1 or 2, 100 mg (1.4 mmol) KCI, 6 units CPO and 17 jii (280 pmol) of 50 %aq. H20 in 34 ml of 6:4 0.2 M potassium phosphate-DMF buffer, pH 3.4.Commercial chloroperoxidase obtained from Sigma Chemical Co.3. Nephroma Iaevigatum chioroperoxidase (CPC) reaction: 3 mg (10 j.tmol)of 1 or 2, 100 mg (1.4 mM) KCI, 6 units lichen CPO and 17 jil (280 pmol)of 50 % aq. H20 in 34 ml of 6:4 0.2 M potassium phosphate-DMF buffer,pH 3.4.4. All products were characterized by MS and 1H NMR spectra.Emodin (1)CIFigure 13. Biohalogenation experiments.ReactionErnodin(1)7-Chloroemodin(2)5-Chloroemodin(3)5,7-Dichloroemodin(4)Control(1)2++(40hours)+(40hours)+(68hours)Control(2)2+-Corn.CPO(l)++(20hours)+(20hours)+(68hours)Corn.CPO(2)++(40hours)N.I.CPO(i)++(20hours)--N.I.CPO(2)+-‘Symbolkey:+(present)or-(absent).Thetimewhentheproductfirstappeared(byTLC)isshowninparentheses.AllreactionproductscharacterizedbyUV,ElMSand1 HNMRspectra.2 Noenzyme.3 Commercialfungalchloroperoxidase(Caldariomycesfumago);SigmaChemicalCo.4 NephromaIaevigatumsemi-purifiedchloroperoxidase.U)coTable15.Reactionproductsfromchlorinationexperimentswithemodin(1)and7-chloroemodin(2).’86genate was filtered through cheese cloth, and centrifuged at 10,000 rpm for1 hour. The extraction process was repeated three times, and the filteredextracts centrifuged. After centrifugation, the supernatants were combined,and adjusted to 40 % saturation with ammonium sulphate. The extract was leftat 00 for 24 hours. After centrifugation at 10,000 rpm (1 hour), the supernatantwas brought to 60 % saturation with ammonium sulphate, and left standing at00 for several hours. The pellet was suspended in 0.1 M potassium phosphatebuffer (pH 5.8), and then dialyzed against 10 mM phosphate (pH 5.8) for 24hours (fraction 1). The 60% ammonium sulphate solution was centrifuged at10,000 rpm for 40 minutes, and the pellet collected. The pellet was suspendedin 0.1 M phosphate buffer (pH 5.8), and dialyzed against 10 mM phosphate (pH5.8) for 24 hours (fraction 2). Fractions 1 and 2 were assayed by the dimedonmethod (Neidleman and Geigert, 1986). Fraction 1 displayed little activity; fraction 2 was purified on a DEAE-Sephadex A 50 column by gradient elution with0.1 M to 0.2 M potassium phosphate buffer, pH 6.0. The column fractions wereassayed, and the active portions combined, and dialyzed against 10 mM phosphate buffer (pH 5.0) for 16 hours. The semi-purified enzyme preparation waslyophilized, and the crude chloroperoxidase (in 25 ml of 0.1 M potassium phosphate, pH 5.0) assayed once again. Two isolations and purifications of lichenchloroperoxidase were undertaken. The yield of crude chioroperoxidase per runwas 6 mg, with a specific activity of 0.67 units/mg of protein for each purification(for a combined total of 12 mg chioroperoxidase). The protein content and specific87activity were calculated according to the Bradford dye-binding method (Bradford,1976) and dimedon assay (Neidleman and Geigert, 1986), respectively. The semi-purified chloroperoxidase was used in the experimental protocols described in Figure 13.The UV-visible absorption spectrum of the chloroperoxidase shown in Appendix 6 shows the characteristic absorption band (280 nm) for protein, but the absence of absorption bands for a heme or flavin prosthetic group suggests that thelichen enzyme lacks such a prosthetic group. Several nonheme chloroperoxidases are known from fungi and bacteria (Franssen and van der PIas, 1992).4.3 RESULTS AND DISCUSSIONIn order to examine whether N. Iaevigatum chloroperoxidase was capableof chlorinating anthraquinones in vitro, emodin (1) and 7-chloroemodin (2) wereincubated with the chloroperoxidase preparation from the lichen. For comparison,commercial chioroperoxidase (from the fungus Caldariomyces fumago) was alsotested for its ability to chlorinate the anthraquinones.The results of the incubation experiments can be seen in Table 15. Incubation of emodin (1) with N. Iaevigatum chloroperoxidase (at pH 3.4) produced 7-chloroemodin (2), but not 5,7-dichloroemodin (3); similarly, incubation of 7-chloro-emodin with the lichen enzyme failed to give 5,7-dichloroemodin. In contrast, commercial fungal chloroperoxidase catalyzed the conversion of emodin to 7-chloro-emodin and 5,7-dichloroemodin; 7-chioroemodin was also converted to 5,7-dichloro-emodin. The latter reaction took place with extraordinary efficiency; after 40 hours88at ambient temperature, the 7-chloroemodin had been completely converted to5,7-dichloroemodin. The control reaction (no enzyme) also converted emodin to7-chloroemodin and 5-chloroemodin, as well as a small quantity of 5,7-dichioro-emodin; surprisingly, 7-chioroemodin was not further chlorinated in the controlreaction (Table 15).The time course of the reactions suggests that chlorination in the lichen andfungal enzyme reactions are, indeed, catalyzed by the chloroperoxidase, and arenot simply a consequence of hydrogen peroxide-mediated chlorination. 5-Chloro-emodin and 7-chioroemodin were detected, by TLC, at least 20 hours earlier thanin the control reaction (Table 15). This indicates that both enzymes increased thereaction rate for the formation of the product. 5,7-Dichloroemodin was formed, fromemodin, at the same rate for both the fungal enzyme and control reactions, but wassynthesized from 7-chioroemodin only in the fungal chioroperoxidase reaction.The results of the incubation experiments are consistent with the knownchemistry of Nephroma IaevIgatum. The chlorination of 7-chloroemodin to 5,7-di-chioroemodin by the lichen chloroperoxidase would not be expected, since neither5- nor 5,7-dichloroemodin is found in the lichen (Cohen and Towers, 1995a). While5,7-dichloroemodin is known from the lichen Heterodermia obscurata (Yosioka etal., 1968b; Cohen and Towers, 1995b), until now only a few 5-chloroanthraquinoneshave been isolated from fungi (Steglich et al., 1969; Ayer and Trifonov, 1994).Steglich and coworkers (1969) proposed a biogenetic scheme in which late chlorination of an anthraquinone in ring position 5 might take place in the genus Der89mocybe. However, no isotope incorporation experiments in Dermocybe with 36C1have been performed, nor has a chloroperoxidase been isolated from a basidiomycete.It is thus presently unclear why Nephroma Iaevigatum would produce the5-chioro-substituted emodins in the 13C feeding experiment, while they were not evident, by TLC, in either the sample selected for tracer studies, the other two incubation studies, or in all the previous work on the lichen. It is further unclear why thechloroperoxidase, once isolated from the lichen, appears incapable of effecting 5-chlorination. A typical electrophilic chlorination (Cl) reaction should give both the5 and 7-chloro-substituted products. This expectation was observed with the hydrogen peroxide-mediated chlorination of emodin (control reaction) and in the reaction catalyzed by the fungal enzyme. The 13C incubation mixture (which produced the 5-chloro isomers) was not supplemented with exogenous chloride ion,but the lichen chloroperoxidase medium contained 40 mmol KCI. Perhaps excesschloride ion inhibits the ability of this enzyme to effect 5-chlorination. It is also conceivable that 5-chlorination was effected by another enzyme, which was not removedfrom lichen tissue or was lost during purification. Finally, it is conceivable that the5-chioro isomers are present in the wild lichen, but were not previously detected dueto their low concentrations and Rf values so close to their 7-chloro counterparts.4.4 PHYSICAL AND CHEMICAL DATA FOR REACTION PRODUCTS4.4.1 PRODUCTS FROM CONTROL (NO ENZYME) REACTION907-Chloroemodin (2) was obtained as orange crystals from ethyl acetate:ElMS m/z[M] 306 (33), 304 (100). TLC (8:2 chloroform:methanol): R 0.50.5,7-Dichloroemodin (3) was obtained as red crystals from methanol: ElMSm/z[M] 384 (2), 382 (14), 380 (22), 342 (12), 340 (66), 338 (100). TLC: R, 0.20.5-Chloroemodin (4) was obtained as orange crystals from ethyl acetate:ElMS m/z [M} 306 (37), 304 (100); 1H NMR (DMSO-d6,400 MHz) ö 7.38 (s, 1 H,H-4), 7.03 (s, 1H, H-2), 6.60 (s, 1H, H-7), 2.48 (s, 3H, Me). TLC: R 0.60.4.4.2 PRODUCTS FROM COMMERCIAL CHLOROPEROXIDASE REACTION7-Chloroemodin (2) was obtained as orange crystals from ethyl acetate:ElMS m/z [M] 306 (33), 304 (100). TLC: R 0.505,7-Dichloroemodin (3) (from emodin) was obtained as red crystals frommethanol: ElMS m/z[M]342 (12), 340 (68), 338 (100). TLC: R 0.205,7-Dichloroemodin (3) (from 7-chloroemodin) was obtained as red crystals from methanol: ElMS m/z[M]’ 342 (13), 340 (68), 338 (100); 1H NMR (DMSOd6, 400 MHz) 7.40 (s, 1 H, H-4), 7.05 (s, 1 H, H-2), 2.40 (s, 3H, Me). TLC: R 0.20.5-Chloroemodin (4) was obtained as orange crystals from ethyl acetate:ElMS mlz 306 (37), 304 (100); ‘H NMR (DMSO-d6,400 MHz) ö 7.35 (s, 1 H, H4), 7.05 (s, 1H, H-2), 6.60 (s, 1H, H-7), 2.48 (s, 3H, Me). TLC: Rf 0.60.4.4.3 PRODUCTS FROM N. LAEVIGATUM CHLQROPEROXIDASE REACTION7-Chloroemodin (2) was obtained as orange crystals from ethyl acetate:ElMS mIz[M] 306 (33), 304 (100). TLC: R, 0.50.91OH 0 OHHO — CH30Emodin (1) 7-Chloroemodin (2)CH3ciHO5,7-Dichloroemodin (4)05-Chloroemodin (3)Figure 14. Substrates and products from chlorination experiments.92CHAPTER 5: ANTIVIRAL ACTIVITIES OF LICHEN QUINONOID PIGMENTS5.1 INTRODUCTIONHypericin, and a number of structurally related anthraquinones and bianthrones, have been shown to posses antiviral activities against several animalviruses with membranes, including HIV-1 (Meruelo et al., 1988; Schinazi et al.,1990; Tang et al., 1990; Kraus et al., 1990; Anderson et al., 1991; Sydiskis et al.,1991; Barnard et al., 1992). None of these earlier studies, however, examined therole of light in mediating virucidal activity. Hypericin is a known photosensitizer,and its virucidal activity has been shown to be dependent on light (Hudson et al.,1991; Carpenter and Kraus, 1991; Hudson et al., 1993; Lenard etal., 1993).This may also be true for anthraquinones and bianthrones. In addition, the anti-viral activity of hypericin has been shown to be affected by a variety of assayparameters (Anderson et al., 1991; Hudson et al., 1993). Thus, relative activitiesmust be compared under uniform and optimal conditions.The compounds used in this study (Figure 15) were isolated either from thelichens Nephroma Iaevigatum and Heterodermia obscurata, or were natural productsobtained previously. All of the compounds, with the exception of hypericin, areknown constituents of lichens.5.2 MATERIALS AND METHODSThe anti-HSV assays follow the protocols described previously (Marles et al.,93CH3R = R’ = H; Emodin (1)R = CI, R’ = H; 7-Chloroemodin (2)R = R’ = CI; 5,7-Dichloroemodin (5)R OH, R =COOH; Endocrocin (7)R R’ = H; Chrysophanol (8)R = R” = R’” H, R’ = CH3; 6-0-Methylemodin (2)R = CI, R’ = R”’ = H, R” = CH3; 7-Chloro-1 -0-methylemodin (4)R Cl, R’ = H, R” = CH3, R”’ = OH; 7-Chloro-1-O-methyl-co-hydroxyemodin (13)R = H; Flavoobscurin A (9)R = CI; Flavoobscurin B (10)R H; Hypericin (11)R = CI; 7,7’-Dichlorohypericin (12)R 0 0R0OH 0 OHSkyrin (6)OH 0 OHFigure 15. Structures of lichen compounds used in antiviral assays.0)Table16.MinimuminhibitoryconcentrationsoflichencomroundsaciainstHSV-1virus.1CompoundCompleteInactivationPartialInactivationEmodin(1)216-Q-Methylemodin(2)inactiveinactive7-Chloroemodin(3)217-Chloro-1-Q-methylemodin(4)20.55,7-Dichloroemodin(5)0.250.125Skyrin(6)inactiveinactiveEndocrocin(7)inactiveinactiveChrysophanol(8)inactiveinactiveFlavoobscurinA(9)inactiveinactiveFlavoobscurinB(10)inactiveinactiveHypericin(11)<0.06<<0.067,7’-Dichlorohypericin(12)<0.06<<0.067-Chloro-1-Q-methyl-o-hydroxyemodin(13)inactiveinactiveigImI.951992). Briefly, the standard reaction mixtures were prepared by adding a knownamount of HSV-1 (100 pfu or plaque-forming units) to serial two-fold dilutions ofthe test compound made up in Dulbecco MEM medium plus 0.1 % serum, previously shown to be the optimum conditions for the antiviral activity of hypericin(Hudson et al., 1994). The range of concentrations of the compounds was 2.0jig/mI down to 0.06 jig/mI. The reaction mixtures were irradiated at 9 Kjoules,supplied by fluorescent lamps, for 30 minutes. The irradiation was provided byseveral General Electric F20 T12/cw lamps which gave a measured incident energy of 1.5 mW/cm2(the emission was between 380 and 700 nm). Controls included reactions kept dark by covering the assay plates with aluminum foil, andvirus without compound. All reactions were carried out in duplicate, and everycompound was tested at least twice.The cultures were inspected daily microscopically for residual HSV-1 infectivity, as characteristic CPE (cytopathic effects), and for comparison with virus onlyand cells only controls. In the case of the untreated HSV-1, CPE involved the entireculture by day 3-4. At this time, cultures that still displayed no viral CPE were considered free of infectious virus (complete viral inactivation in the reactions).Compounds that displayed good anti-HSV activity in the end-point tests wereevaluated in more detail by use of plaque assays. Reaction mixtures were preparedwith several concentrations of the compounds and HSV-1 (1 o pfu/ml). Followingirradiation with light as before, and with appropriate controls, the mixtures were Se-cc0)Table17.InhibitionofHSV-1virusinilaciueassay.Compound1#ofPlagues2%ofPlagues3None(control)936100Emodin(1)784847-Chloroemodin(3)904977-Chloro-1-Q-methylemodin(4)10621135,7-Dichloroemodin(5)0<0.1Hypericin(11)0<0.17,7’-Dichlorohypericin(12)1<0.111.0igImI.2 pfu/mix102dilutionfactor.3 Relativetocontrol.r-.0)Table18.EffectoflightonHSV-1inhibitionatdifferentsubstrateconcentrations.1CompoundDark/Lightl.0p.gImI0.1jig/mI0.01jig/mI5,7-Dichloroemodin(5)Dark100--Light<0.0152100Hypericin(11)Dark34100100Light<0.010.33247,7’-Dichlorohypericin(12)Dark43100100Light0.17501001 Valuesreportedas%pfu(plaque-formingunits)remaining.98rially diluted 10to ion. The 102 and i0 dilutions were each inoculated in duplicateonto monolayers of Vero cells in culture dishes (60 mm diameter) to permit adsorption of remaining infectious virus to the cells (60 minutes at 37°C). The inoculawere then removed by aspiration and replaced by molten agarose overlays (at42°C), which consisted of 0.5 % final concentration of agarose, Dulbecco MEMand 5 % serum. When the overlays had set, the cultures were returned to theincubator until plaques could be visualized (4 days). In order to facilitate theenumeration of plaques, the cell monolayers were fixed in 10 % formalin in phosphate-buffered saline, and stained with 1 % crystal violet in water. Plaques appeared as discrete round “holes” in the uniform blue monolayer of intact cells.The number of plaques at each dilution was calculated as pfu/mI, and comparedto the corresponding number in the untreated virus control.5.3 RESULTS AND DISCUSSIONTable 16 presents the results of the survey of 12 lichen compounds andhypericin for their inhibitory activity against HSV-1 virus. Emodin (1), 7-chloro-emodin (3) and 7-chloro-1-O-methylemodin (4) completely inactivated HSV-1at a concentration of 2 pg/ml. Partial inactivation was also seen for compounds1 (1 tg/ml), 3 (1 rig/mI) and 4 (0.5 pg/ml). The most active anthraquinone was5,7-dichloroemodin (5). Complete inactivation of the virus was attained at a concentration of 0.25 igIml, and 0.125 .tg/ml gave partial inactivation. The remaining99five anthraquinones (2, 6, 7, 8, 13) were completely inactive, even at 5 pjg/mI.The bianthrones flavoobscurin A (9) and flavoobscurin B (10) were also inactiveat 5 jig/mI. Finally, hypericin (11) and 7,7’-dichlorohypericin (12) showed comparable inhibitory activity against HSV-1; complete inactivation took place at lessthan 0.06 jig/mI.Table 17 compares the antiviral activities of the six active compounds in aplaque assay. As was the case with the initial end-point CPE method, this assaywas conducted following irradiation with light. Of the six compounds tested, 5,7-dichloroemodin (5), hypericin (11) and 7,7’-dichlorohypericin (12) completely inhibited HSV-1 at 1.0 jig/mI. Thus, of the 11 anthraquinones and bianthrones tested, only 5,7-dichloroemodin exhibited light-mediated anti-HSV activity comparableto that of hypericin and 7,7’-dichlorohypericin.This result is illustrated in Table 18, where the effects of light and dark activities of the three compounds are compared. At a higher concentration (1.0jig/mi), both hypericin and 5,7-dichloroemodin were appreciably more active than7,7’-dichlorohypericin in the light. In the dark, however, 5,7-dichloroemodin wastotally inactive; the two hypericins exhibited comparable moderate anti-HSV activity.At an intermediate concentration (0.1 gIml), the activities of 5,7-dichioroemodinand 7,7’-dichlorohypericin in the light started to decrease. At this concentration,all three compounds were inactive in the dark. Finally, at the lowest. concentration(0.01 jig/mI), only hypericin showed any anti-HSV activity in the light.100There are several significant findings to be gleaned from this study. Thefirst observation is the higher inhibitory activity of 5,7-dichioroemodin, whencompared with 7-chloroemodin and emodin. There also appears to be nodifference in activity between the monochlorinated anthraquinones and thenonchiorinated parents. This finding suggests that, in the emodin series at least,anti-HSV activity occurs when both the C-5 and C-7 ring positions are substituted;the possibility that 5-chioroemodin might also be active cannot be excluded, however. Replacement of the 1-OH in emodin by a 1-OMe group (compound 4) resulted in an initial partial inactivation, but the plaque assay results suggest that theviral CPE were merely delayed, and that eventually the CPE reached 100 %.Thus, for 1,3 and 4, lower concentrations of the compound gave little or no decrease in the progress of CPE, while higher concentrations gave complete inactivation of the virus.Modifications of the basic emodin structure (Figure 15) by placing an OMegroup at C-6 (compound 2), a COCH group at C-2 (compound 7), removal of theOH group at C-6 (compound 8), and substitution of a CH2O for a methyl group(compound 13) all gave inactive compounds. Of the dimeric structures, only thehypericins (compounds 11 and 12) showed anti-HSV properties; skyrin (6) (bianthraquinone) and flavoobscurins A (9) and B (10) (bianthrones) were completelyinactive.In general, the results suggest that the phenanthroperylenequinone structureof hypericin and 7,7’-dichlorohypericin is more active than either the less condensed101bianthrone structure, or the monomeric/dimeric anthraquinones (Table 16). The presence of several chlorine atoms in flavoobscurin A and B did not appear to positivelyinfluence their activities against HSV; in fact, the structural conformations of the twocompounds may account for their lack of activity against the virus. In contrast, placing chlorines in ring positions 5 and 7 of emodin resulted in greatly enhanced virucidal activity for 5,7-dichloroemodin. In the case of 7,7’-dichlorohypericin, substitution of two chlorine atoms in the 7 and 7’ ring positions of hypericin slightly reducedthe antiviral activity of the halogenated analogue in the light. Meruelo et al. (1988)had shown that pseudohypericin (with a CH2O group in place of a CH3) was significantly less active than hypericin. Compound 13, which also has a CH2Q group inplace of a methyl group, was completely inactive.In conclusion, this study provides evidence for the anti-HSV activities of several lichen anthraquinones and a chlorinated hypericin derivative. In particular, hypericin, 7,7’-dichlorohypericin and 5,7-dichloroemodin demonstrated significantvirucidal properties in light at a concentration of 1.0 p.g/ml. Hypericin and 7,7’-di-chlorohypericin also displayed moderate anti-HSV activities in the dark at 1.0 jig/mI.Hypericin is known to be a photodynamic compound that mediates biological activities via singlet oxygen (Thomas and Pardini, 1992; Thomas et al., 1992; De Witteet al., 1993; Hudson et al., 1994). It is not known, however, if the dark reaction follows the same mechanistic course as the light-mediated process. In fact, there maybe several different light and dark processes operating concurrently, or subject to102varying reaction conditions. This would appear to be the case with 5,7-dichloro-emodin, which showed the same light-mediated virucidal activity as hypericin,but was totally inactive in the dark at a concentration of 1.0 pjg/mI.103CHAPTER 6: CONCLUSIONSThis thesis addresses several different aspects of the chemistry, biochemistry and medicinal properties of quinonoid constituents of the lichens NephromaIaevigatum and Heterodermia obscurata. I have undertaken the isolation andcharacterization of twelve anth raquinone, bianthrone and phenanth roperylenequinone pigments from the two lichens, examined in vitro and in vivo the biogenesisof several anthraquinones, and tested thirteen structurally diverse lichen quinonoid compounds for their potential antiviral activities.The first study dealt with the isolation and identification of pigments inN. Iaevigatum and H. obscurata. Four emodin and two hypericin derivatives wereisolated from N. Iaevigatum collected in British Columbia. This lichen was foundgrowing only in the littoral zones of islands between the mainland and VancouverIsland. The preferential lichen substratum was granitic rock along the shoreline,but occasionally the lichen could be found on the trunks and branches of Big-LeafMaple (Acer macrophyllum). TLC surveys of several samples collected from different geographical locations and substrata showed a uniformity in chemicalcomposition. Although minor constituents, such as 7,7’-dichlorohypericin, 2,2’,7,7’-tetrachlorohypericin and 7-chloro- 1-O-methyl-o-hydroxyemodin gave onlyfaint TLC spots, they were present in all lichen extracts examined. There did notappear to be, therefore, any chemical variation within the regions and differentlichen substrata examined. Two chemical races of N. Iaevigatum are known104(Table 2), but they differ only in their triterpene compositions. Occasionally, nonpigmented forms of the lichen have been collected in Madeira, Canary Islands andin Portugal (James and White, 1987).The only previous chemicalstudies of Nephroma Iaevigatum were those ofBendz et al. (1967) and Bohman (1968). Bohman (1968) identified five anthraquinones in the lichen growing in Sweden. Of the five compounds characterized byBohman, I was able to isolate three, which are apparently common to both theBritish Columbian and Swedish samples (emodin, 7-chloroemodin and 7-chioro-1-O-methylemodin). I did not, however, find either 7-chloro-6-O-methylemodin or7-chloro-1 ,6-di-Q-methylemodin in any of my samples. Furthermore, Bohman didnot indicate in her paper how much lichen material was used in the study. Thus,her failure to find 7-chloro- 1- O-methyl-o)-hydroxyemodin, 7,7’-dichlorohypericinand 2,2’,7,7’-tetrachlorohypericin may reflect her having worked with insufficientmaterial, or this may truly represent chemical variation between two differentsamples of the same species (Cohen and Towers, 1995a).Three anthraquinones, two bianthrones and 7,7’-dichlorohypericin were isolated from Heterodermia obscurata collected in Maryland. As with N. Iaevigatum,H. obscurata displayed chemical uniformity, according to TLC, for samples collectedfrom different locations in Cedarville State Forest. In addition, with the exceptionof 7,7’-dichlorohypericin, the compounds obtained from the lichen growing in Maryland were identical to those found in a Japanese sample studied by Yosioka et al.105(1968a, 1968b) (Cohen and Towers, 1995b). Again, 7,7’-dichlorohypericin maynot have been identified by the Japanese group as a result of a lack of sufficientlichen material, or it may be restricted to particular varieties of H. obscurata.Thus, a systematic chemical study of N. Iaevigatum and H. obscurata from a variety of different geographical and environmental conditions is necessary beforegeneralizations about the chemistry of these two lichens can be made.The presence of emodin anthrones in both N. Iaevigatum and H. obscuratawas suggested by the TLC of lichen extracts. Several pale yellow spots were evident in the thin-layer chromatographs of all extracts of both lichens. Although notdefinitive, the Rs of the spots correspond closely to purported emodin anthrones inN. Iaevigatum (Bohman, 1968) and H. obscurata (Yosioka et al., 1968c; Kurokawa,1973).My second objective was to examine the biogenesis of emodin and chlorinated emodin derivatives in N. Iaevigatum using radioactive and stable isotopes.Lichen anthraquinones are believed to be formed from acetate, in the same manner as fungal anthraquinones. The polyketide pathways leading to fungal anthraquinones have been well established (Franck, 1984; Gill, 1994, 1995). I have demonstrated, using sodium [2-14C]acetate and sodium [1-13C}acetate, the polyketideorigins of emodin, 7-chloroemodin and 7-chloro-1-O-methylemodin in N. Iaevigaturn. Figure 16 shows the percentage incorporations for the14C-labelled emodins,and the percentage ‘3C enrichments (over and above the 1.1 % natural abundance)1068 x14CH3O2Na0 O 0CO2Hif8 xCH313O2NaO%02HIf7-Chloroemodin (0.1 %)CH37-Chloro- 1-O-methylemodin (0.02 %)HO CH3Emodin (0.01 %)CH37-Chloro-1 -O-methylemodinFigure 16. The lichen polyketide pathway in Nephroma Iaeviga turn.107at each alternately-labelled carbon in 7-chloro-1-O-methylemodin. Absolute incorporation rates of 0.1 % or higher for polyketide biosynthesis in fungi wereconsidered significant by Franck (1984); similar values for incorporations of [1-14Clacetate and [2-14C]acetate into Rumex alpinus and Rhamnus frangula anthraquinones were also regarded as significant by Leistner (1971, 1973). Intheir study of the biosynthesis of the aliphatic acid, (+)-protolichesterinic acid,Bloomer et al. (1968, 1969) obtained incorporation rates of 0.01 -0.08 % of [1-14C]acetate into the lichen Cetraria islandica. Finally, Gill and Giménez obtained,in their study of the biosyntheses of anthraquinones in Dermocybe sanguinea,13C atom % enrichment levels ranging from 0.3 to 1.3 % from sodium [1-3C]ace-tate (1990b).The degree of labelling is highly subject to the experimental conditionsand target organism of the feeding study: higher plants, bacteria, marine algae,and some lower fungi tend to have higher metabolic turnover rates than mostbasidiomycetes and lichens. Not surprisingly, therefore, there have been fewfeeding studies with mushrooms and lichens. In general, based on previousstudies with plants and fungi, the degree of labelling of any isolated product willdepend on the amount of isotope supplied, the metabolic or reproductive stageof the organism at the time of precursor application, the number of potential biochemical pathways to which the labelled precursor may contribute, cellular compartmentalization of metabolites and the purity of the final labelled product (Luck-108ner, 1990). Several of the problems associated with whole-organism feedingstudies can be reduced by using cell cultures or enzyme preparations, which bypass complications associated with metabolite compartmentalization. In addition,use of purified enzymes and structurally-pertinent labelled precursors reducesthe likelihood of unwanted biochemical transformations, yet can result in thedemonstration of direct precursor-product relationships.This has been shown for several transformations of anthraquinones: 14C-labelled emodin .converted to radioactive geodin and dihydrogeodin in Aspergillusterreus (Fujimoto et al., 1975), conversion of3H-emodin to radioactive parietin ina cell-free preparation from Aspergilus parasiticus (Anderson, 1 986b), deoxygenation of emodin to chrysophanol by emodin deoxygenase from Pyrenochaetaterrestris (Ichinose et aL, 1993), and oxidation of emodin and chrysophanol an-thrones to the corresponding anthraquinones by emodinanthrone oxygenasefrom Aspergillus terreus (Chen et al., 1995).In particular, Sankawa and coworkers have established the enzymatic processes leading to the oxidation of emodin anthrone to emodin, and the 0-methyl-ation of emodin to 8-0-methylemodin, in Aspergillus terreus (Fujii et al., 1982,1991; Chen et al., 1992, 1995). Two enzymes have been isolated from the fungalculture: emodin anthrone oxygenase and emodin 0-methyltransferase. In a survey of ten anthraquinone-producing microorganisms, Fujii et al. (1991) found an-throne oxygenase activity in all of them; this confirms the ubiquity of the biosyn109thetic process of anthrone oxidation to anthraquinone in microorganisms. EmodinO-methyltransferase exhibited a broad substrate specificity with respect to methylation, when tested on sixteen anthraquinones and anthrones (Fujii et al., 1982;Chen et al., 1992). Emodin showed the highest relative activity (100 %), followedby 4-hydroxyemodin (80 %), -hydroxyemodin (22 %) and 7-chloroemodin (18 %).The remaining substrates showed little or no activity. The implications of these resuits for the formation of 7-chioro- 1- O-methylemodin and 7-chloro- 1-O-methyi-ohydroxyemodin in Nephroma Iaevigatum are: 1) it is reasonable to assume thatanthraquinone-producing lichens contain analogous enzymes catalyzing the formation of anthraquinones from anthrones; and 2) Q-methylation of emodin occurspreferentially, but not exclusively, before chlorination of the anthraquinone ring.Taking into account the slow growth and metabolism of most lichens, I anticipated that the in vivo absolute incorporation rates and enrichments of 14C and13C-labelled acetate, respectively, into N. Iaevigatum would be lower than might beexpected for analogous feeding studies with plants or fungi. Therefore, future experiments designed to ascertain the biosynthetic pathways leading to anthraquinones in lichens should include the use of cell-free preparations or purified enzymes,‘4C-labelled anthrones and anthraquinones as precursors, [1 ,2-3C]acetate(which would provide unequivocal evidence for the transformations of octaketideprecursors into anthraquinones), and in vivo time-course studies in order to examine the patterns and progressions of precursor metabolism in the “cultured” lichen.110Since there is tentative evidence for the presence of anthrones in lichens,and anthrones have been established as anthraquinone precursors in some fungi (Sankawa et al., 1973; Franck, 1984; Gill and Giménez, 1991; Gill et al., 1992;Gill, 1994, 1995) and higher plants (Labadie et al., 1972; Yagi et al., 1978; Vederasand Nakashima, 1980; Grün and Franz, 1980, 1981; Sigler and Rauwald, 1994;Chen et al., 1995), biosynthetic transformations of anthrones to anthraquinones inlichens should be studied using labelled precursors in both in vivo and in vitro systems.Finally, the origin(s) of the 5-chloro-substituted emodins, obtained from the[1-3C]acetate feeding experiment, should be examined in greater detail. Systematic chemical surveys of natural populations of Nephroma Iaevigatum (and other lichens) should be undertaken in order to determine if 5-chloroanthraquinones areubiquitous or occasional constituents of lichens; or if they represent anomalousnatural products produced by N. Iaevigatum only under laboratory conditions. Theproduction of the unexpected compounds may be a result of having perturbed themetabolism of the lichen by using a relatively large amount (1 gram) of 0.12 M sodium [1-3C]acetate, or a consequence of changes in the ionic strength or pH of thelichen’s environment during the course of the experiment (Holker et al., 1974).The third area of research involved the study of the chlorination of anthraquinonoid pigments in N. Iaevigatum. Incorporation of sodium36chloride into thelichen established that exogenous chloride could be used by the lichen to makechlorinated anthraquinones, although this experiment by itself cannot rule out111the possibility of nonenzymatic chlorination, as lichens may contain sufficientamounts of hydrogen peroxide and endogenous chloride to carry out chlorinations in situ.An attempt was made, therefore, to isolate a chloroperoxidase (chlorinating enzyme) from N. Iaevigatum. Ultimately, a semi-purified enzyme fractionwas obtained from the lichen by a series of purification steps. Using the Dimedonand Bradford assays, I determined the specific activity and protein content of thecrude chloroperoxidase, respectively.Chlorination experiments with the semi-purified lichen ch loroperoxidase,and a commercially available fungal chloroperoxidase, demonstrated substratespecificity for both enzymes. The lichen chloroperoxidase failed to catalyze theformation of 5,7-dichloroemodin from either emodin or 7-chloroemodin as anticipated, while 5,7-dichloroemodin was synthesized in excellent yield from 7-chloro-emodin by the commercial enzyme, and from emodin in the control (no enzyme)reaction. In addition, 5-chloroemodin was produced from emodin in both the commercial and control reactions. Interestingly, although 5-chloroemodin was foundto be produced by the lichen during the [1-3C]acetate feeding experiment, it hasnot been previously identified in N. Iaevigatum, nor was it produced in the lichenchloroperoxidase reaction. This supports the hypothesis that the lichen is not normally capable of biosynthesizing 5-chloroemodin from emodin, and the presenceof the 5-chloroemodin derivatives in the “cultured” lichen is a probable consequenceof having perturbed the lichen’s normal metabolism during the course of the [1-13C]acetate incorporation study.112Chloroperoxidases represent a fairly large class of halogenating enzymes,known primarily from fungi and bacteria (Neidleman and Geigert, 1986; Asplund,1992; Franssen and van der Plas, 1992). A majority of the chioroperoxidasescontain heme prosthetic groups, but several nonheme chioroperoxidases havebeen obtained from these sources (Neidleman and Geigert, 1986; Franssen andvan der Plas, 1992). In particular, a nonheme chloroperoxidase isolated fromPseudomonas pyrrocinia was shown to catalyze the chlorination of indole to 7-chloroindole, although the product was verified only by comparison of the HPLCretention time with authentic samples of chloroindoles (Wiesner et al., 1986). Thisenzyme would thus represent the first haloperoxidase for which a regioselectivehalogenation has been demonstrated, as all previous studies of haloperoxidasecatalyzed halogenation reactions have shown a complete absence of any regloselectivity or stereoselectivity (Morrison and Bayse, 1973; Ramakrishnan et al.,1983; Neidleman and Geigert, 1986; Franssen and van der Plas, 1992).The results of the biohalogenation reactions suggest that the N. Iaevigatumchioroperoxidase exhibits regioselectivity with respect to the substrate emodin.The enzyme catalyzed the chlorination of emodin to give only 7-chloroemodin.Although the exact mechanism for any chloroperoxidase-mediated chlorination hasyet to be fully established, the fact that the lichen haloperoxidase catalyzed theformation of only one of the two expected products of electrophilic chlorination ofemodin suggests that the enzymatic chlorination mechanism in the lichen may be113somewhat different than the mechanisms proposed for fungal and algal haloperoxidases.The final topic of my thesis dealt with the antiviral properties of lichen compounds. Several lichen anthraquinones, bianthrones and hypericin derivativeswere examined in end-point CPE (viral cytopathic effects) and plaque assays withHSV-1 (herpes simplex virus type 1). Emodin, 7-chloroemodin, 7-chloro-1-O-methylemodin, 5,7-dichioroemodin, hypericin and 7,7’-dichlorohypericin showedfair to good virucidal activity in the presence of light. In the plaque assay, hypericin, 7,7’-dichlorohypericin and 5,7-dichloroemodin completely inhibited thevirus at 1.0 jig/mI concentrations; only hypericin was active, however, at 0.01jig/mI. The remaining compounds were found to be inactive at a concentration of 5.0 jig/mI.The results of the HSV-1 assays demonstrate that substitution of twochlorine atoms on the emodin structure enhances the virucidal activity substantially. On the other hand, 7-chloroemodin showed the same activity as emodinin the assay. Substitution of chlorine atoms in the hypericin structure did not enhance its activity, and the two chlorinated bianthrones (flavoobscurin A and B)were completely inactive. In addition, Q-methylation at position 1 of 7-chloro-emodin did not effect the antiviral activity. An unexpected result was that 5,7-dichloroemodin exhibited virucidal activity (1.0 jig/mI) only under the influenceof light.114Anthraquinones, and a number of structurally related compounds, havebeen shown to possess antiviral and associated activities (Brownet al., 1980;Konoshima et al., 1989; Kraus et al., 1990; Schinazi et at., 1990; Andersen et al.,1991; Sydiskis et al., 1991; Barnard et al., 1992; Tagahara et at., 1992; Cohenet al., 1995). In addition, hypericin and related perylenequinones have demonstrated antiviral activities against a variety of viruses (Meruelo et al., 1988; Lavieet al., 1989; Kraus et al., 1990; Schinazi et at., 1990; Tang et at., 1990; Andersenet at., 1991; Carpenter and Kraus, 1991; Hudson et al., 1991; Lopez-Bazzocchiet at., 1991; Barnard et al., 1992; Hudson et al., 1993; Lenard et at., 1993; Hudson et al., 1994; Cohen et al., 1995). The role of light in mediating the virucidalactivity of hypericin has been shown by Hudson et at. (1991, 1993), Carpenter etal. (1991) and Lenard et al. (1993). Until now, however, light dependence hadnot been demonstrated for anthraquinones or anthrones. Thus, 5,7-dichloroemo-din represents the first anthraquinone requiring light for its antiviral activity (Cohenetal., 1995). While most of the reports involve the testing of plant, fungat or synthetic anthraquinones and hypericin derivatives against viruses, the results presented here represent the first reports of antiviral naturally occurring chlorinatedanthraquinones and hypericin derivatives.Finally, experiments planned for the future should include the testing of5-chlorinated anthraquinones, as well as of other halogenated and nonhalogenated structural analogues of emodin and hype ricin, in order to ascertain the struc115ture-activity relationships that determine the natural product’s activities againstdifferent viruses under the influence of a host of reaction parameters.116BIBLIOGRAPHYAberhart, D. J., K. H. Overton, and S. 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Approximate phytogeographical distribution of Heterodermia obscuratain North America.142APPENDIX 3OH 0 OH81 8a I 9aEmodin (anthraquinone)OH OH8J.8a JI.9a 117i(HO4 H36’ 1 Ob lb 3’HO -(1Oc( CH37’1O2’8’(10flf laOH 0 OHEmodin bianthrone (bianthrone)OH 0 OH81 8a J.. 9a Ii7926JJ9cJHO (8b(9b- CH3HO JiOb J.. 1 b J.3b CH1Oc7’ 10 2’8hb0,faOH o OHHypericin (phenanthroperylenequinone)Figure 19. Numbering systems in quinonoid natural products.APPENDIX 4 143Sample Calculations of 13C Carbon Chemical Shifts— c = 128.5 + z1 (all aromatic carbons)\ / X = 195.2+z (C-9andC-10)Substituent x z1 z2 z3 z4-H 0.0 0.0 0.0 0.0-CH3 9.2 0.7 -0.1 -3.0-Cl 6.3 0.4 1.4 -1.9-OH 26.9 -12.8 1.4 -7.4-OCH3 31.4 -14.4 1.0 -7.7-COphenyl 9.3 1.6 -0.3 3.7OH 0 OCH3HOCH35-Chloro-1 -Q-methylemodinCarbon Calculation Value Obs.1 128.5 + 31.4(OCH)-0.1 (CH3) -0.3 (C=O) + 3.7(0=0) 163.2 160.22 128.5 - 14.4(OCH - 0.3 (C=O) + 0.7 (CH3) 114.5 120.03 128.5 + 9.2(CH3)+ 1.0 (OCH3)+ 3.7 (C=O) 142.4 145.84 128.5 + 0.7 (CH3) - 7.7 (OCH3)+ 3.7 (C=O) - 0.3 (C=O) 124.9 119.85 128.5 - 0.3 (C=O) + 3.7 (C=O) - 7.4 (OH) - 12.8 (OH) + 6.3 (Cl) 118.0 118.66 128.5 + 26.9 (OH) + 3.7 (C=O) + 0.4 (Cl) + 1.4 (OH) 163.3 161.07 128.5 + 1.4 (CI)- 12.8 (OH) -12.8 (OH)+ 3.7(C=O) 108.0 112.68 128.5 + 26.9 (OH) + 1.4 (OH) - 1.9 (CI) - 0.3 (C=O) + 3.7 (C=O) 159.1 160.29 195.2 + 1.0 (OCH3)+ 1.4 (OH) + 3.7 (C=O) - 1.9 (Cl) 199.4 191.810 195.2 - 3.0 (CH3) -7.7 (OCH3)+ 3.7 (C=O) - 7.4 (OH) -7.4 (OH) +1.4 (CI) 175.2 183.64a 128.5-0.1 (CH3)+ 1.0 (OCH3)+ 1.6 (C=O) -0.3 (C=O) - 1.9 (Cl) 128.8 130.68a 128.5 + 1.6 (C=O) - 0.3 (C=O) - 12.8 (OH) + 1.4 (CI) - 7.4 (OH) -7.7 (OCH3) 103.5 112.69a 128.5 -14.4 (OCH3)- 3.0 (CH3) + 1.6 (C=O) - 7.4 (OH) - 0.3(C=O) 105.0 113.41 Oa 128.5 + 1.6 (C=O) - 0.3 (C=O) + 1.4 (OH) + 1.4 (OH) + 0.4 (CI) 133.0 134.8Figure 20. Calculations of 13C carbon chemical shifts using the method ofEwing (1979).144APPENDIX 5Calculations of Protein Content and Specific Activity:1. A standard curve is prepared from standard solutions of BSA (Bovine Serum Albumin), containing 10 to 100 ig protein per 0.1 ml buffer, using the Bradford dye-binding assay. The weight of the protein is plotted against corresponding absorbances.2. The protein content in a sample of the active semi-purified lichen chioroperoxidase fraction is determined from the standard curve.3. Calculation:a. 100 j.iI of semi-purified lichen chloroperoxidase gave absorbances of 0.184 and0.200.b. A595 nm (0.184) is equivalent to 29.2 tg protein and A595 nm (0.200) is equivalent to31.6 ig protein.c. Average weight of protein is (29.2 ig + 31.6 p.g)/2 30.4 jig.d. Protein concentration is 30.4 pgI1 00 p1 or 0.3 .tg/p.l or 0.3 mg/mI.e. Total protein content in semi-purified lichen chloroperoxidase:20 ml (total volume of fraction) x 0.3 mg/mI = 6 mg total protein.f. 10 j.tl semi-purified lichen chloroperoxidase converted 2 x 1 o moles MCD(monochlorodimedon) to DCD (dichlorodimedon) in 100 minutes (6000 seconds)in dimedon assay.2 x iO moles DCD formed/6.0 x iO seconds = 3.33 x 10.11 moles sec1; for 20 mltotal volume of fraction, this is 6.66 x 108 moles sec1 or 0.066 iimoles sec1.g. 1 Unit of enzyme catalyzes the formation of 1 jimole product per minute underdefined conditions. Therefore, (0.066 j.imoles secj x (60 seconds/minute) = 4pmoles mm1 or 4 units of lichen chloroperoxidaseh. Specific Activity = 4 units/6 mg protein = 0.67 units/mg protein.Figure 21. Calculations of protein content and specific activity of semi-purifiedchloroperoxidase from Nephroma Iaevigatum.145APPENDIX 6Figure 22. UV spectrum of semi-purified chioroperoxidase from Nephroma Iaevigatum(the spectrum was recorded in 0.1 M potassium phosphate buffer, pH 5.0,and a protein concentration of 0.3 mg/mI).eAPPENDIX 7146Figure 23. Lichen feeding experiments with stable and radiolsotopes.IAPPENDIX8Table19.HPLCdataforlichenanthraquinones,bianthronesandhypericinderivatives.1CompoundSourceRetentionTime2SolventandElutionSystemEmodinN.Iaevigatum4.581:1MeOH-2%AcOHinCH3 CN7-ChloroemodinN.Iaevigatum4.201:1MeOH-2%AcOHinCHCN7-ChioroemodinN.Iaevigatum4.231:1MeOH-2%AcOHinCHCNto100%MeOH(gradient;15mm)7-ChloroemodinH.obscurata4.1560:40MeOH-H2 0to100%MeOH(gradient;15mm)7-Chloro-1-O-methylemodinN.Iaevigatum4.241:1MeOH-2%AcOHinCH3 CN7-Chloro-1-Q-methyl-w-hy-N.Iaevigatum3.801:1MeOH-2%AcOHinCH1 CNdroxyemodin5,7-DichloroemodinH.obscurata2.9760:40MeOH-H2 0to100%MeOH(gradient;15mm)FlavoobscurinAH.obscurata3.761:1MeOH-2%AcOHinCHCNFlavoobscurinBH.obscurata3.531:1MeOH-2%AcOHinCHCNHypericinSigmaChem.Co.13.4460:40MeOH-H2 0to100%MeOH(gradient;10mm)7,7’-DichlorohypericinN.Iaevigatum13.8560:40MeOH-H2 0to100%MeOH(gradient;10mm)7,7’-DichlorohypericinSynthetic13.9360:40MeOH-H2 0to100%MeOH(gradient;10mm)2,2’,7,7’-TetrachlorohypericinN.Iaevigatum14.1860:40MeOH-H2 0to100%MeOH__________________________________________________________(gradient;10mm)1Allcompoundswereelutedfromareversed-phaseWatersBondapakC 18column(3.9x300mm).2Retentiontimesareinminutes.

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