Triterpenes epoxidations with

Various attempts to oxidize 7,8- and 8,9-unsaturated tetracyclic triterpenes such as (1) to the corresponding epoxides by reaction with perbenzoic acid afforded mixtures of epoxide with the 7,9(1 l)-diene. Fried et al. found that the difficulty is eliminated by use of m-chloroperbenzoic acid in chloroform, which gave exclusively... 

Double Sharpless epoxidation was also applied to the synthesis of a key intermediate 10 to the meso-compound 11 that is related to teurilene, abioactive polycyclic triterpene isolated from red alga Laurencia obtusa,4 The reaction of ( , )-Bisallylic alcohol 9 gave the bisglycidic alcohol 10 with 80% de and 89% ee for each epoxidation (Scheme 6AA.4). 

Van Tamelen has examined extensively the cyclization of monoepoxide-polyolefins (94). For example, he found (95) that the treatment of racemic epoxide 247 with SnCl in nitromethane provided a major product (35% yield) which was identified as the racemic tetracycle 248. This represents an over all, close simulation of the squalene tetracyclic triterpene bioconversion (except for optical activity). 

Limonoids are C2(, nortriterpenoids deriving from a C30 triterpene precursor. The best known limonoids are the Azadirachta indica (neem tree) antifeedant azadirachtin (C50L C40 C6 -C60(epoxide methylene cross-link) furan) and the Citrus species (Rutaceae) bitter antifeedant limonin (G50L G40 G6 G6 C50L(epoxide)-furan). Limonin gives a delayed bitter taste to Citrus fruit. The limonoids are typically bitter compounds with insect antifeedant activity... 

An impressive cationic domino polycyclization has been developed by Corey and coworkers in their short and efficient enantioselective total synthesis of aegicer-adienol (1-150), a naturally occurring pentacyclic nor-triterpene belonging to the 3-amyrin family [42]. Thus, the treatment of the enantiopure monocyclic epoxy tet-raene 1-147 with catalytic amounts of methylaluminum dichloride induces a ca-tion-JT-tricyclation by initial opening of the epoxide to form the tetracyclic ketone 1-148 in 52 % yield, and its C-14 epimer 1-149 in 23 % yield, after silylation and chromatographic separation (Scheme 1.37). Further transformations led to aegicer-adienol (1-150) and its epimer 1-151. 

Squalene oxidases enantiospecifically and regiospecifically epoxidize the terminal double bond of squalene 1 to give (31S )-squalene epoxide 258. This, in turn, is the precursor of triterpenes and sterols (e.g., 3) in both plants and animals. Squalene oxidases are found in higher forms of life with the probable exception of insects and terrestrial annelids. They are also present in algae and other lower forms with the possible exception of some bacteria. The enzyme system requires molecular oxygen, NADPH and FAD as well as supernatant protein and phospholipids. There is no evidence for the participation of cytochromes P-450. 

Several pentacyclic triterpenes found in ferns also appear to be formed from squalene (1), but not via the intermediacy of the 2,3-epoxide (2) (Fig. 23.4). Compounds, such as fer-nane (16), are found in several ferns (Goodwin, 1973). 24-Ethoxyhopane (17) has been reported from the fern Onoclea neriifolia. Although these triterpenes apparently are distributed widely in ferns, triterpenes derived from condensation of squalene 2,3-oxide frequently co-occur with them. 

A large number of triterpenes and all steroids in plants are derived by cyclization of squalene (as the 2,3-epoxide) via a chair-boat-chair-boat transition state (Fig. 23.5). The cyclic structures formed can all be rationalized in terms of the ways that the squalene 2,3-epoxide (2) may be folded (pseudo-chair-and-boat conformations) on the enzyme surface, with due consideration given to the stereoelectronic requirements for cyclization. Cyclization usually is initiated by acid-catalyzed opening of the ring of (35)-2,3-squalene... 

Seedlings of cucumber, Cucumis sativus, converted the triterpene (51) into cucurbitacin C (52) microsomal fractions from seedlings of Cucurbita maxima converted squalene 2,3-epoxide into 10a-cucurbita-5,24-dien-33-ol. Neither parkeol (53) (Fig. 23.1), cycloartenol (3), 11-ketocycloartenol, or 24,25-dihydro-9a,lla-epoxyparkeol were incorporated significantly, and no products with a cu-curbitane skeleton were isolated (Balliano et al., 1983 Harrison, 1985). 

In the oleanane-ursane group of triterpenes, postulated hydride shifts [as proposed by Eschenmoser and co-workers (1955)] have now been demonstrated. Incorporation of [4- C]MVA into oleanolic acid (56), ursolic acid (63), and several other metabolites was accomplished with tissue cultures of Isodon japonicus (Lamiaceae). Following introduction of [4- C]mevalonate and [l,2- C2]acetate, the pentacyclic triterpene compounds were examined by C-NMR spectroscopy. Rings D and E were formed by the routes predicted by Eschenmoser et al. (1955). The labeling pattern at carbons 4,23, and 24 of 3- p/-maslinic acid (64) indicates that this series is formed from (35)-squalene epoxide, rather than from the (3R)-epimer. 

Many metabolically altered triterpenes contain epoxides, lactones, furans, and cyclopentanoid systems, functional units that are associated with biological activity in other... 

For example, as shown in Scheme 6.35, in the presence of the appropriate catalyst(s) it is known (Sir John Cornforth, et al., vide supra) that the triterpene squalene (2,6,20,15,19,23-hexa-methyltetracosa-2,( )6,(F)10,( )14,( )18,22-hexaene [C30H50]) undergoes oxidation (forming an epoxide) and then via a zipper-like cascade, with one double bond adding to the next, cycUzation and rearrangement with migration of hydride and methide (or their equivalent) follow to produce lanosterol. 

Triterpenes are much rarer than sesquiterpenes or diterpenes, and can be divided into two distinct groups. The first group comprises squalene and its three possible epoxides which are also characterized by having all their asymmetric carbons with the configuration (S) 2,3(S), 6(S),7(S), and 10(S),ll(S)-epoxysqualene. These four substances were isolated from a single species, Caulerpa prol era (De Napoli et al., 1980, 1982). 

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