Chapter 21

Structural Diversity in Lignans and Neolignans

Adrian F. A. Wallis

CSIRO Forestry and Forest Products, Private Bag 10, South Clayton MDC, Victoria 3169, Australia

Lignans are a diverse group of optically active plant phenols formed by the union of two phenylpropane units through the center carbon atoms (C-8 /C-8') of their sidechains. Phenylpropane dimers which are linked through atomic centers other than the 8,8' carbons are frequently termed neolignans. The compounds can contain free phenolic hydroxyl groups or aromatic alkoxy groups. The range of most of the naturally occurring lignan and neolignan structures are outlined in this chapter, and methods for their structural elucidation are discussed. Some pitfalls in the determination of structures by N M R spectroscopy are given. Approaches to the biomimetic synthesis of lignans and neolignans are illustrated by the oxidative coupling reactions of isoeugenol and 4-propenylsyringol. In some cases the laboratory preparation of lignans by oxidative coupling has occurred in advance of their isolation from plant materials. The remarkable stereo-specificity noted in the oxidative coupling of p-hydroxyphenyl-propenes through the C-8/C-8' carbon atoms is rationalized in terms of mechanisms passing through favored intermediate complexes. However, this stereospecificity is not observed in every case, so that more study is required for a more complete understanding of the reaction mechanisms involved.

The term 'lignan' was given by Haworth in 1936 to a group of optically active phenylpropanoid dimeric extractives linked through the central (C-8/C-8')carbon atoms of their propane sidechains (1) (7). The family of compounds was extended in 1972 by Gottlieb to include 'neolignans', phenylpropanoid dimers linked through atoms other than the C-8/C-8' carbon atoms of the sidechains (2). However, Gottlieb later revised the definition to cover only those compounds formed by the oxidative dimerization of allyl or propenylphenols (3). The revised classification has not gained general acceptance, and the original definition of neolignans is more frequently used (4, 5). Related compounds occur in the form of lignans and neolignans such as sesquilignans, dilignans and trilignans with 3, 4 and 6 phenyl­propane units, respectively. Hybrid lignans are exemplified by flavonolignans, and

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norlignans have one or more carbon atoms from their propane sidechains missing (6). Examples of lignans with different skeletons include one acyclic structure (2), three monocyclic structures (3-5), two bicyclic structures (6 and 7) and two tricyclic structures (8 and 9). Neolignans have more varied skeletons, and examples include a 3,3'-linked biphenyl (10), 8,1'- and 8,3'-linked 1,2-diarylpropanes (11 and 12) and the 3-0-4'- and 8-0-4'-linked ethers (13 and 14).

Lignans were originally thought to be low molecular weight lignin fragments, although unlike lignin they are optically active, which implies enzymic control of the final steps in their biosynthesis. Both lignans and neolignans are understood to be formed by enzyme-catalyzed oxidative coupling of phenolic precursors, and the resulting dimers can subsequently undergo oxidation, reduction or alkylation to give the final product (see Chapters 22 and 25, this volume). They are widely distributed throughout the plant kingdom, occurring in up to 70 plant families in roots, rhizomes, wood, stems, leaves, fruits and seeds (5).

Lignans have been the subject of several reviews, notably in two books entirely devoted to them (5, 7), in chapters of monographs (8, 9), and several journal reviews (6, 10-18). They have become topical in recent times because increasing numbers of them have been described, from 14 in Haworth's 1936 review (7) to 440 in 1987 (5) (exclusive of neolignans), and because of their strong and varied biological activities. Notably, derivatives of podophyllotoxin (15) are used clinically in cancer chemotherapy (79), kadsurenone (16) is a potent antagonist of a mediator of inflammatory diseases (20), enterolactone (17) has digitalis-like activity (27) and masoprocol (18) is useful for treating various skin disorders (22).

Structural Elucidation of Lignans and Neolignans

Structure determination for the lignans and neolignans has followed similar lines to that for other classes of natural products. Various forms of chromatography have been used for the separation and tentative identification of unknown compounds—for instance capillary gas chromatography (GC) for separation of lignan glucosides (23), pyrolysis-GC (24) and high performance liquid chromatography (25) for lignans. Once separated, the components can be identified by spectroscopic comparison with authentic compounds, or by synthesis. GC itself has been used as a tool for the classification of lignans (26).

Ultraviolet spectrophotometry has also been a useful technique for establishing the class to which an unknown lignan belongs. A l l lignans show a basic U V absorption pattern typical of aromatic compounds, with three bands in the regions of 210, 230 and 280 nm (5). The infrared spectrum is also a useful tool in structure determination for lignans. However, lignans often occlude small guest molecules such as the solvent from which they are crystallized, that can distort the information obtained from their infrared spectra (27).

Mass spectrometry applied to lignans leads to extensive fragmentation and, although the spectra do not show many daughter ions of diagnostic significance, they do allow the characterization of aromatic rings and basic features of dimeric structures (5). Direct vapor injection is not an appropriate technique for obtaining mass spectra of lignan glycosides, because they are generally too involatile. For such compounds, chemical ionization (28), field desorption and fast atom bombardment (29) are important ancillary mass spectral techniques.

The most useful spectral technique for structural elucidation of lignans and neolignans is nuclear magnetic resonance (NMR) spectroscopy. The technique is particularly sensitive to structural variations (30), and there are substantial differences between the spectra of different classes of lignans. Both ! H and 1 3 C (37) NMR spectroscopy are useful and complementary methods; indeed the 1 3 C N M R spectroscopy of lignans has been the subject of extensive reviews (32, 33).

Although NMR spectroscopy has been a very powerful technique for elucidating the structures of lignans and neolignans, some errors in assignment have occurred through incorrect interpretation of the spectra. For instance for thomasic acid, a

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lignan isolated from the heartwood of the American rock elm Ulmus thomasii, a cis configuration was assigned to the C-7' and C-8' substituents in 19a on the basis of the small coupling constant J7,8, in its ! H NMR spectrum, indicative of a small dihedral angle between the C-H bonds (34). This was later revised to the trans configuration 19b, when it was recognized that the compound assumed a tran s-diaxial conformation (35). This conformation has subsequently been assigned to other dihydronaphthalene lignans (36, 37). A similar correction has been made to the assignment of a guaiacyl-dihydronaphthalene (38).

In spite of the current sophistication of NMR spectroscopic techniques, errors still occur in the assignment of configuration to neolignans on the basis of ] H and 1 3 C N M R spectra. Green et al. (39) originally proposed the unusual 8-0-3 ' structure for a lignan from the roots of Piper capense, with 1,3,5-substitution of the etherified ring, as a result of NMR spectral analysis. The structure was later confirmed to be that of the conventional 8-0-4' neolignan 20 (40). Corrections have been made to the assignment of configuration for two similar structures, the 8-0-4' linked isomers of the neolignan dehydrodiconiferyl alcohol (41) isolated from Arum italicum (42). The original errors of assignment were made on the basis of the deceptively simple pattern for the three aromatic proton signals (two singlets) in their 1 H NMR spectra, caused by coincident chemical shifts of the two protons at C-5' and C-6'.

N M R spectroscopy has been used to ascertain the conformation of lignans and neolignans in solution. The probable conformations of isomeric tetrahydrofuranoid lignans were obtained from their ! H (43) and 1 3 C (44) NMR spectra. Those of 8-0-4' neolignan isomers were also determined by lH (45, 46) and 1 3 C (47) N M R spectroscopic examination, and it was concluded that hydrogen bonds between the benzylic hydroxyls and adjacent ether oxygen atoms influenced the conformation. This was confirmed in the solid state by x-ray crystal structure determinations (48, 49).

Synthesis of Lignans and Neolignans

The biosynthesis of lignans and neolignans has received attention in various reviews (5, 7, 8), and is discussed in more detail in Chapters 22 and 25 of this volume. Both classical organic syntheses and biomimetic syntheses involving phenolic oxidative coupling have been employed as approaches to the synthesis of lignans and neolignans in the laboratory (5, 17, 50). These syntheses have value for structural confirmation of the compounds, and for possible pharmaceutical applications. The absolute configurations of lignans have been assigned through optical rotatory dispersion, circular dichroism and x-ray crystallography (5). Several enantioselective syntheses for lignans and neolignans have been outlined (51-53).

Biomimetic syntheses of lignans are illustrated in the lignans derived from the oxidative dimerization of isoeugenol and 4-propenylsyringol. Oxidation of iso-eugenol (21) with one-electron oxidants gives rise to a mesomeric free radical where the unpaired electron resides on the oxygen, C-5 and C-8 atoms, and the coupling of the various radical mesomers gives rise to a variety of products.

Erdtman (54) found that the arylcoumaran 22a (dehydrodiisoeugenol) was the product of the oxidative dimerization of isoeugenol in the presence of ferric chloride. This observation led him to propose that similar radical intermediates were involved in the biosynthesis of lignin. Formation of the arylcoumaran arises from radical coupling between the Ο and C-5 atomic centers to give an intermediate quinone-methide and subsequent cyclization leads to the arylcoumaran product. Forty years after Erdtman's paper describing the structure of dehydrodiisoeugenol, the compound was reported as a 8-3'-neolignan in the wood of Licaria aritu (55) and in nutmeg (Myristica fragrans) (56), and since that time it has been isolated from other plant materials also (57-61).

Oxidation of Ζ-isoeugenol (21b) with hydrogen peroxide catalyzed by a peroxidase enzyme gave, as one of the products, the arylcoumaran 22b with the sidechain retaining its Ζ stereochemistry (45, 62). However, the trans arrangement of

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R1 F*2 19 a H CH2OH

b CH2OH H

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substituents on the coumaran ring implies rotation about the C-7-C-8 bond of the quinonemethide intermediate prior to cyclization (62). There have been no reports of the isolation of naturally occurring neolignans with Z-propenyl sidechains, although arylcoumarans analogous to 22 with cis arrangements of the substituents on their coumaran rings have been proposed as structures for neolignans from the needles of Picea abies (63) and the inner bark of Betula pendula (64).

The products of the oxidation of ^-isoeugenol (21a) by hydrogen peroxide with peroxidase included a mixture of the threo and erythro isomers of the 8-0-4' neolignans 23a and 24a (45). The Z-isomer of isoeugenol 21b treated similarly gave rise to neolignans 23b and 24b with the Z-propenyl sidechains retained (45). These products arise from radical coupling between the Ο and C-8 atomic centers followed by addition of water to the resulting quinonemethide. Both threo and erythro neolignans 23a and 24a have been identified in the bark and root of Machilus thunbergii (60) and in mace (from Myristica fragrans) (65), while the erythro isomer has been found in the roots of Nardostachys jatamansi (66). The methyl ethers of threo 23a (66, 67) and erythro 24a (39, 40, 66) have also been found in plant extracts.

Both E- and Ζ-isoeugenol gave on oxidation by hydrogen peroxide with peroxidase, the tetrahydrofuranoid lignans 25a and 26a. These lignans are derived from stereospecific threo radical coupling between the two C-8 atomic centers to give a bisquinonemethide ; water adds to one quinonemethide to give a benzyl alcohol, which subsequently attacks the second quinonemethide with the formation of the tetrahydrofuran ring. The stereospecific formation of the threo bisquinonemethide has been rationalized in terms of an intermediate complex involving overlapping aromatic rings. The complex 27 which leads to threo coupling is more sterically favored than that (28) which would give erythro coupling. For 8-8' coupling of radicals from a phenol with bulky tert-buty\ substituents, both threo and erythro coupling products have been found (68); it was postulated that here the bulky substituents hindered the formation of the intermediate complex.

Both tetrahydrofuranoid lignans 25a and 26a have been isolated from mace (from Myristica fragrans) (69), and compound 26a has also been described in the extracts of Jatropha grossidentata (70). The dimethyl ethers of 25 and 26 are known as galbelgin (25b) and veraguensin (26b), and were first isolated from Himantandra belgraveana (71) and Ocotea veraguensis (72), respectively.

Reaction of £-4-propenylsyringol (29a) with ferric chloride in aqueous acetone has afforded the 8-0-4'-coupled threo and erythro dehydrodimers 30a and 31a, respectively, as the major products (46). Similar reaction of Z-4-propenylsyringol (29b) with ferric chloride has yielded the corresponding products with Z-propenyl sidechains, 30b and 31b. The yields of the compounds were improved when silver oxide was used as the oxidant (73). The erythro isomer 31a has been isolated from the fruit of Virola carinata ( 74), whereas the threo isomer 30a has been obtained as its aryl methyl ether from Piper polysyphorum ( 75).

Whereas the 8-0-4' coupled products predominated in the oxidation mixture fromZs-4-propenylsyringol (29a) in the acidic ferric chloride medium, reaction of 29a with hydrogen peroxide in the presence of peroxidase yielded a mixture of the threo-coupled tetrahydrofurans 32a and 33a as the sole products (45). However, reaction of Z-4-propenylsyringol with hydrogen peroxide in the presence of peroxidase gave the four tetrahydrofurans 32a-35a, resulting from both threo and erythro coupling (45). Three of the isomers 32a-34a have been found as components of mace (from Myristica fragrans) (69), while 32a has been isolated from the fruit of Licaria aurea (76) and the leaves of Virolapavonis (77). The dimethyl ethers, 32b (76, 78) and 34b (79), have also been found in plant materials.

The absence of stereospecificity in coupling between the two C-8 atomic centers of radicals derived from Z-4-propenylsyringol is surprising in view of the stereo-specificity in 8-8' coupling between the radicals from isoeugenol and £-4-propenyl-syringol. Stereospecific threo coupling has been noted in other oxidative coupling reactions of 1-arylpropenes (80-87). More study is required for a clearer understanding of the factors influencing the stereochemistry of oxidative coupling

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a Ar = 4-hydroxy-3,5-dimethoxyphenyl

b Ar = 3,4,5-trimethoxyphenyl

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processes leading to the formation of lignans, although a recent study showed some eryi/iro-coupled product from the oxidation of methyl sinapate (88). The number and type of structures found in this remarkable group of natural products will undoubtedly increase apace, particularly because of the intense interest in their biological properties.

Literature Cited

1. Haworth, R. D. Ann. Rep. Progr. Chem. 1936, 33, 266-279. 2. Gottlieb, O. R. Phytochemistry 1972, 11, 1537-1570. 3. Gottlieb, O. R. Fortschr. Org. Chem. Naturst. 1978, 35, 1-72. 4. Moss, G. P. Lignan and Neolignan Nomenclature; IUPAC-IUB Joint

Commission on Biochemical Nomenclature, 1987, 25.3, 1. 5. Ayres, D. C.; Loike, J. D. Lignans—Chemical, Biological and Clinical

Properties; Cambridge University Press: Cambridge, 1990. 6. Ward, R. S. Nat. Prod. Rep. 1995, 12, 183-205. 7. Rao, C. B. S. Chemistry of Lignans; Andhra University Press: Waltair, India,

1978. 8. Davin, L . B.; Lewis, N . G. In Phenolic Metabolism in Plants; Stafford, Η. Α.,

Ibrahim, R. K., Eds.; Plenum Press: New York, NY, 1992; pp 325-375. 9. Gottlieb, O. R.; Yoshida, M. In Natural Products of Woody Plants, I; Rowe, J.

W., Ed.; Springer-Verlag: Berlin, 1989, pp 439-511. 10. Castro, Μ. Α.; Gordaliza, M.; Del Corral, J. M. M.; San Feliciano, A .

Phytochemistry 1996, 41, 995-1011. 11. Ward, R. S. Nat. Prod. Rep. 1993, 10, 1-28. 12. Jensen. S.; Hansen, J.; Boll, P. M. Phytochemistry 1993, 33, 523-530. 13. Whiting, D. A. Nat. Prod. Rep. 1990, 7, 349-364. 14. Massanet, G. M.; Pando, E.; Rodriguez-Luis, F.; Zubia, E. Fitoterapia 1989, 60,

3-35. 15. Whiting, D. A. Nat. Prod. Rep. 1987, 4, 499-525. 16. Whiting, D. A. Nat. Prod. Rep. 1985, 2, 191-211. 17. Pelter, A. Recent Advances in Phytochemistry 1986, 20, 201-241. 18. Chatterjee, Α.; Banerji, Α.; Banerji, J.; Pal, S. C.; and Bhosal, T. Proc. Indian

Acad. Sci. (Chem. Sci.) 1984, 93, 1031-1057. 19. Anon. Chem. Brit. 1981, 17, 457. 20. Chang, M. N.; Han, G.-Q.; Arison, Β. H.; Springer, J. P.; Hwang, S.-B.; Shen, T.

Y. Phytochemistry 1985, 24, 2079-2082. 21. Braquet, P.; Senn, N . ; Robin, J.-P.; Esanu, Α.; Garay, R. P. J. Hypertension,

1986, 4 (suppl. 5), S461-S464. 22. Olsen, Ε. Α.; Abernethy, M. L . ; Kulp-Shorten, C.; Callen, J. P.; Glazer, S. D. ;

Huntley, Α.; McCray, M . ; Monroe, A. B.; Tschen, E.; Wolf, J. E. J. Am. Acad. Dermatol. 1991, 24, 738.

23. Kraus, R.; Spiteller, G. Liebigs Ann. Chem. 1990, 1205-1213. 24. Galletti, G. C.; Ward, R. S.; Pelter, A. J. Anal. Appl. Pyrol. 1991, 21, 281-293. 25. Betto, P.; Gabriele, R.; Galeffi, C. J. Chromatogr. 1992, 594, 131-135. 26. Ayres, D. C.; Chater, R. B. Tetrahedron 1969, 25, 4093-4098. 27. Hartwell, J. L.; Schrecker, A. W. In Progr. Chem. Org. Nat. Prod.; Zechmeister,

L., Ed.; Springer: Vienna, 1958; Vol. 15, pp 83-166. 28. Milne, G. W. Α.; Fales, H. M.; Axelrod, T. Anal. Chem. 1971, 43, 1815-1820. 29. Chapman, J. R. Practical Organic Mass Spectrometry—A Guide for Chemical

and Biochemical Analysis; 2nd Edition; Wiley: Chichester, 1993. 30. Günther, H. NMR Spectroscopy—Basic Principles, Concepts, and Applications

in Chemistry; Wiley: New York, NY, 1994. 31. Breitmaier, E.; Voelter, W. Carbon-13 NMR Spectroscopy; 3rd edition; V C H

Verlag: Weinheim, 1987. 32. Agarwal, P. K.; Thakur, R. S. Magn. Reson. Chem. 1985, 23, 389-418.

Dow

nloa

ded

by U

NIV

MA

SSA

CH

USE

TT

S A

MH

ER

ST o

n Se

ptem

ber

26, 2

012

| http

://pu

bs.a

cs.o

rg

Pub

licat

ion

Dat

e: A

ugus

t 13,

199

8 | d

oi: 1

0.10

21/b

k-19

98-0

697.

ch02

1

In Lignin and Lignan Biosynthesis; Lewis, N., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1998.

332

33. Agarwal, P. Κ.; Pathak, A. K. Magn. Reson. Chem. 1994, 32, 753-773. 34. Seikel, M . K.; Hostettler, F. D.; Johnson, D. B. Tetrahedron 1968, 24, 1475-

1488. 35. Wallis, A. F. A. Tetrahedron Lett. 1968, 5287-5288. 36. Ayres, D. C.; Harris, J. A. Chem. Commun. 1969, 1135-1136. 37. Agata, I.; Hatano, T.; Nishibe, S.; Okuda, T. Phytochemistry 1989, 28, 2447-

2450. 38. Wallis, A. F. A. Chem. Ind. 1974, 259. 39. Green, T. P.; Galinis, D. L . ; Wiemer, D. F. Phytochemistry 1991, 30, 1649-1652. 40. Reilly, M . J.; Wallis, A. F. Α.; Lundquist, K.; Stomberg, R. J. Wood Chem.

Technol. 1992, 12, 471-483. 41. Gellerstedt, G.; Lundquist, K.; Wallis, A. F. Α.; Zhang, L. Phytochemistry 1995,

40, 263-265. 42. Greca, M . D.; Molinaro, Α.; Monaco, P.; Previtera, L. Phytochemistry 1994, 35,

777-779. 43. Sarkanen, Κ. V.; Wallis, A. F. A. J. Heterocyclic Chem. 1973, 10, 1025-1027. 44. Fonseca, S. F.; Barata, L. E. S.; Rúvera, Ε. Α.; Baker, P. M. Can. J. Chem. 1979,

57, 441-443. 45. Sarkanen, Κ. V.; Wallis, A. F. A. J. Chem. Soc. Perkin Trans. I 1973, 1869-

1878. 46. Wallis, A. F. A. Aust. J. Chem. 1973, 1571-1576. 47. Braga, A. C. H. ; Zacchino, S.; Badano, H . ; Sierra, M . G.; Rúveda, E. A .

Phytochemistry 1984, 23, 2025-2028. 48. Wallis, A. F. Α.; Lundquist, K.; Stomberg, R. Acta Chem. Scand. 1991, 45, 508-

516. 49. Stomberg, R.; Lundquist, K . ; Wallis, A. F. A. J. Crystallogr. Spectrosc. Res.

1982, 23, 319-333. 50. Ward, R. S. Chem. Soc. Rev. 1982, 11, 75-125. 51. Zacchino, S. Α.; Badano, H. J. Nat. Prod. 1988, 51, 1261-1265. 52. Zacchino, S. Α.; Badano, H. J. Nat. Prod. 1991, 54, 155-160. 53. Zacchino, S. A. J. Nat. Prod. 1994, 57, 446-451. 54. Erdtman, H. Liebigs Ann. Chem. 1933, 503, 283-294. 55. Aiba, C. J.; Corrêa, R. G. C.; Gottlieb, O. R. Phytochemistry 1973, 12, 1163-

1164. 56. Forrest, T. P.; Forrest, J. E.; Heacock, R. A. Naturwissenschaften 1973, 60, 257-

258. 57. Forrest, J. E. ; Heacock, R. Α.; Forrest, T. P. J. Chem. Soc. Perkin Trans. I 1974,

205-209. 58. Hattori, M.; Hada, S.; Watahiki, Α.; Ihara, H. ; Shu, Y.-Z. ; Kakiuchi, N.; Mizuno,

T.; Namba, T. Chem. Pharm. Bull. Japan 1986, 34, 3885-3893. 59. Achenbach, H.; Groβ, J.; Dominguez, Χ. Α.; Cano, G.; Star, J. V . ; Brussolo, L.

C.; Muñoz, G.; Salgado, F.; López, L. Phytochemistry 1987, 26, 1159-1166. 60. Shimomura, H. ; Sashida, Y.; Oohara, M . Phytochemistry 1987, 26, 1513-1515. 61. Yoshida, M. ACS Symp. Ser. 1995, 588, 85-92. 62. Sarkanen, Κ. V.; Wallis, A. F. A. Chem. Commun. 1969, 298-299. 63. Lundgren, L. N.; Popoff, T.; Theander, O. Phytochemistry, 1981, 20, 1967-1969. 64. Smite, E.; Pan, H . ; Lundgren, L. N . Phytochemistry, 1995, 40, 341-343. 65. Hada, S.; Hattori, M.; Tezuka, Y. ; Kikuchi, T.; Namba, T. Phytochemistry, 1988,

27, 563-568. 66. Bagchi, Α.; Oshima, Y.; Hikino, H. Planta Med. 1991, 57, 96-97. 67. Barata, L . E. S.; Baker, P. M . ; Gottlieb, O. R.; Rùveda, E. A . Phytochemistry

1978, 17, 783-786. 68. Sarkanen, Κ. V.; Wallis, A. F. A. J. Chem. Soc. Perkin Trans. I 1973, 1878-

1881. 69. Hattori, M.; Hada, S.; Kawata, Y.; Tezeka, Y. ; Kikuchi, T.; Namba, T. Chem.

Pharm. Bull. Japan 1987, 35, 3315-3322.

Dow

nloa

ded

by U

NIV

MA

SSA

CH

USE

TT

S A

MH

ER

ST o

n Se

ptem

ber

26, 2

012

| http

://pu

bs.a

cs.o

rg

Pub

licat

ion

Dat

e: A

ugus

t 13,

199

8 | d

oi: 1

0.10

21/b

k-19

98-0

697.

ch02

1

In Lignin and Lignan Biosynthesis; Lewis, N., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1998.

333

70 Schmeda-Hirschmann, G.; Tsichritzis, F.; Jakupovic, J. Phytochemistry 1992, 31, 1731-1735.

71. Birch, A. J.; Milligan, B.; Smith, E. ; Speake, R. N. J. Chem. Soc. 1958, 4471-4476.

72. Crossley, N . S.; Djerassi, C. J. Chem. Soc. 1962, 1459-1462. 73. Zanarotti, A. J. Chem. Res. (M) 1983, 2625-2637. 74. Cavalcante, S. H.; Yoshida, M.; Gottlieb, O. R. Phytochemistry 1985, 24, 1051-

1055. 75. Ma, Y. ; Han, G. Q.; Li, C. L . ; Cheng, J. R.; Arison, B. H.; Hwang, S. B. Yaoxue

Xuebao 1991, 26, 345-350; Chem. Abstr. 1991, 115, 247446. 76. Bárbosa-Filho, J. M.; da Silva, M . S.; Yoshida, M.; Gottlieb, O. R.

Phytochemistry 1989, 28, 2209-2211. 77. Ferri, P. H . ; Barata, L. E. S. Phytochemistry 1991, 30, 4204-4205. 78. Holloway, D.; Scheinmann, F. Phytochemistry 1974, 13, 1233-1236. 79. Conserva, L. M . ; da Silva, M . S.; Filho, R. B. Phytochemistry 1990, 29, 257-260. 80. Erdtman, H. Svensk Kem. Tidskr. 1935, 47, 223-758. 81. Cartwright, N . J.; Haworth, R. D. J. Chem. Soc. 1944, 535-537. 82. Freudenberg, K.; Rasenak, D. Chem. Ber. 1953, 86, 755-758. 83. Freudenberg, K.; Schraube, H. Chem. Ber. 1955, 88, 16-23. 84. Freudenberg, K.; Harkin, J. M.; Reichert, M . ; Fukuzumi, T. Chem. Ber. 1958, 91,

581-590. 85. Freudenberg, K.; Lehmann, B. Chem. Ber. 1960, 93, 1354-1366. 86. Chapman, O. L.; Engel, M . R.; Springer, J. P.; Clardy, J. C. J. Am. Chem. Soc.

1971, 93, 6696-6698. 87. Wallis, A. F. A. Aust. J. Chem. 1973, 26, 1571-1576. 88. Setälä, H . ; Pajunen, Α.; Kilpeläinen, I.; Brunow, G. J. Chem. Soc. Perkin Trans.

I 1994, 1163-1165.

Dow

nloa

ded

by U

NIV

MA

SSA

CH

USE

TT

S A

MH

ER

ST o

n Se

ptem

ber

26, 2

012

| http

://pu

bs.a

cs.o

rg

Pub

licat

ion

Dat

e: A

ugus

t 13,

199

8 | d

oi: 1

0.10

21/b

k-19

98-0

697.

ch02

1

In Lignin and Lignan Biosynthesis; Lewis, N., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1998.