insect pheromones synthesized by oxidative

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Mar 16, 2005 - α,ω-difunctionalized 52. ... Epoxidation of alkene 57, which was produced by extension of the C chain of starting (R)-30by a Schlosser— ...
Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

INSECT PHEROMONES SYNTHESIZED BY OXIDATIVE TRANSFORMATIONS OF NATURAL MONOTERPENOIDS

G. Yu. Ishmuratov,1 R. Ya. Kharisov,1 R. R. Gazetdinov,1 and G. A. Tolstikov2

UDC 547.3+632.936.2

The literature on oxidative transformations of natural monoterpenoids to synthesize insect pheromones was reviewed. Key words: monoterpenoids, oxidation, insect pheromones. The literature on pheromone chemistry is broad and reflects the consistent interest in this area over the last 20 years in the form of monographs [1-7] and reviews [8-15]. Insect pheromones known today are rather simple molecules (less than four asymmetric centers and four functional groups). Therefore, the "ideal" substrate (chiral or achiral) for most such structures is a moderately functionalized molecule, in particular, hydroxy- and amino-acids in addition to monoterpenoids. The last class of compounds is the most accessible for this series and is especially convenient for the synthesis of molecules with a branched C skeleton, primarily isoprenoid pheromones. Oxidative methods of transforming monoterpenoids are the most convenient and widely used methods for carrying out various transformations of starting molecules and introducing most known functional groups. Considering this aspect, articles on the synthesis of insect pheromones that for one reason or another did not appear in previous reviews are reviewed herein.

ALLYLIC OXIDATION BY SELENIUM DIOXIDE One of the most common methods for functionalizing the C skeleton of unsaturated monoterpenoids containing an isopropylidene group is regio- and stereoselective oxidation at the allylic position, for example, by SeO2, of geraniol (1) and its derivatives or similar compounds. This method enables introduction into molecules of hydroxy- or oxo- functions that can be used in further transformations. For example, treatment of geranylacetate (2) with a stoichiometric or catalytic (with t-BuOOH) amount of SeO2 followed by hydride reduction gives unsaturated hydroxyacetate 3, which is widely used to synthesize insect pheromones [16-19]. In particular, 3 was oxidized to aldehyde 4, which was then converted to the olefin by n-propylidenephosphorane [16], to prepare racemic 5,9-dimethylheptadecane (7), the sex pheromone of the pear leaf blister moth (Leucoptera scitella Zeller). Organocuprate coupling of the resulting allylic acetate 5 with n-hexylmagnesium bromide led to 5,9-dimethyl-3,5,9heptadecatriene (6), exhaustive hydrogenation of which gave the desired pheromone 7. Condensation of bromoacetate 8 wth sodium malonic ester was used [17] to synthesize 10-hydroxy-4,8-dimethyl-4E,8Edecadienoic acid (11), an acyclic precursor of ferrulactone I (12), which is the principal component of the aggregation pheromone of the rusty grain beetle (Cryptolestes ferrugineus Stephen). Decarboxylation of the resulting triester 9 and subsequent saponification gave the key hydroxyacid 11, lactonization of which by bis-(4-t-butyl-N-isopropylimidazol-2yl)disulfide gave the desired 12. 1) Institute of Organic Chemistry, Ufa Scientific Center, Russian Academy of Sciences, 450054, Ufa, pr. Oktyabrya, 71, fax (3472) 35 60 66, e-mail: [email protected]; 2) N. N. Vorozhtsov Novosibirsk Institute of Organic Chemistry, Siberian Division, Russian Academy of Sciences, 630090, Novosibirsk, pr. Akad. Lavrent′eva, 9. Translated from Khimiya Prirodnykh Soedinenii, No. 6, pp. 509-522, November-December, 2005. Original article submitted December 15, 2004. 0009-3130/05/4106-0617 ©2005 Springer Science+Business Media, Inc.

617

d

O

63.4%

5

4

1, 2

90%

c 87.8%

CH3(CH2)3

30%

b

C6H13

6 f

1: R = H 2: R = Ac

a

e

55.6%

OR

g

HO

95%

3

EtO2C

h

Br

j

EtO2C

76%

8

9 O

k

i EtO2C

HO2C

42% from 8

OH

10

(CH2) 7CH3 7

O

42%

11

12

a. Ac2O, Py; b. SeO2, t-BuOOH; c. PCC; d. C3H7P+Ph3Br-, t-BuOK; e. n-C6H13MgBr, CuI; f. H2, Pd-C; g. CBr4, PPh3; h. NaCH(CO2Et)2; j. NaCl, DMF, H2O, 160°C; i. NaOH, dihlim; k. BID, PPh3

Odinokov et al. [18] changed the condensation and decarboxylation conditions in the synthesis of 11 from 3. In the former instance, they used chloride 13 instead of bromide 8 and carried out the reaction in the presence of Pd(OAc)2–PPh3. This increased the yield of 9 to 90%. They used LiCl in aqueous DMF for the decarboxylation of triester 9 whereas NaCl was used previously [17]. They isolated from the reaction mixture the expected 10 in addition to its acetoxy hydrogenolysis products as a mixture (7:3) of esters of regioisomeric ∆8,9-14 and ∆9,10-15 4,8-dimethyldecandienoic acids. Alkaline hydrolysis of 10 isolated by column chromatography gave the key hydroxyacid 11.

3

a

b

Cl

OAc

85%

92%

9

c 58% for10

+

10 + EtO2C 14 f

O

15

d, e HO

17

16

a. COCl2, DMF, Py; b. NaCH(CO2Et)2, Pd(OAc)2-PPh3, THF; c. DMF, LiCl, H2O, 173°C; d. H2, Pd-C; e. DIBAH; f. PCC (91%)

The resulting mixture of 14 and 15 was used to synthesize 4,8-dimethyldecanal (17), a racemic analog of the red flour beetle (Tribolium castaneum and T. confusum) aggregation pheromone, for which they were reduced to saturated alcohol 16, oxidation of which gave the desired aldehyde 17. Use of hydroxyacetate 3 is very convenient for preparation of 1,5-dimethyl-branched pheromones [19]. a

HO

OAc 3

89% e-g

OAc 20

62.6%

OAc

CH3(CH2)5

97%

18

d 89%

THPO

b, c

AcO

h, i

AcO

C12H25 21

(CH2)6CH3 19

74%

CH3(CH2)13

(CH2)13CH3 22

a. Ac2O, Py; b. n-C5H11MgBr, CuI; c. H2, Pd-C; d. DHP, TsOH; e. n-C12H25MgBr, CuI; f. TsOH, MeOH; g. Ac2O, Py; h. n-C13H27MgBr, CuI; i. H2, Pd-C

The C skeleton of 3 was extended by a one-step dialkylation of diacetate 18 with n-pentylmagnesium bromide in the synthesis of 7,11-dimethyloctadecane (19), a pheromone of the yellow fever mosquito (Aedes aegypti).

618

The same hydroxyacetate (3) was used by these same researchers to approach 15,19-dimethyltriacontane (22), a pheromone of the stable fly (Stomoxys calcitrans), for which a selective two-step lengthening of the chain of diether 20 was achieved. Allylic oxidation of 2 by SeO2 gives 3 and a side product of acetoxyaldehyde 4, which was used in a three-step synthesis of (S)-3,7-dimethyl-2E-octen-1,8-diol (23), a secretion of the plain tiger butterfly (Danaus chrysippus) [20]. Reduction of 4 by Baker's yeast, which transformed only the (S)-isomer, and subsequent alkaline hydrolysis gave diol 23 in 97% optical purity. a OAc

b, c

O

OAc

12%

2

4

HO

OH

20%

23

a. SeO2; b. BY, H2O, 25°C; c. KOH

A (phenylthio)acetate of geraniol (24) was used to synthesize lactone 12 [17]. Allylic oxidation of 24 by SeO2 followed by mild hydride reduction led to unsaturated monosubstituted diol 25. Halogenation and further cyclization of bromide 26 gave in high yield S-containing macrolide 27, desulfurization of which gave the desired pheromone 12.

1 g, h

O

a

O

99% 24

38%

S C H 6 5

b, c

d

OH

Br

100%

46% 25

26

28

O

j O O

OCOCH2Br

S C6H5

27 f 45%

i 47%

O

52%

OBz O

O

e

O

78%

29

12

a. PhSCH2COCl, Py; b. SeO2; c. Na(CN)BH3; d. CBr4, PPh3; e. NaH, HMPA; f. Ni-Ra, EtOH; g. BrCH2COBr, Et3N; h. SeO2 on SiO2, t-BuOOH; i. SmI2, THF, BzCl, DMAP; j. SmI2, THF, HMPA, pivalic acid

In another synthesis [21] of lactone 12, aldehyde 28, the product of allylic oxidation of geraniol bromoacetic ester by t-BuOOH in the presence of a catalytic amount of SeO2 on silica gel, underwent lactonization in the presence of SmI2 to cyclic benzoate lactone 29, further deacyloxylation of which by SmI2 and pivalic acid gave the desired compound 12. A key step in the approach to a diastereomeric mixture of (3S,6R/S)-(-)-methyl-6-isopropenyldecanol acetate (34), a component of the sex pheromone of the California red scale (Aonidiella aurantii) [22], was the cross coupling of n-butyllithium with unsaturated bromoacetate 33, which was prepared by allylic oxidation of the double bond of citronellylacetate (31) with subsequent two-step substitution of the hydroxyl by bromine. The coupling with n-butyllithium proceeds by an SN21 mechanism and produces an isopropenyl substituent. b R 30, 31 a

c, d

HO

e, f

Br

OAc

45% 32

33

34

30: R = OH 31: R = OAc

a. Ac2O, Py; b. SeO2, t-BuOOH; c. TsCl, Py; d. LiBr, Me2CO; e. n-BuLi, CuI; f. BF3 ⋅Et2O

Poppe et al. [23] proposed an interesting approach using allylic oxidation by SeO2 for the synthesis of (5S,9S)-5,9dimethylheptadecane (40a), the principal component of the sex pheromone of the pear leaf blister moth (Leucoptera scitella). Its (5R,9S)-isomer (41a) and (5SR,9S)-40b and (5R,9S)-dimethylpentadecanes (41b) are possible sex attractants of the coffee leaf miner moth (Perileucoptera coffeella).

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(R, S)-30

b

a 21%

c R

O 35

36a, b

d 70% O

+

HO

37a, b

38a, b

e 50% f

+

R

83% 39a, b

R 41a, b

40a, b

R = Et (a), H (b)

a. BY; b. R(CH2)4P+Ph3Br-, NaOEt; c. SeO2; d. PDC; e. n-PrP+Ph3Br-, NaOEt; f. H2, Pd-C

(R)-Citronellal (35), which was prepared by incubation of racemic citronellol (30) with Baker's yeast, was coupled with n-hexyliden or n-butylidenylides to produce the corresponding dienes 36a and b, allylic oxidation of which gave a mixture of aldehydes 37a and b and the accompanying alcohols 38a and b, which were converted to the required aldehydes 37a and b by the standard method. Then lengthening of the C chain by Wittig olefination with n-propylidenphosphorane and exhaustive hydrogenation of the resulting trienes 39a and b completed the synthesis of the target pheromones 40a and b and 41a and b.

MONOTERPENOIDS FUNCTIONALIZED BY EPOXIDATION Selective epoxidation of the isopropylidene group of geraniol derivatives and similar structures is often used in synthetic approaches to insect pheromones from monoterpenoids. m-Chloroperbenzoic acid (MCPBA) is used most often for this. Thus, epoxidation of the double bond of geraniol tetrahydropyranyl ether (42) was used to isomerize it into the terminal position using aluminum isopropylate in the synthesis of ferrulactone (12) [24]. Claisen rearrangement of the resulting allylic alcohol 43 gave the trisnorfarnesane structure in 44. Its hydrolysis gave the key hydroxyacid 11, which was cyclized under standard conditions into the desired pheromone 12. b

a OTHP

77%

c

99%

O

43 OH

42

11

OT HP

O

f

d, e EtO2C

95%

O

98%

44

12

a. MCPBA, AcONa; b. (i-PrO)3Al; c. MeC(OEt)3, EtCO2H, 135°C; d. PPTS; e. KOH; f. BID, PPh3, PhMe

An analogous approach with isomerization of the double bond of another monoterpenoid, nerol (45), was used to prepare of 3,7-dimethyl-2Z,7-octadien-1-ol propionate (48), a component of the California red scale (A. aurantii) pheromone [25, 26]. Here the double bond in the terminal position was substituted by treatment of epoxyester 46 with t-butylhypochlorite, which produced allyl alcohol 47, deoxygenation of which gave the required 48. c

a, b

45: R = H

OR

92%

O 46: R = COEt

a, d, e OH 47

a. EtCOCl, Et3N; b. MCPBA; c. t-BuOCl; d. MsCl, Et3N; e. LiAlH4

620

48

OCOEt

Epoxidation of the ∆6,7-bond of geraniol was used to shorten the C chain in the synthesis of racemic 9-methylgermacrene-B (56), a pheromone produced by male sand flies (Lutzomyia longipalpis) [27]. Compound 1 was fragmented through intermediate epoxide 49 of the corresponding TBS ether using periodic acid. The resulting aldehyde 50 was alkylated by a Grignard reagent to allyl alcohol 51, Claisen rearrangement of which gave the trimethyl-branched diene α,ω-difunctionalized 52. Then 52 was converted by known methods to aldehyde 53, which was transformed successively through cyano derivatives 54 and 55 into the required pheromone 56. a

b

1

c

OT BS

d

O

71%

55%

37% from 1

O 49

f, g

e

Et 2OC

51 OH

50

52 CN

h EtO2C

OEE

i-k

Cl

O

Cl

Cl

53 l, m

n, o

CN

17% from 52

54

10%

OEE 55

56

a. TBSCl, DMF; b. MCPBA; c. HIO4; d. MeC=CMeMgBr, THF; e. MeC(OEt)3, EtCO2H, ∆; f. PPTS, MeOH; g. PPh3, CCl4; h. DIBAH, -78°C; i. TMSCN, KCN, 18-crown-6; j. BnMe3NF, THF, H2O; k.CH2=CHOEt, PhH; l. NaHMDS; m. PPTS; n. NaOH, Et2O; o. Me2CBr2, Sm, CrCl3, SmI2, THF

An analogous method for shortening the C chain was used in a convergent synthesis of (3S,11S)-3,11-dimethyl-2heptacosane (70), a component of the sex pheromone of male German cockroaches (Blattella germanica), using (R)-citronellol (30) and ethyl-(R)-3-hydroxybutyrate (61) as starting materials [28].

(R)-30

a, b

c

d

(CH2)15Me

O

57 g, h

58 i, e, a, f

CO2Et

CO2Et

I

62

61

TBSO

66

60 n

l, m OEE

63 o

O

O 64

OH

p, q O

O

(CH2)15Me

CO2Me O

67

I

59

j, k

OEE

OH

OH

e, a, f

O

65 r-t

68

u, v (CH2) 15Me

(CH2)15Me OH

69

O

70

a. TsCl, Py; b. Me(CH2)13MgBr, Li2CuCl4; c. MCPBA; d. HIO4; e. LiAlH4, Et2O; f. NaI, Me2CO; g. LDA, THF, HMPA; h. MeI, -60°C; i. CH2=CHOEt, TsOH; j. MeCOCH2CO2Me, K2CO3; k. NaOH, l. AcOH, THF, H2O; m. recristallization from hexane; n. TBSCl; o. CO(OMe)2, NaH, dioxane; p. 60, K2CO3, MeCOEt; q. NaOH, n-Bu4NOH, THF, H2O; r. NaBH4; s. MsCl, Py, DMAP; t. LiBEt3H; u. HF, DME, H2O; v. PCC

Epoxidation of alkene 57, which was produced by extension of the C chain of starting (R)-30 by a Schlosser—Grignard reaction, followed by periodate cleavage of epoxide 58 led to methyl-branched aldehyde 59. Then 59 in three standard steps was converted to the first chiral synthon, iodide 60. The second chiral synthon was constructed starting with hydroxyester 61. Its stereoselective methylation in the presence of two equivalents of lithium diisopropylamide gave a mixture of the syn- and anti-isomers of methyl-branched 62 in

621

a 4:96 ratio. After protection of the hydroxyl, the ester was transformed in three steps into an iodomethylene group. Lengthening of the C chain in the resulting 63 by alkylation with acetoacetic ester gave ketoether 64. Removal of the protecting group gave hemiacetal 65, recrystallization of which enabled the syn-isomer to be completely removed. Treatment of 65 with t-butyldimethylsilylchloride and methoxycarbonylation of the intermediate ketoether 66 gave the β-ketodifunctionalized 67. Alkylation of 67 through the sodium derivative by iodide 60 followed by deoxygenation gave silyl ether 69, deprotection of which produced the desired pheromone 70. An original approach to the synthesis of ipsdienol (73), an aggregation pheromone of Ips bark beetles, was based on myrcene (71) [29]. Regioselective epoxidation of triene (71) at the ∆6,7-bond formed the epoxide 72, in which the oxirane ring was opened by MeMgI at room temperature after 1 d to give the target pheromone 73. a 73%

OH

b 64%

O

73

72

71

a. MCPBA; b. MeMgI

Epoxidation of geraniol and similar monoterpenoids can also be carried out selectively at the ∆2,3 bond by treatment with t-BuOOH in the presence of catalytic amounts of VO(acac)2. Such an approach was used to prepare racemic 3,7-dimethyl2-oxo-6-octen-1,3-diol (77c), which may be active as an aggregation pheromone of the Colorado beetle (Leptinotarsa decemlineata) [30]. OH

O a, b OH 1

OAc

OH OH

c, d OH

92% 74

75a - c

c, d

OH

e

OT BDPS

OH

f

OT BDPS

OH

O

76a - c

g OH

OH a

OH O

78a, b

79a, b

O a=R b =S c = racemic

77a - c

a. t-BuOOH, VO(acac)2; b. Ac2O, Py; c. HClO4, DMF; d. K2CO3, MeOH; e. TBDPSCl; f. DMSO, (COCl)2, Et3N; g. n-Bu4NF

Regioselective epoxidation followed by acylation converted 1 into epoxyacetate 74, opening of the oxirane ring of which and hydrolysis of the acetate gave triol 75c. After selective protection of the primary hydroxyl, Swern oxidation of the secondary hydroxyl in 76c followed by deprotection produced 77c as a racemic mixture. Separation of the racemate 77c by chiral GC gave two compounds, one of which was identical to the natural pheromone. The isomers (R)-77a and (S)-77b were obtained from the linalool enantiomers (R)-78a and (S)-78b, respectively, to establish the absolute configuration of the natural pheromone. In particular, epoxidation of the terminal double bond of the (R)-isomer and then opening of the oxirane ring in (R)-79a gave triol (R)-75a. By analogy for the racemic species, the desired (R)-77a was obtained from (R)-75a. The isomer (S)-77b was synthesized by the same synthetic pathway from (S)-78b. Biological tests showed that unsaturated oxodiol (S)-77b corresponded completely with the natural aggregation pheromone produced by male Colorado beetles whereas its (R)-isomer, i.e., 77a, was inactive. Regioselective epoxidation of the exocyclic double bond of carvone (80) by MCPBA was used to introduce a hydroxyl in establishing the absolute configuration of natural α-phellandren-8-ol (p-mentha-1,5-dien-8-ol) (85), a monoterpene isolated from bark and pine beetles (Ips sexdentatus Born, I. acuminatus Gyl, Dendroctonus ponderosae Hopkins) [31]. Both

622

enantiomers of 85 were synthesized from (R)- and (S)-carvone (80), respectively. Reduction of epoxide 81 involved the oxo group also, which was regenerated from the resulting diol 82 by selective oxidation with MnO2 to ketoalcohol 83. Deoxygenation occurring with deprotonation of the ketone by the Bamford—Stevens method [32] led through tosylhydrazone 84 to the desired dienol (S)-85. (R)-85 was prepared analogously from (R)-carvone (80). O

O

a quant.

*

*

NNHT s

d 73%

82%

*

OH 84

OH 83

82

e 67%

*

*

OH

81

O

c

*

O

80

OH

b quant.

OH 85

a. MCPBA; b. LiAlH4; c. MnO2; d. TsNHNH2, HCl, MS 4 A°, THF; e. MeLi

The longhorn beetle (Vesprus xatarti) is a dangerous grape pest. Both enantiomers of 10-oxoisopiperitenone (vesperal) (93), a component of its sex pheromone, were synthesized in high optical purity and with a prepared C10-template starting from (S)- and (R)-limonenes (86) [33] in order to determine its absolute configuration. OH SeP h

O a-c

d quant.

30% * 86

*

f

e 86%

*

OH

87

*

OAc

OAc 89

88

+

87%

*

SeP h OH

90

* OH

OH 91

OH h

g

91

65%

54% *

OH 92

O

*

O 93

a. n-BuLi, TMEDA; b. O2; c. Na2SO3; d. Ac2O, Py; e. MCPBA; f. Ph2Se2, NaBH4; g. H2O2, THF, H2O; h. PCC, NaOAc

The hydroxyl was introduced on C-10 of diene (R)-86 using the classical method of Crawford et al. [34] involving lithiation and oxidation of the organometallic intermediate, which gave unsaturated alcohol 87 in 30% yield without racemization at C-4. The second oxygen function was introduced at C-3 via organoselenium intermediate 91. For this, a mixture of diastereomers of 89, prepared from 88 using MCPBA, was treated with phenylselenide anion as before [35]. In parallel with hydrolysis of the acetate, this gave a mixture of diols 90 and 91, which were separated by chromatography. Then 91 was oxidatively cleaved by hydrogen peroxide to give bisallyl alcohol 92, oxidation of which using Corey reagent gave the target (S)-vesperal (93). The (R)-isomer of 93 was synthesized from (S)-limonene by an analogous scheme. According to tests, the (R)-isomer was completely identical to the natural pheromone. The examples given above [16-23] involve allylic oxidation of an isopropylidene group by SeO2. Italian chemists [36] proposed a scheme in which the first step was epoxidation of (S)-citronellol (30) after protection of the hydroxyl in a simple and effective synthesis of (3S,6R/S)-3-methyl-6-isopropenyl-9-decen-1-ylacetates (99), components of the sex pheromone of California red scale (A. aurantii). Subsequent isomerization of oxirane 94 by aluminum isopropylate gave unsaturated allyl hydroxyether 95. The butenyl radical was introduced into the molecule using a Grignard reaction and benzothiazolesulfide as the leaving group, which was added to substrate 95 by treatment with benzothiazole disulfide in toluene in the presence of triphenylphosphine. The resulting mixture of isomeric sulfides 96 and 97 was converted completely by irradiation into 97. Alkylation of 97 by 3-buten-1-ylmagnesium bromide in the presence of Cu(I) bromide proceeded through an SN21 mechanism exclusively to unsaturated ether 98, from which the required acetate 99 was produced as a mixture of two diastereomers. It

623

should be noted that conditions for removing the benzyl protecting group by hydrogenolysis while not affecting the double bonds in the molecule were successfully found.

(S)-30

a, b

c

O

d

OBn 93%

87% 94

95

f 89%

+ BtzS 96

OH

BtzS

OBn

97

98

e quant.

g, h OAc 99

a. BnBr, NaH; b. MCPBA; c. (i-PrO)3Al; d. BtzS2, PPh3; e. hv; f. CH2=CH(CH2)2MgBr, CuBr; g. H2, Pd-C; h. AcCl, Py

Alkaline epoxidation of the conjugated double bond of (R)-carvone (80) was used to contract the ring in a synthesis of (+)-grandisol (110), a component of the boll weevil sex pheromone (Anthonomus grandis) [37]. O

(R)-80

O

a 75%

HO HO

O

b

c, d

MeO MeO

O

e

CO2Me i

f-h

90%

62%

69%

I

CO2Me

78%

78%

103 101

100 CO2R

j, k 78%

CO2R

+ H

106a: R = Me b: R = H

H

104

102

CO2Me H 105

O l

O I

78%

107a: R = Me b: R = H

m

106b

n

OH

35%

H 108

g, o

CN H

109

OH

44%

78% H

p, n H 110

a. H2O2, NaOH, MeOH; b. HCIO4, THF; c. NaIO4, MeOH; d. HC(OMe)3, MeOH, TsOH; e. AcOH, H2O; f. NaBH4; g. TsCl, Py; h. NaI, Me2CO; i. LDA, HMPA, THF; j. LDA, then MeI, THF; k. NaOH, MeOH; l. KI, I2, NaHCO3, CH2Cl2; m. Zn, NH4Cl, EtOH; n. LiAlH4; o. NaCN, HMPA, H2O; p. DIBAH

Opening of epoxide 100 by acid gave ketodiol 101. Periodate cleavage of 101 gave acetalester 102, simple transformations of which produced haloester 104. Enolization of 104 with subsequent intramolecular alkylation provided a path to the required cyclobutane 105. A methyl group of the required configuration was introduced by low-temperature methylation of the corresponding lithium derivative of ester 105, which led to a diastereomeric mixture of 106a,b and 107a,b. Subsequent iodolactonization of this mixture cyclized only the cis-isomer of 106b to form iodolactone 108, which after reduction by zinc dust was regenerated into the single acid 106b. Reduction of 106b gave the alcohol (109), which was converted to the desired pheromone 110 through the cyanide homolog. An isopropylidene group in linear monoterpenoids can be epoxidized both directly by peracids and by dehydrohalogenation of vicinal bromohydrins prepared, for example, using N-bromosuccinimide [38-39]. Thus, whereas Novak et al. [26] proposed using MCPBA to epoxidize the ∆5,6 double bond of nerol propionate (48), Mori [38] used hydroxybromination of nerylacetate (111) followed by dehydrobromination of the resulting bromohydrin 112 under alkaline conditions to give epoxide 113 in the synthesis of threo-(+)-4-methylheptan-3-ol (118), a pheromone of the elm bark beetle (Scolytus multistratus). Periodate cleavage of 113 gave ω-acetoxyaldehyde 114, which was converted by a Huang—Minlon modification into unsaturated acetate 115. The hydroxyl was added regioselectively to C-2 using an organoboron intermediate. Subsequent methylation of C-1 with dimethyllithiumcuprate of epoxide 117 corresponding to diol 116 completed the synthesis of the desired 118.

624

a 111

O

58% from 122

OH

OAc

d

b, c

Br

82%

j, k OH 116

68%

67%

113

112

g, h, i

e, b, f

O 114

115

OAc

l

29%

90%

O

OH

OH

117

118

a. NBS, DMF, H2O; b. KOH; c. Ac2O, Py; d. HIO4; e. N2H4 ·H2O, (CH2OH)2; f. DHF, TsOH; g. B2H6; h. 30% H2O2, NaOH; i. H3O+; j. HBr, AcOH; k. KOH, (CH2OH)2; l. Me2CuLi

Double bonds are also fragmented via cleavage of the corresponding epoxide through an intermediate vicinal alcohol. This approach was used in the synthesis of 13,17,21-trimethyltri-(124a)-, -penta-(124b), and -hepta-(124c)-triacontanes, components of the sex pheromone of the tobacco hornworm (Manduca sexta L.) [39]. a

b

Cl

(CH2)10Me

O

119

120

38% from 120

121

g

c-f

h

O

R 123a - c

122

R

C12H25 124a - c

R = n-C12H25 (a); n-C14H29 (b); n-C16H33 (c) a. AcOCH2CO2Et, base; b. Me(CH2)10CH=PPh3; c. NBS, DME, H2O; d. i-PrONa; e. HClO4; f. Pb(OAc)4; g. R-C(Me)=PPh3; h. H2, Pd-C

Triene 121, the product of Wittig olefination of geranylacetone (120) by n-dodecylidenephosphorane, which was prepared in turn from geranylchloride (119), was used as a substrate for hydroxybromination. It was cleaved by a sequence of hydroxybromination, cyclization—decyclization, and fragmentation reactions using lead tetraacetate to aldehyde 122. Olefination by the appropriate phosphoranes and exhaustive hydrogenation of the resulting trienes 123a,b,c led to the required pheromones 124a,b,c. The methyl ester of acetic acid α-sulfochloride, like NBS, reacts regioselectively with acylic terpenoids. For example, its reaction with geraniol benzyl ether (125) was used to synthesize 3,7-dimethyl-2E,6E-decadien-1,10-diol (129), a component of the sex pheromone of the plain tiger butterfly (D. chrysippus) [40]. Dehydrohalogenation of the resulting α-chlorosulfide 126 led to an allylsulfide (127) with a terminal double bond. An intramolecular rearrangement of 127 in the presence of base gave .-mercaptoester 128 in good yield, desulfurization of which with further hydride reduction of the ester and deprotection led to the desired diol 129. b

a OBn 125

72%

126 SH

c

Cl

d-f

MeO2C

56% 128

SCH2CO2Me

73% 127

HO

SCH2CO2Me

OH 129

a. MeO2CCH2SCl; b. DMF, 60°C; c. t-BuOK, THF, DMSO; d. Ni-Ra; e. LiAlH4; f. H2, Pd-C

An analogous rearrangement was used to prepare the racemic acetate of 2E,3-methyl-6-isopropenyl-9-decadien-1-ol (135), a component of the sex pheromone of the California red scale (A. aurantii) [41, 42]. Geranylacetate (2) was transformed by a Stevens rearrangement into sulfide 131 through an intermediate trifluoroacetate dimethylsufonium salt 130. Subsequent conversions of sulfide 131, culminating in its oxidation and alkylation of the resulting hydroxysulfone 132 by allylbromide, gave sulfone 133, reductive desulfonylation of which gave alcohol 134, which was easily converted to the target natural product 135.

625

2

a

OAc

90% CF3CO2

b, c

130

131

f, g OH

88%

77 from 2

SMe2

MeO2S

e

MeO2S

d

MeS

h

SO2

c OAc

47%

77%

132

133

134

135

a. (CF3CO)2O, DMSO; b. MeONa; c. Ac2O, Py; d. MCPBA; e. LiAlH4; f. n-BuLi; g. CH2=CHCH2Br, THF, HMPA; h. Na, NH3

OZONOLYSIS Yet another oxidative method is ozonolytic cleavage of double bonds. It has also been widely used to synthesize insect pheromones [43-54] and is a convenient and practical method for preparing O-containing compounds. An example is the synthesis of all stereoisomers of 3,13-dimethylheptadecane (145a-d), the principal component of the western false hemlock looper (Nepytia freemani) sex pheromone [43]. The key step in this scheme was alkylation of phenylsulfones (R)- and (S)-140 with a reactive α-methylene that were prepared from the corresponding enantiomers citronellol, (R)- and (S)-30.

(S)-,(R)-30

a, b

c

*

83%

a, d

HO

70%

137

a, g, h

HO 90%

5

61% HO

81%

136 f

e

I

PhSO2

72%

139

138 * 5

(S)-140, (R)-140

a. TsCl, Py; b. EtMgBr, Li2CuCl4, THF; c. O3, then NaBH4; d. NaI, Me2CO; e. n-BuLi, HC≡CH(CH2)3OH, THF, HMPA; f. H2, PtO2, EtOH; g. PhSH, NaOH, MeOH; h. MCPBA

The aforementioned organosulfur compounds were synthesized starting with alkylation of intermediate citronellyltosylate with ethylmagnesium bromide. Ozonolytic cleavage of the double bond of dimethyl-branched alkene 136 followed by hydride reduction of the peroxide ozonolysis products led to alcohol 137, which was transformed into iodide 138. Then, 138 was used to alkylate the lithium derivative of 4-pentyn-1-ol. Exhaustive hydrogenation, phenylthiylation of 139, and oxidation by MCPBA gave the corresponding synthons (R)- and (S)-140. The second building block was synthesized using the required enantiomer of methyl 2-methyl-3-hydroxypropionate (141) as the substrate. Through a series of known transformations [44], including preparation of optically active 2-methylbutanol (142), it was converted to iodide 143. HO *

a CO2Me

(S)-, (R)-141

b, c

*

OH

47% from141

142 d

e, f 5

144

I

(R)-,(S)-143

SO2Ph

(R)-140 + (S)-143 72%

*

85% 145a

5

a. Ref. [44]; b. TsCl, Py; c. NaI, Me2CO; d. n-BuLi, THF, HMPA; e. Na, Hg, Na2HPO4, EtOH; f. H2, PtO2

626

Combination of the lithium derivatives of sulfone 140 and iodide 143 through a cross-coupling reaction gave sulfone 144 with two optically pure chiral centers. Desulfurization and hydrogenation completed the synthesis of the desired pheromones 145a-d. A synthesis of all stereoisomers of 10,14-dimethyloctadec-1-ene (152a-d), 5,9-dimethyloctadecane (153a-d), and 5,9-dimethylheptadecane (40a, 41a, and 155a,b), components of the apple leafminer (Lyonetia prunifoliella) sex pheromone, was proposed starting with citronellol enantiomers (30) and methyl 2-methyl-3-hydroxybutanoate (149) [45]. a, b

(S)-, (R)-30

*

AcO

66%

c, d

OH

73%

HO

147

h CO2Me

*

*

*

O

7 (S)-, (R)-148

Br

(R)-,(S)-151

150 b

j-l

152a

c, i

OH

56%

(S)-,(R)-149

(S)-151

40%

7

146 *

e-g

*

AcO

36%

60%

7

152a

C9H19

153a

c, n

m 154a

152b

7 OH

84% from 152a

40a

C8H17

m, c, n 155a

C8H17

a. Ac2O, Py; b. O3, then NaBH4; c. TsCl, Py; d. H2C=CH(CH2)4MgBr, Li2CuBr2 ·Me2S ·PhS, THF, HMPA; e. KOH; f. SiO2 AgNO3; g. PCC; h. ref. [47]; i. LiBr; j. Mg, then 148; k. MsCl, Py; l. LiBEt3H, THF; m. H2, PtO2; n. LiAlH4, THF

Required isomers of 148 were prepared from (R)- and (S)-citronellol (30), which was first converted to acetoxyalcohol 146 [46] and then alkylated through the corresponding tosylate [47]. Hydrolysis of the resulting unsaturated acetate 147 with subsequent oxidation led to (S)- and (R)-aldehydes 148, respectively. Synthon 151 was prepared by a known method from the corresponding enantiomers of methyl 2-methyl-3hydroxybutanoate (149) [47] through optically active 2-methylhexanol (150) [47]. Combinations of possible cross-couplings of (S)- and (R)-isomers of synthons 148 and 151 with subsequent simple transformations that did not affect the chiral centers led to all stereoisomers of the olefinic components of pheromone, 152a-d. Catalytic hydrogenation of olefins 152a-d produced the alkane components of the pheromone, 153a-d. The remaining stereosiomers of the pheromone 40a, 41a, and 155a,b were synthesized from 152a-d by ozonization followed by hydride reduction and deoxygenation of the intermediate alcohols 154a-d. Two oxidative methods were used to prepare key blocks for a convergent synthesis of (+)-cembrene A (165), a highly effective tracking pheromone of termites (Nastitermes exitiosus) [48]. a, b

c

1

SO2Ph 156

d, e

O

82%

RO

SO2Ph

90% 157

158, 159 158: R = H 159: R = THP

a. PBr3; b. NaSO2Ph, DMF; c. O3, CH2Cl2, then Me2S; d. NaBH4, MeOH; e. DHP, TsOH

A key step in the synthesis of the first synthon 159 was ozonolytic fragmentation of geranylphenylsulfone 156. For some reason the peroxide ozonolysis products of 156 were reduced in two steps, first to the aldehyde and then to the alcohol. However, in this instance it would have been simpler to convert immediately the peroxide ozonolysis products into the required alcohol 158 using NaBH4. 627

To prepare the second synthon 161, oxidation of geranylacetone 120 using sulfuryl chloride proceeded with allyl rearrangement to give secondary chloroketone 160, which contained the isopropenyl group. By introducing dithiane protection of the oxo group, this block was prepared for cross-coupling with the first synthon, which was carried out successfully in aqueous base in the presence of an interphase-transfer catalyst. Further standard transformations of the resulting sulfone 162 led to the acyclic precursor oxodithiane 164, intramolecular cyclization of which gave the required sesquiterpene 165. a

120

S

b O

82%

Cl

90% Cl

160 SO2Ph

d-f

T HPO

c 65%

S

161

g

HO

65%

62% 162

163 h 82%

O 164

165

a. SO2Cl2, Na2CO3, CH2Cl2; b. HS(CH2)3SH, BF3·Et2O; c. NaOH, 159, n-Bu4NBr; d. PPTS; e. Na, Hg, NaH2PO4; f. HgCl2, CaCO3; g. PCC; h. TiCl3-AlCl3 (3:1), Zn-Cu, DME

The principal component of the abdominal gland secretion of the aposematic shield bug (Cantao parentum White) [Hemiptera: Scutelleridae] was established as (2S,4R,6R,8S)-trimethyl-1,7-dioxaspiro[5.5]undecane (174). This was the first example of a branched spiroacetal in the insect kingdom [49]. O

a O

O

b

e, f

d

c O

CO 2H

OH

167

166 O

g, h 169 I

NNMe2

i

170

j

171

NNMe2 OT HP

O

k

OT HP

m

l O

168

OH

OH

O

OT HP

HO l

m

O

e, n

O

174

OT HP

O

O

173 HO

I 172

a. Ref. [52]; b. MeLi, -78°C; c. HO(CH2)2OH, TsOH; d. O3, -78°C, then NaBH4; e. TsCl; Py; f. NaI; g. t-BuOK; h. AcOH, 80°C; i. H2NNMe2, AcOH; j. LDA, -78°C, then 172; k. SiO2; l. HCl; m. AD-mix β, 0°C; n. LiAlH4

The enantioselective synthesis of this unique spiroacetal was carried out [50] starting with (R)-(+)-pulegone (166) through an intermediate acetonide of unsaturated ketone 167. Successive ozonolysis of the double bond in 167 and reduction

628

of the peroxide products gave hydroxyketal 168, dehydrogenation of which through the corresponding iodide 169 gave after acid hydrolysis ketoolefin 170. The required C chain was constructed by alkylation of the lithium derivative of the corresponding tosylhydrazone 171 by optically active substituted iodohydrin 172 [51]. The hydroxyl was introduced by oxidation of the double bond using the chiral osmium reagent AD-mix β. Simultaneous incorporation into the precursor of alcohols at the δ and δ′ positions relative to the oxo group caused ketal formation, which led to hydroxyketal 173. Deoxygenation of 173 gave the target spiroketal 174. Ozonolysis was used as a functionalization method in the preparation of both synthons in a convergent synthesis of (11R,17S)-dimethylhentriacontane (182), a communication pheromone of ants Camponotus vagus [53]. Thus, phosphonium salt 178 was synthesized starting from (R)-citronellol (30), ozonolysis of which with subsequent reduction gave a mixture of hydroxyaldehyde 175 and hemiacetal 176. Wittig olefination of this mixture gave (Z)-unsaturated alcohol 177, which was converted further through the bromide to salt 178. OH a

(R)-30

OH

O

98%

O

+

OH

b 5

75%

175

(R)-178 g

a 10

92% 179

P Ph 3Br 5

(R)-177

176

e, f

(R)-35

c, d 73%

O 30%

98%

OT s

(R)-180 h, i

4

78%

181

12

8

182

a. O3, -78°C, then Me2S; b. n-C7H15PPh3Br, n-BuLi, THF; c. CBr4, PPh3; d. PPh3, MeCN; e. Mg, Me(CH2)11Br, 10% (CH2Br)2, THF; f. TsCl, Py; g. (R)-178, n-BuLi, THF; h. LiAlH4, NaH; i. H2, Pd-C

The second synthon was synthesized by ozonolytic cleavage of intermediate tosylate 179, the esterification product of (R)-citronellal (35) alkylated with n-dodecylmagnesium bromide. The resulting aldehyde (R)-180 underwent Wittig olefination by the phosphorane from phosphonium salt 178. Reduction of tosyloxydiene 181 completed the synthesis of the desired pheromone 182. This same group [54] proposed a convergent synthesis of [3H2]-(11S,17R)-dimethylhentriacontane (187), a tritiated derivative of the C. vagus ant communication pheromone. Like in the scheme given above, the synthesis was performed from the two synthons tosyloxyaldehyde (S)-180 and a transformation product of aldehyde (S)-35 prepared by a previously described method [53]. Saturated bromide 184, prepared from (S)-citronellol (S)-30, was used for phosphonium salt (S)-185. Wittig olefination of (S)-180 gave tosyloxyolefin 186, into which tritium was introduced using Wilkinson rhodium catalyst after deoxygenation. These transformations produced the target compound 187 in 22% overall yield calculated for starting (S)-35.

(S)-30

a 98%

b

(S)-177

89%

OH 6

c

Br

98%

d

PPh 3Br

75%

183

184

185 3

OT s a

(S)-35

90%

(S)-180

f, g

e 28%

6

10

186

87%

8

3

H 12

H 187

a. Ref. [54]; b. H2, (PPh3)3RhCl, THF; c. CBr4, PPh3; d. PPh3, MeCN; e. 206, n-BuLi, THF; f. LiAlH4; g. Tr2, Rh(PPh3)3Cl, THF

629

(R)-4-Menthenone, (R)-189, which is accessible from l-menthol (188), presents excellent synthetic possibilities for synthesizing optically pure methyl-substituted natural compounds [55-60]. We developed ozonolytic fragmentation of enone (R)-189 to prepare the versatile optically pure bifunctional synthon (R)-190, methyl (R)-5,5-dimethoxy-3-methylpentanoic acid [61]. b

a 87%

OH

O

c, d

96%

OAc

e

87%

188

O

87%

MeO

O

MeO

OMe 190

(R)-189

a. PCC; b. Ac2O, TsOH; c. Br2, CCl4; d. MeOH; e. O3, c-C6H12 (or CCl4)-MeOH then MeOH, TsOH

As an example of the use of this compound, we proposed convenient approaches to the synthesis of (R,R)-17- and (R,S)-17-4,8-dimethyldecanal, components of the aggregation pheromone of Tribolium flour beetles. OMe

O

MeO

(R)-191

OMe 190 c, f, g

Br

l, f, m O

O

a-e

(S)-189 n, c, f, g

T sO

Br

OBn 192

(R)-193

(S)-194

h

i-k

i-k

(R, R)-17

OBn (R, R)-195

OBn

(R, S)-17

(S, R)-195

a. DIBAH; b. BnCl, KOH; c. PPTS, H2O; d. NaBH4; e. TsCl, Py; f. N2H4·H2SO4, KOH; g. Ag2O, then Br2; h.. Mg, Li2CuCl4; i. H2/PtO2; j. PBr3/Py; k. Mg, then DMF; l. H2O2, NaOH, MeOH; m.. PCC; n. O3, c-C6H12 (or CCl4)-MeOH then MeOH, TsOH

It was noted in a convergent synthesis of (R,R)-17 that both optically active key synthons, toslyate 192 and bromide (R)-193, were synthesized from the aforementioned optically pure substrate (R)-189. Synthon 192 was prepared using chemically selective hydride reduction of the ester in acetalester 190 to the alcohol, benzylation of which enabled the other end of the molecule to be transformed into the tosylate. The second building block 193 was synthesized starting with conversion of the deprotected carbonyl in acetalester 190. Subsequent Huang—Minlon reduction of the aldehyde and simultaneous hydrolysis of the ester produced the required bromide (R)-193 after Hunsdiecker decarboxylation. The key step included catalyzed alkylation of the tosyl group in 192 by the Grignard reagent from bromide (R)-193, which gave dimethyl-branched hydroxybenzyl ether 195, subsequent simple transformations of which produced the desired aldehyde (R,R)-17. The (R,S)-17 isomer was synthesized analogously. Instead of bromide (R)-193, its (S)-192 isomer prepared from (S)-4-methen-3-one, (S)-189, was used. (S)-189, in turn, was the configuration inversion product of (R)-189. (R)-189 was transformed into (S)-189 using Warton reduction of epoxyketones [62], which occurs with allyl rearrangement and forms allyl alcohols, oxidation of which gives unsaturated conjugated enones. Thus, carrying out these reactions caused inversion of the configuration of the asymmetric center in the starting chiral enone (R)-189. Furthermore, using the synthesis of (14S)-methyloctadec-1-ene (201) [63], the sex pheromone of the apple leaf miner (Lyonetia clerkella), as an example, the novel synthetic possibilities of optically pure (R)-189 were demonstrated. These were based on the susceptibility of conjugated enones for selective 1,2-addition of organometallic reagents with subsequent oxidative rearrangement of the resulting tertiary allyl alcohols by Cr(VI) [64, 65]. Ozonolytic decyclization of the resulting

630

(S)-ethylmenthenone (196) and subsequent methanolysis gave ketoester 197, Huang—Minlon deoxygenation of which accompanied by saponification of the ester gave (3S)-methylheptanoic acid (198), which was converted by standard methods through alcohol 199 and tosylate 200 into the target pheromone 201. a O

O

c

b OH

O

d, e

MeO

O

197 196

(R)-189 O

f, g

h RO

HO

11

199, 200

198

201

199: R = OH; 200: R = OTs

a. EtLi; b. PCC; c. O3, c-C6H12-MeOH then MeOH, TsOH; d. N2H4·H2SO4, KOH; e. KOH; f. LiAlH4; g. TsCl, Py; h. H2C=CH(CH2)9MgBr, Li2CuCl

All aforementioned methods are used "classically" in organic synthesis. However, rather specific reagents, for example oxidizers based on Os(VIII) oxide [66-68] or singlet oxygen [69], are used just as often in constructing molecules. The use of OsO4 was proposed for preparing vicinal diol 202 from geraniol acetate (2) in the syntheses of suspensolide (214), anastrephin (215), and epianastrephin (216), components of the pheromone of the Caribbean fruit fly (Anastrepha suspensa Loew), a very dangerous pest of citrus in Central and North America [66]. The C skeleton of the key synthon hydroxyacid 213, which contains two trisubstituted double bonds, was constructed using the transformations shown in the scheme.

1

a 84%

2

c

b OAc HO OH

j 72%

206 Ts n

Ts 210

SPh MeO2C

HO2C

207 o

204

k 93%

h, i 86%

205

m

l, e, f

MeO2C

Br

58% from 208

208 p, q, r

HO2C

OH

95%

203

MeO2C

g

Br

45% from 202

202

OT HP

d-f

O

209 s

HO2C

OH 213

212

O

9%

O

t 78% H

214 H

O

+

O O

O 215

216

a. Ac2O, Py; b. OsO4, N-methylmorpholine-N-oxide; c. NaIO4; d. Li(t-BuO)2AlH; e. TsCl, Py; f. LiBr, Me2CO; g. LiC≡CH, NH3, DMSO; h. TsOH, DHP; i. n-BuLi, THF, then MeOCOCl; j. PhSH, NaOH, MeOH; k. MeMgBr, CuI, THF; l. LiAlH4; m. TsNa, DMF; n. n-BuLi, THF, then CO2; o. Na-Hg, MeOH; p. CH2N2; q. TsOH; r. K2CO3, MeOH; s. EtCO2N=NCO2Et, PPh3, PhH; t. BF3·Et2O

Geranylacetate (2) was oxidized by OsO4 in the presence of co-oxidant N-methylmorpholine-N-oxide followed by periodate cleavage of diol 202 to aldehyde 203, which was converted by standard methods to bromide 204. Subsequent acetylenation of 204 in a mixture of liquid NH3 and DMSO gave enyne alcohol 205. The second trisubstituted double bond was introduced using a literature method [70]. Michael addition of phenylthiol to unsaturated ester 206, prepared by treatment of 205 with methylchloroformate, led to sulfide 207, methylation of which gave (2E)-unsaturated ester 208. Then 208 was converted by standard transformations to sulfone 210. The C chain was lengthened by addition of CO2 to the lithium derivative of 210 with subsequent desulfurization of 211, which gave the THP ester of 212, 631

which was converted to key hydroxyacid 213. Lactonization of 213 by the method of Mitsunobu et al. [71] gave the target suspensolide 214. The previously described [72] acid-catalyzed cyclization of 214 gave racemic mixtures of (±)-215 and (±)-216. The resulting compounds were purified by chromatography and separated using diastereomeric amides. The use of chromic anhydride as a co-oxidant enabled Monteiro and Schpector [67] to fragment (R)-citronellol (30) protected as the methyl ether (217) at the double bond to give ω-methoxyacid 218, which was used to synthesize (+)-grandisol (110). In this synthesis, the use of successive bicyclization with subsequent partial decyclization was proposed. a

b, c

(R)-30

OMe

SO2P h

O

SO2Ph

219

O SO2P h h

220

SO2P h

HO i

f

PhO2S

O 218

g

N2

e

PhO2S O

217 O

d

MeO

O SO2Ph

SO2Ph j

k, l

+ OMe

I

221

223

222 OH SO2P h

m

226

O

OH n

HO

225

224

+ p

o

HO

110

A=



BF4

N Et

227

HO

O

228

a. NaH, then MeI, DME; b. OsO4, CrO3, Me2CO; c. H+, MeOH, CH2Cl2; d. BnONa, THF, DMSO; e. A, NaN3, NaOAc, MeOH; f. Rh2(OAc)4, PhH; g. NaI, TMSCl, MeCN; h. NaH, THF; i. MeMgI, THF, Et2O; j. SOCl2·Py; k. H2O2, HCO2H, 100°C; l. NaOH, MeOH; m. t-BuOK, DMSO; n. Na-Hg; o. NaIO4, RuCl3, H2O; p. Ref. [68]

Disubstituted 218 was transformed into ketosulfone 219 and then into diazo derivative 220 in order to introduce the tertiary and quaternary C atoms of the required configuration. Carbenoid cyclization of 220 proceeded stereoselectively and was catalyzed by rhodium acetate to give cyclopentane sulfone 221. Substitution of the methoxy by halide followed by another cyclization with asymmetry generation gave the key cyclobutane fragment of the target molecule. 1,2-Addition of methylmagnesium iodide to the ketone of 223 produced tertiary alcohol 224, which was transformed into unsaturated sulfone 226. The cyclopentane ring was opened by treatment of 227 with a mixture of ruthenium chloride and sodium periodate. The resulting ketoacid 228 was converted by a known method [68] into the desired pheromone 110. Oxidation by singlet oxygen of myrcene (71), which occurred with allyl rearrangement of the trisubstituted double bond, was used in a three-step synthesis of farnesene (231), a component of the fire ant (Solenopsis invicta) tracking pheromone [69]. Claisen rearrangement of the resulting triene alcohol 229 gave ester 15, low-temperature hydride reduction of which with subsequent Wittig olefination of aldehyde 230 by isopropylidenetriphenylphosphorane led to the desired sesquiterpene 231.

71

b

a 31%

OH

91% 229

c

15

d

O

80% 230

231

a. O2, h, n-Bu4 NBr; b. CH(OEt)3, H+, 139°C; c. DIBAH, -78°C; d. i-PrPh3P+I -, n-BuLi

The synthetic pathway for α -geranylpropionate (235), a component of the sex pheromone of the San Jose scale (Quadraspidiotus perniciosus), that is based on isomerization of the isopropylidene groups of geraniol into an isopropenyl group is interesting [72, 73]. Two approaches were used for this. The first went through allyl chloride 233, which was prepared by low-temperature chlorination of geranylpropionate 232 and subsequent dehalogenation; the second, through the dimethylsulfonium salt of geranylpropionate 234 with subsequent electrolytic reduction.

632

a OCOEt

98%

232 b

OCOEt Cl

233

c 62%

89% d OCOEt

+

Me2S ClO4



234

OCOEt

20% 235

a. SO2Cl2, CH2Cl2, -60°C; b. DMSO, (CF3CO)2O, LiClO4; c. Zn, NiCl2, PPh3, NaI, DMF, H2O; d. e−

Thus, the literature indicates that oxidative methods are used in a wide range of transformations of monoterpenoids. Chiral substrates can be functionalized with retention of the absolute configuration of the asymmetric centers. This is one of the fundamental aspects of insect-pheromone synthesis.

REFERENCES 1. 2.

3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.

K. Mori, "The Synthesis of Insect Pheromones," in: Total Synthesis of Natural Products, J. Simon, ed., J. Wiley, New York (1981), Vol. 4. H. J. Bestmann and O. Vostrovsky, in: Chemie der Pflanzenschutz- und Schaedlingsbekaempfungsmittel, Band 6: Insektizide, Bakterizide, Oomyceten Fungizide, Biochemische und Biologische Methoden, Naturstoffe, R. Wegler, ed., Springer-Verlag, Berlin, Fed. Rep. Ger. (1981). C. A. Henrick, R. L. Carney, and R. J. Anderson, in: Insect Pheromone Technology: Chemistry and Application Symposium, 182nd Meeting Am. Chem. Soc., New York, Aug. 1981, ACS, Washington, DC (1982). R. Baker and J. W. S. Bradshaw, Aliphatic and Related Natural Products Chemistry, The Chemical Society, London (1983), Vol. 3. K. V. Lebedeva, V. A. Menyailo, and Yu. B. Pyatnova, Insect Pheromones [in Russian], Nauka, Moscow (1984). H. E. Hummel and T. A. Miller, Techniques in Pheromone Research, Springer-Verlag, New York (1984). V. N. Odinokov and E. P. Serebryakov, Synthesis of Insect Pheromones [in Russian], Gilem, Ufa (2001). R. M. Silverstein, Science, 213, 1326 (1981). V. N. Odinokov and G. A. Tolstikov, Izv. Akad. Nauk Kaz. SSR, Ser. Khim., 44 (1984). A. M. Moiseenkov, K. V. Lebedeva, and B. A. Cheskis, Usp. Khim., 5, 1709 (1984). E. D. Matveeva, A. L. Kurts, and Yu. G. Bundel′, Usp. Khim., 55, 1198 (1986). K. Mori, Tetrahedron, 45, 3233 (1989). S. Chattopadhyay and V. R. Mampadur, J. Indian Chem. Soc., 74, 51 (1994). G. Yu. Ismuratov, M. P. Yakovleva, R. Ya. Kharisov, and G. A. Tolstikov, Usp. Khim., 66, 1095 (1997). V. N. Odinokov, Khim. Prir. Soedin., 12 (2000). B. G. Kovalev and A. M. Sorochinskaya, Khim. Prir. Soedin., 844 (1991). A. C. Oehlschlager, J. W. Wong, V. Verigin, and H. D. Pierce, J. Org. Chem., 48, 5009 (1983). V. N. Odinokov, G. Yu. Ishmuratov, I. M. Ladenkova, R. R. Muslukhov, and G. A. Tolstikov, Khim. Prir. Soedin., 272 (1991). V. N. Odinokov, G. Yu. Ishmuratov, I. M. Ladenkova, and G. A. Tolstikov, Khim. Prir. Soedin., 818 (1990). P. Gramatica, G. Giardina, G. Speranza, and P. Manitto, Chem. Lett., 1395 (1985). T. Moriya, Y. Handa, J. Inanaga, and Y. Masaru, Tetrahedron Lett., 29, 6947 (1988). N. Carniery, D. R. Kelly, W. Kurler, and A. A. Marcondes, Agric. Biol. Technol., 3, 421 (1993). L. Poppe, L. Novak, J. Devenyi, and C. Szantay, Tetrahedron Lett., 32, 2643 (1991). B. A. Cheskis, N. A. Shapiro, and A. M. Moiseenkov, Izv. Akad. Nauk SSSR, Ser. Khim., 2602 (1989).

633

25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67.

634

C. Szantay, L. Novak, A. Kis-Tamas, B. Majoros, I. Ujvary, and L. Poppe, Hungarian Pat. No. 182,391 (1986); Ref. Zh. Khim., 130114P (1987). L. Novak, L. Poppe, A. Kis-Tamas, and C. Szantay, Acta Chim. Acad. Sci. Hung., 118, 17 (1985). J. G. Hamilton, A. M. Hooper, H. C. Ibbotson, S. Kurosawa, K. Mori, S. Muto, and J. A. Pickett, J. Chem. Soc., Chem. Commun., 2335 (1999). H. Takikawa, K. Fujita, and K. Mori, Liebigs Ann. Chem., 815 (1997). O. A. Kozhich, N. E. Pyzh′yanova, G. M. Segal′, and I. V. Torgov, Bioorg. Khim., 9, 1658 (1983). J. E. Oliver, J. C. Dickens, and T. E. Glass, Tetrahedron Lett., 43, 2641 (2002). K. Mori and Y. Igarashi, Liebigs Ann. Chem., 93 (1988). D. Barton and W. D. Ollis, eds., Comprehensive Organic Chemistry. The Synthesis and Reactions of Organic Compounds, Vol. 2, Pergamon, New York (1978). K. Domon and K. Mori, Eur. J. Org. Chem., 3783 (2000). R. J. Crawford, W. F. Erman, and C. D. Broaddus, J. Am. Chem. Soc., 94, 4298 (1972). K. B. Sharpless and R. F. Lauer, J. Am. Chem. Soc., 95, 2697 (1973). V. Calo, L. Lopez, and V. A. Fiandanese, Gazz. Chim. Ital., 120, 577 (1990). K. Mori and K. Fukamatsu, Liebigs Ann. Chem., 489 (1992). K. Mori, Tetrahedron, 3, 289 (1977). P. E. Sonnet, Org. Prep. Proced. Int., 7, 261 (1975). Y. Masaki, K. Sakuma, and K. Kaji, Chem. Lett., 1061 (1980). A. M. Moiseenkov, V. A. Dragan, and V. V. Veselovskii, Izv. Akad. Nauk SSSR, Ser. Khim., 365 (1989). V. A. Dragan, A. M. Moiseenkov, and V. V. Veselovskii, Izv. Akad. Nauk SSSR, Ser. Khim., 1143 (1989). H. Takikawa, Y. Shirai, M. Kobayashi, and K. Mori, Liebigs Ann. Chem., 1695 (1996). K. Mori and H. Takikawa, Liebigs Ann. Chem., 497 (1991). H. Tamagawa, H. Takikawa, and K. Mori, Eur. J. Org. Chem., 973 (1999). K. Mori and M. Kato, Liebigs Ann. Chem., 2083 (1985). K. Mori and J. Wu, Liebigs Ann. Chem., 439 (1991). W.-D. Li, Y. Li, and Y.-L. Li, Chin. J. Chem., 10, 92 (1992). C. J. Moore, A. Hubener, Y. Q. Tu, W. Kitching, J. R. Aldrich, S. Schulz, and W. Francke, J. Org. Chem., 59, 6136 (1994). Y. Q. Tu, C. J. Moore, and W. Kitching, Tetrahedron: Asymmetry, 6, 397 (1995). M. V. Perkins, M. F. Jacobs, W. Kitching, P. J. Cassidy, J. A. Lewis, and R. A. I. Drew, J. Org. Chem., 57, 3365 (1992). C. G. Overberger and J. K. Werse, J. Am. Chem. Soc., 90, 3525 (1968). D. Pempo, J. Viala, J. L. Parrican, and M. Santelli, Tetrahedron: Asymmetry, 7, 1951 (1996). D. Pempo, J. C. Cintrat, J. L. Parrain, and M. Santelli, Tetrahedron, 56, 5493 (2000). O. Wallach, Liebigs Ann. Chem., 397, 181 (1913). L. A. Yanovskaya and A. P. Terent′ev, Zh. Obshch. Khim., 22, 1598 (1952). V. W. Armstrong, H. C. Najmul, and R. Ramage, Tetrahedron Lett., 373 (1975). H. C. Brown and C. P. Garg, J. Am. Chem. Soc., 83, 2952 (1961). W. Treibs and H. Albrecht, J. Prakt. Chem., 13, 291 (1961). T. Shono, Y. Matsumura, K. Hibino, and S. Miyawaki, Tetrahedron Lett., 1295 (1974). R. Ya. Kharisov, R. R. Gazetdinov, O. V. Botsman, R. R. Muslukhov, G. Yu. Ishmuratov, and G. A. Tolstikov, Zh. Org. Khim., 1047 (2002). P. S. Warton and D. H. Bohlen, J. Org. Chem., 26, 3615 (1961). R. Ya. Kharisov, E. R. Latypova, R. F. Talipov, R. R. Muslukhov, G. Yu. Ishmuratov, and G. A. Tolstikov, Izv. Akad. Nauk, Ser. Khim., 2146 (2003). S. Torii, T. Inokuchi, and R. Oi, J. Org. Chem., 48, 1944 (1983). A. Nangia and G. Prasuna, Synth. Commun., 24, 1989 (1994). K. Mori and Y. Nakazono, Liebigs Ann. Chem., 167 (1988). H. J. Monteiro and J. Zukerman-Schpector, Tetrahedron, 52, 3879 (1996).

68. 69. 70. 71. 72. 73.

F. X. Webster and R. M. Silverstern, J. Org. Chem., 51, 5226 (1986). P. Baeckstroem, L. Li, M. Wickramaratne, and T. Norin, Synth. Commun., 20, 423 (1990). T. Mukaiyama, H. Toda, and S. Kobayashi, Chem. Lett., 535 (1975). T. Kurihara, Y. Nakajima, and O. Mitsunobu, Tetrahedron Lett., 2455 (1976). A. Saito, H. Matsushita, and H. Kaneko, Chem. Lett., 729 (1984). V. N. Odinokov, O. S. Kukovinets, R. A. Zainullin, E. Yu. Tsyglintseva, V. R. Sultanmuratova, V. V. Veselovskii, V. A. Dragan, T. Ya. Rubinskaya, B. A. Cheskis, A. M. Moiseenkov, and G. A. Tolstikov, Khim. Prir. Soedin., 419 (1989).

635

Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

INFLUENCE OF REDUCED TEMPERATURE ON LIPID COMPOSITION OF Viburnum opulus SEEDS

A. R. Karimova, S. G. Yunusova, M. S. Yunusov, and E. G. Galkin

UDC 547.915:665.33

The influence of reduced temperature (frost) on the lipid composition of Viburnum opulus seeds was studied. Low temperatures changed lipid metabolism by forming hydroxyglycerides in NL, lowering significantly the GL and PL contents, and changing their class composition. The structures of hydroxyacids in the hydroxyglycerides were established. Key words: Viburnum opulus, neutral lipids, glycolipids, phospholipids, lipophilic components, fatty acids, hydroxyacids, frost. Various negative environmental factors such as increased temperature, drought, and freezing can induce stress and disrupt the vital functions of plants in nature. Lipids are one of the principal classes of natural compounds in all higher plants and can undergo various changes [1, 2]. The present work studied the influence of frost on the lipid composition of Viburnum opulus L. because fruits are collected until late autumn. We studied the compositions of neutral and polar glyco- and phospholipids, lipophilic components, and fatty acids. The results were compared for ripe seeds from fruits collected in October before frost (Specimen I) and in November, when the temperature decreased to -10-20°C (Specimen II). Lipids were identified by the usual methods using qualitative reactions, comparison of TLC mobilities with authentic compounds, physicochemical analytical methods, and chemical transformations. The weight of the fruit decreased after frost because of desiccation of the pericarp (Table 1). The oil content of the seeds and the acid number (AN) increased slightly whereas the carotinoid content decreased. Apparently the changes in the oil content of the seeds and the AN of the oil were due to hydrolysis in combination with the influence of cold temperatures on the plant [3-7]. Such trends were noted during a study of the influence of freezing on lipids from leaves of certain coffee varieties [8] and peppers [9]. Neutral lipids (NL) were separated into classes by column chromatography (CC) (Table 2). The content of triterpene alcohols and their esters and of triacylglycerides (TAG) and monoacylglycerides (MAG) decreased significantly in NL of Specimen II. The content of diacylglycerides (DAG) doubled. A new class of compounds appeared and was tentatively identified as hydroxyacylglycerides (HAG) (11%) that were absent in seeds collected before frost. TLC using system 4 of the HAG produced two main spots with Rf values 0.36 and 0.17. This mobility is the same as that of HAG isolated from castor oil. The IR spectrum of these compounds contained vibrations for OH at 3200-3600 cm-1 and ester at 1744 cm-1. The IR spectrum of the product from mild alkaline hydrolysis gave vibrations corresponding to OH. The band characteristic of ester disappeared. Vibrations characteristic of a carboxylic acid carbonyl appeared at 1714 cm-1. TLC using system 4 of the hydrolysis products of methylated HAG produced three bands. The least polar products had the same mobilities as methyl esters of ordinary fatty acids (FAME). The product with Rf 0.53 (I) corresponded with the methyl ester of ricinolic acid. There were also components (II) with Rf 0.35. They were separated using preparative TLC (PTLC) and system 4. The FAME were identified using GC (mass %): 12:0, 0.4; 14:0, 0.2; 15:0, tr.; 16:0, 11.8; 17:0, 0.2; 18:0, tr.; 18:1, 59.3; 18:2, 22.6; 18:3, 1.7; 22:0, 3.8. Institute of Organic Chemistry, Ufa Scientific Center, Russian Academy of Sciences, 450054, Ufa, pr. Oktyabrya, 71, fax (3472) 35 60 66, e-mail: [email protected]. Translated from Khimiya Prirodnykh Soedinenii, No. 6, pp. 523-528, November-December, 2005. Original article submitted July 19, 2005. 636

0009-3130/05/4106-0636 ©2005 Springer Science+Business Media, Inc.

TABLE 1. Properties of Fruit and Seed Oil of Viburnum opulus Property

Specimen I

Specimen II

59.0 8.6 15.4 1.5 4.0

51.0 9.4 16.0 2.5 3.4

Weight of 100 fruits, g Moisture, absolute dry seed mass, % Oil content, absolute dry seed mass, % Acid number, mg KOH Carotinoid content, mg % OT MS

OT MS

O

O

CH 3 (CH2) 4 CH CH (CH 2) 2 CH CH 2 CH CH C (CH2) 3 C 12

9

6

227

5

271

O

9

CH3

OCH3

(CH2) 6 CH CH CH (CH 2) 3 11

5

129

299

CH CH CH

O 12

CH 2 CH 2

17

95

263

SMT O

4

OT MS

SMT O

(CH2) 8 CH CH CH2 (CH2) 5 CH 2 C 10

303

O

9

259

OCH3

259

215

3 SMT O

O

SMT O OT MS O CH 3 CH 2 (CH2) 6 CH CH CH2 (CH2) 5 CH2 C OCH 3 10 9

11

CH2 CH2 (CH 2)8 CH2 C OCH3

13

67

C

2

81 CH2

(CH2) 3 129

1

15

C

10

CH3 CH2 (CH2) 5 CH CH

OCH3

11

201

5 SMT O 11

CH 3 (CH2) 5 CH CH 12

187

OT MS CH (CH 2)7 CH2 C 10

OT MS

OT MS CH (CH 2)6 CH2 C 9

OT MS

259

O OCH3

6

O OCH3

273 7

We used GC—MS analysis to identify the hydroxyacids in the HAG. The hydroxyacids were analyzed as the TMS derivatives of their methyl esters. According to GC—MS, the hydroxyacids were a rather complicated mixture of oxidation products. Owing to the complexity of the mixture, only the main components could be identified. The main compounds in I were 5-oxo-9-hydroxyoctadeca-6,12-dienoic (1), 5-oxo-11-hydroxyoctadec-9-enoic (2), and 15,17-octadecadienoic (3) acids. The mass spectrum of 1 contained principal peaks with m/z 73 (100%), 129 (40), 227 (70), and 271 (35). According to the literature [10], the peaks with m/z 271, 227, and 129 are important for identifying the acid. The spectrum indicated that the compound contained two double bonds, a carbonyl, and an initial C9 hydroxyl. The fact that the peak with m/z 227 was more than two times as strong as that with m/z 271 was consistent with one of the double bonds in the position allylic to the C-OTMS group on C6 and C7. The peak with m/z 271 is due to brake of bond between C9 and C10. Take into account that peak with m/z 271 was weak the second double bond was not located in the position allylic to the initial hydroxyl and was most likely located between C12 and C13. The peak with m/z 129 is most probably due to a carbonyl on C5. Acid 2 was identified analogously from the principal characteristic peaks with m/z 73 (100%), 299 (50), and 129 (25). In addition to the aforementioned acids, specimen I also contained 3, the mass spectrum of which gave peaks with m/z 294 (25%) [M]+, 263 (10) [M - 31]+, 220 (5) [M - 31 - 43]+, and peaks for fragments with m/z 95 (45) and 67 (100) that identified it as 15,17-octadecadienoic acid (the UV spectrum of I had an absorption at 232 nm characteristic of a conjugated double bond). It is not clear why this acid appeared in the hydroxyacid fraction. Degradation products of fatty acids of varying degree of unsaturation were also observed. These were 9-oxononanoic, octanedicarboxylic, nonanedicarboxylic, and decanedicarboxylic acids.

637

TABLE 2. Neutral Lipids of Viburnum opulus Seeds NL mass % NL class Triterpene esters (TE) Fatty acid methyl esters (FAME) Triacylglycerides (TAG) Free fatty acids (FFA) Triterpene alcohols (TA) Sterols Diacylglycerides (DAG) Hydroxyglycerides Monoacylglycerides (MAG)

Specimen I

Specimen II

2.0 0.1 93.1 0.8 0.6 1.1 1.3 1.0

0.7 0.1 83.2 0.9 Tr. 1.0 2.6 10.9 0.6

TABLE 3. Lipophilic Components of Viburnum opulus Seeds Free, %

Bound, %

Triterpene I

II

4,4′-Demethylsterols (sterols)

β-Sitosterol Campesterol Isofucosterol Stigmasterol Cholesterol Stigmastan-3-ol

88.7 4.5 3.2 0.8 0.5 2.3

I

II

76.1 5.6 8.1 5.7 4.5

78.2 8.4 13.4 -

64.9 21.7 11.8 1.6

72.9 21.6 3.5 2.0

a

71.2 7.7 17.3 1.7 2.1 -

4,4′-Dimethylsterols (triterpene alcohols)b

α-Amyrin β-Amyrin 24-Methylenecycloartanol Lupeol

71.4 15.0 11.7 1.9

-

______ Specimen I was fruits collected before frost; specimen II, after frost. a% of total sterol mass; b % of total triterpene alcohol mass. The principal components of II were saturated di- and trihydroxyacids: 9,10-dihydroxyoctadecanoic (4), 9,10,18trihydroxyoctadecanoic (5), 9,10,11-trihydroxyoctadecanoic (6), and 10,11,12-trihydroxyoctadecanoic (7) acids. The mass spectrum of 4 had principal peaks with m/z 73 (100%), 215 (30), and 259 (30). The identical strengths of the peaks with m/z 215 and 259 indicated that the compound had two OH groups in the 9- and 10-positions. Peaks for 5 with m/z 259 (70%) and 303 (30) can be explained based on the structure of trihydroxydecanoic acid and location of the initial hydroxyls in the 9-, 10-, and 18-positions. The location of the hydroxyl in the terminal position on C18 explains why it is not involved in the fragmentation. The mass spectra of 6 and 7 are consistent with two isomers involving the hydroxyls. The spectra of these compounds gave peaks for fragments with m/z 201 (45%) and 259 (30) for 6 and 187 (50) and 273 (30) for 7. These indicated that the structures were 9,10,11- and 10,11,12-trihydroxyoctadecanoic acids. One of the most typical properties of unsaturated fatty acids, especially polyenoic (PUFA), is their suscepibility to peroxide oxidation. Auto-oxidation is considered the most characteristic for animal tissues [11, 12] whereas enzymatic oxidation, for example, lipoxygenase, is dominant in plants. The compounds formed by these reactions are called oxylipins [11, 12]. The 18:2 and 18:3 acids are the main source of their formation in plants. Oxylipins include hydroperoxides, hydroxides, epoxy- and keto-derivatives of fatty acids, volatile aldehydes, and cyclic compounds. A particular enzyme carries out the oxidation at each stage [11, 12].

638

TABLE 4. Neutral Lipids of Viburnum opulus Seeds Content, % Class Specimen I

Specimen II

Phospholipids N-Acylphospholipids (N-acyl-PL) Cardiolipin (CL) Phosphatidylglycerine (PG) Phosphatidylethanolamine (PE) Phosphatidylcholine (PC) Lysophosphatidylethanolamine (lyso-PE) Lysophosphatidylcholine (lyso-PC) Unidentified component Phosphatidylinosite (PI) Phosphatidic acid (PA)

0.14 4.6 3.9 2.0 21.2 30.3 2.7 1.3 10.6 19.0 4.4

0.7×10-3 43.4 12.1 33.7 4.8 3.6 Tr. Tr. Tr. 2.4

Glycolipids Acylmonogalactosyldiacylglycerides (Ac-MGDG) Monogalactosyldiacylglycerides (MGDG) Digalactosyldiacylglycerides (DGDG) Sulfoquinovosyldiacylglycerides (SQDG)

1.1 21.4 40.6 25.0 13.0

0.13 18.6 30.3 38.9 12.2

The qualitative set of free and bound sterols in Specimen II became simpler after frost (Table 3), especially in the bound state. β-Sitosterol remained the main component but its mass in the free state decreased. The content of isofucosterol increased by almost six times; of stigmasterol, by two times. Stigmasterol was not found in the bound state. The same tendency was observed for isofucosterol. The ratio (isofucosterol+stigmasterol)/β-sitosterol in the free state increased by six times compared with that in seeds collected before frost. Isofucosterol, which is an isomer of stigmasterol with respect to the position of the double bond in the side chain, may play the same role as the latter and to a certain extent influences the packing and elasticity of the membrane bilayers. This has been noted in studies of pepper and tomato lipids [5, 9]. The composition of the free triterpene alcohols (TTA) of seeds collected after frost could not be established because of the small quantity available. The qualitative composition of the bound TTA did not change upon cooling. The main component of Specimen II remained α-amyrin. However, the quantity of it increased as did that of lupeol. The content of 24-methylenecycloartanol decreased significantly. The significant decrease in the content of GL and PL, the mass of which decreased by 10 and 1000 times, respectively (Table 4), confirmed that acylhydrolase enzymes [13] were activated upon cooling and membrane lipids were the first ones affected. The PL were almost completely destroyed. This same phenomenon has been noted several times previously [3, 6, 9, 14, 15]. The increased decomposition of total PL did not cause FFA to accumulate but simply kept their level stable by stimulating oxidative processes [4] that destroyed the plant cells. Table 4 shows the changes in the class composition of membrane lipids. The content of classes with monosaccharides decreased in GL of Specimen II: Ac-MGDG, MGDG, and SQDG. The MGDG content decreased by 10% whereas that of glycolipids with disaccharides (DGDG) increased by 14%. It is thought [1, 8] that the DGDG/MGDG ratio and the resistance of plants to frost are definitely related. The content of PC, PE, and PI decreased (by 10, 4, and 19 times, respectively) in PL of Specimen II (Table 4). The amounts of so-called "high melting" PG and di-PG (cardiolipin) increased by 16 and 3 times, respectively. The amount of PA was halved. The FA composition of acyl-containing NL classes in seeds collected after frost changed little compared with those collected before it (Table 5). The qualitative set of acids did not change. The content of saturated FA increased in all classes except DAG because of the 16:0 and 22:0 acids. The content of oleic acid in TE, FAME, and TAG decreased. In these same classes except for TE, the linoleic acid content increased slightly.

639

TABLE 5. Fatty Acid Classes of NL from Viburnum opulus Seeds

NL

Acid 12:0 14:0 15:0 16:0 17:0 18:0 18:1 18:2 18:3 22:0 Σsat.FA Σunsat.FA

TE

FAME

TAG

FFA

DAG

MAG

I

II

I

II

I

II

I

II

I

II

I

II

I

II

Tr. Tr. Tr. 2.3 Tr. Tr. 59.9 37.8 Tr. Tr. 2.3 97.7

0.5 0.3 Tr. 2.9 Tr. Tr. 52.1 42.6 Tr. 1.6 5.3 94.7

Tr. 2.2 Tr. 22.3 Tr. Tr. 37.6 29.0 2.0 6.9 31.4 68.6

1.0 2.4 Tr. 35.7 Tr. 26.6 30.3 4.0 Tr. 39.1 60.9

Tr. 0.4 7.8 59.7 29.9 Tr. 2.2 10.4 89.6

Tr. 0.4 2.2 31.5 47.4 Tr. 18.5 21.1 78.9

Tr. Tr. Tr. 2.3 0.2 58.8 35.7 1.3 1.7 4.2 95.58

Tr. Tr. 3.1 53.8 39.7 3.4 6.5 93.5

Tr. 3.4 8.6 0.7 57.8 29.5 12.7 87.3

Tr. 0.2 0.2 10.4 60.6 22.2 0.8 5.6 16.4 83.6

Tr. Tr. 14.2 Tr. 45.1 38.1 1.3 1.3 15.5 84.5

0.8 0.4 0.1 4.8 0.2 54.6 38.2 0.9 7.2 92.8

0.5 0.8 Tr. 9.3 2.1 Tr. 53.2 32.8 Tr. 1.3 14.0 86.0

Tr. Tr. 7.4 0.4 Tr. 51.2 28.6 12.4 20.2 79.8

______ Specimen I was fruits collected before frost; Specimen II, after frost.

TABLE 6. Total Fatty Acids of GL and PL from Viburnum opulus Seeds

GL

Acid 12:0 14:0 15:0 16:0 16:1 17:0 18:0 18:1 18:2 18:3 20:0 22:0 Σsat.FA Σunsat.FA

PL

Specimen I

Specimen II*

Specimen I

Specimen II

Tr. 0.8 0.2 10.8 Tr. 0.9 Tr. 34.3 38.6 4.5 9.9 22.6 77.4

0.4 0.7 Tr. 20.4 Tr. Tr. 22.3 35.2 14.7 4.7 1.6 45.4 54.6

0.2 0.7 25.8 0.5 Tr. 14.1 53.9 1.5 3.3 30.5 69.5

Tr. Tr. 26.6 Tr. 36.4 18.9 8.9 6.6 2.6 65.6 34.4

______ *9-Hydroxyoctadecanoic acid was present in addition to the indicated fatty acids in Specimen II.

Significant changes in the contents of saturated and unsaturated FA were observed in the GL and PL (Table 6). The overall level of saturated FA in Specimen II doubled in both GL and PL. The 16:0 content in GL rose by 2 times; that of 18:0, by 22 times (i.e., from traces to 22%). The mass of 16:0 in PL did not change whereas that of 18:0 increased from traces to 36%. The content of oleic acid was practically constant. The content of linoleic in GL decreased by more than 2 times; in PL, by 6 times. The amount of linolenic (18:3) acid in PL rose by four times. We found that lipids of these seeds show a decreased content of PC and PE, an accumulation of PG, a significant increase of saturated (16:0 and 18:0) fatty acids in both PL and GL, and the formation of oxylipins. This indicates that low temperatures change the lipid metabolism and eventually destroy membranes.

640

EXPERIMENTAL IR spectra were recorded on UR-20 and Specord M80 spectrometers. Spectrophotometric data were obtained on a Specord M400 instrument; mass spectra, in an MX-1320 spectrometer; GC—MS, using a computerized HP 5890 GC—MS with an HP 5972A mass-selective detector. The analytical conditions have been published [16]. Mass spectra were interpreted using a library based on spectrum—structure correlations and trends and fragmentation features of triterpenes from the literature [1719]. GC was performed on a Chrom-5/DIP chromatograph with a column (1.2 × 3 mm) packed with PDEGS (5%) on Chromaton N-AW-DMCS at 160°C with He carrier gas and flow rate 75 mL/min. CC of total lipids was carried out on L 100160 µm silica gel (Chemapol, Czechoslovakia) at a 1:50 (by weight) extract:adsorbent ratio and a 1:20 cross-section:height aspect ratio. Analytical TLC used Silufol plates and glass plates with a deposited layer of LSL 5/40 µm silica gel with gypsum (13%, Chemapol, Czechoslovakia). PL were identified on standard plates (6 × 6 cm) with KSKG silica gel (5-20 µm fraction, layer thickness 110-150 µm (Lyaene Kalur, Haapsalu). PTLC was carried out on glass plates with LSL 5/40 µm silica gel with gypsum (13%, Chemapol, Czechoslovakia). Lipids were separated, purified, and identified using the following solvent systems: for NL: petroleum ether:diethylether (9:1, 1; 7:3, 2; 1:1, 3; 2:3, 4); for polar lipids: chloroform:methanol:ammonia (25%) (65:25:4, 5); for GL: chloroform:acetone:methanol:acetic acid:water (65:20:10:10:3, 6), acetone:toluene:acetic acid:water (60:60:20:1, 7); for PL: chloroform:methanol:ammonia (25%) (13:7:1, 8), chloroform:acetone:methanol:acetic acid:water (6:8:2:2:1, 9), chloroform:methanol:ammonia (25%) (10:5:2, 10a), chloroform:methanol:acetic acid:water (14:5:1:1, 10b), chloroform:methanol:benzene:ammonia (25%) (13:6:2:1.2, 11a), and chloroform:methanol:benzene:acetone:acetic acid (14:6:2:1:0.8, 11b). The 40-70°C petroleum ether fraction was used. Lipids were detected using I2 vapor for NL, H2SO4 (50% solution) with subsequent heating of the plate until colored spots appeared for triterpenes; α-naphthol and H2SO4 (50%) for GL, and Vaskovsky, Dragendorff, and ninhydrin solutions for PL [20]. Fruits of V. opulus were collected in Iglin region of Bashkortostan from thickets of bushes in the floodplain of Lobovki river. The oil content of seeds and their moisture content [21], acid number [22], and carotinoid content [23] were determined by standard methods. Lipids (NL and polar) were isolated from freshly collected fruits by separating seeds from pulp, grinding them in an electric grinder until homogeneous, and extracting exhaustively with chloroform:methanol (2:1) by standing at room temperature. The combined extracts were reduced to half the volume in a rotary evaporator, treated with NaCl solution (1%), dried over Na2SO4, and evaporated to dryness. Lipids were separated into NL and polar lipids using CC. The eluent was chloroform with subsequent addition of methanol from 0 to 100%. NL elution was monitored by TLC on Silufol using solvent systems 1-4; polar lipids, by TLC on silica gel using solvent systems 5-9. GL and PL were separated initially using precipitation of the latter with acetone [32]. The resulting PL were separated from traces of GL by PTLC on plates (20 × 20 cm) using acetone dried three times over K2CO3. The amounts of NL and GL were estimated gravimetrically; of PL, by spectrophotometry [24]. Total NL were separated into classes using CC with elution by petroleum ether:diethylether (0-100%). Fractions were purified as necessary by PTLC on plates (20 × 20 cm) using solvent systems 1-4. GL were separated into classes also using CC with elution by chloroform:methanol (0-100%). Fractions were purified as necessary by PTLC on plates (20 × 10 cm) using solvent systems 5-7. They were desorbed from the silica gel using chloroform:methanol (2:1). The PL classes were determined qualitatively and quantitatively as before [24]. PL were identified using onedimensional chromatography with authentic compounds and solvent systems 5, 8, and 9 in addition to two-dimensional chromatography and solvent systems 10 and 11 with (a) in the first direction and (b) in the second. Alkaline hydrolysis of acyl-containing lipid fractions was carried out according to the literature [25]. Triterpene esters underwent total alkaline hydrolysis as previously described [16]. Fatty acids were esterified using diazomethane solution in diethylether. Hydroxyacid methyl esters were silylated as before [20].

641

REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25.

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P. J. C. Kuiper, Physiol. Plant., 64, 118 (1985). J. L. Harwood, Prog. Lipid Res., 33, No. 1-2, 193 (1994). Z. Kaniuga, V. Saczynska, E. Miskiewicz, and M. Garstka, Acta Physiol. Plant., 21, No. 1, 45 (1999). Z. Kaniuga, V. Saczynska, E. Miskiewicz, and M. Garstka, Acta Physiol. Plant., 21, No. 3, 231 (1999). B. D. Whitaker, Phytochemistry, 30, No. 3, 757 (1991). H.-L. Yu and C. Willemot, Plant Sci., 113, 33 (1996). H.-L. Yu and C. Willemot, Plant Sci., 125, 21 (1997). P. S. Campos, V. Quartin, J. C. Ramalho, and M. A. Nunes, J. Plant Physiol., 160, 283 (2003). B. D. Whitaker, Physiol. Plant., 93, 683 (1995). R. Kleiman and G. F. Spencer, J. Am. Oil Chem. Soc., 50, 31 (1973). E. Blee, Prog. Lipid Res., 37, No. 1, 33 (1998). H. Zhuang, M. M . Barth, and D. Hildebrand, in: Food Lipids: Chemistry, Nutrition, and Biotechnology [In: Food Sci. Technol. (New York, NY, USA) 2002; 117], C. C. Akoh and D. B. Min, eds., Marcel Dekker, Inc., New York (2002), p. 413. T. Galliard, in: The Biochemistry of Plants, P. K. Stumpf, ed., Academic Press, New York (1980), Vol. 4, p. 85. R. Welti, W. Li, M. Li, Y. Sang, H. Biesiada, H.-E. Zhou, C. B. Rajashekar, T. D. Williams, and X. Wang, J. Biol. Chem., 277, No. 35, 31994 (2002). K. L. Parkin and S.-J. Kuo, Plant Physiol., 90, 1049 (1989). A. R. Karimova, S. G. Yunusova, S. I. Maslennikov, E. G. Galkin, T. S. Yunusov, V. V. Shereshovets, and M. S. Yunusov, Khim. Prir. Soedin., 447 (2000). E. Hettman, Modern Methods of Steroid Analysis, Academic Press, New York (1973), p. 138. N. S. Vul′fson and V. G. Zaikin, Usp. Khim., 42, Vyp. 8, 1379 (1973). L. S. Golovkina, G. V. Rusinova, and A. A. Petrov, Usp. Khim., 53, Vyp. 9, 1493 (1984). M. Kates, Techniques of Lipidology: Isolation, Analysis, and Identification of Lipids, Elsevier, New York (1973). Handbook of Methods for Research and Chemical Monitoring and Accounting of Production in the Oil Production and Fat Processing Industry [in Russian], VNIIZh, Leningrad (1965), Vol. 2, pp. 152, 146. Chemical Monitoring and Accounting of Production in the Oil Production and Fat Processing Industry [in Russian], Pishchpromizdat, Moscow (1958), Vol. 1, p. 306. R. S. Limar′ and O. V. Sakharova, Methods for Comprehensive Study of Photosynthesis [in Russian], (1973), Vol. 2, p. 260. V. E. Vaskovsky, E. Y. Kostetsky, and I. M. Vasendin, J. Chromatogr., 114, No. 1, 129 (1975). A. R. Karimova, S. G. Yunusova, E. G. Galkin, N. I. Fedorov, and M. S. Yunusov, Izv. Akad. Nauk, Ser. Khim., No. 1, 235 (2004).

Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

OZONOLYSIS OF RICINOLIC ACID DERIVATIVES AND TRANSFORMATIONS OF THE OZONOLYSIS PRODUCTS UNDER BARTON REACTION CONDITIONS

G. Yu. Ishmuratov, R. Ya. Kharisov, A. Kh. Shayakhmetova, L. P. Botsman, O. V. Shitikova, and G. A. Tolstikov

UDC 542.85+542.943.5+665.335.5

The possibility of functionalizing the alkyl part of ricinolic acid using the Barton reaction was investigated. Key words: castor oil, ozonolysis, Barton reaction, ricinolic acid. Ricinolic acid (1) is a promising substrate for preparing chiral polyfunctional compounds that can in turn act as convenient building blocks for the synthesis of organic compounds with more complicated structures because of its availability from castor oil and the presence of asymmetric C-12 with the R-configuration. OH (CH2)7CO2H 1

ONO

OR ONO 2, 3 2: R = Ac; 3: R = CH2Ph

4

Herein we report results of investigations on expanding the synthetic potential of 1 by further functionalization of the chemically stable alkyl part of the molecule (C-12—C-18) using the Barton photochemical rearrangement of nitrites 2-4 in the key step. In order to simplify the structure of starting 1, it was fragmented by ozonolysis to produce the desired secondary monoacetate of diol 6. Compound 6 was prepared previously by ozonolysis of methyl R-12-acetoxyoctadec-9-enoic acid (5) in MeOH [1]. Under the conditions described for reduction of the intermediate peroxides (NaBH4—MeOH) in the reaction mixture, the product of thermodynamic control, primary acetate 7 (ratio 6:7, 1:2.5), dominated and was formed as a result of intramolecular trans-esterification. Furthermore, an equimolar amount of methyl 9-hydroxynonanoic acid (8), which is difficult to separate from monoesters 6 and 7, was necessarily formed as a side product. We investigated ways of optimizing the ozonolysis of ricinolic acid to synthesize secondary acetate 6. The first approach was based on transformation of the carboxylic acid in 1 to a methyl, for which we used successive hydride reduction of castor oil 9 (ricinolic acid content ~85%) to diol 10 and subsequent deoxygenation of the primary hydroxyl in it through monotosylate 11. The resulting chiral homoallyl alcohol 12 was converted to acetate 13 or benzyl ether 14. Ozonolytic cleavage of the double bond in 13, carried out in CH2Cl2 in the presence of MeOH (2 equiv.) and subsequent treatment of the peroxide products with NaBH4, gave a mixture (9:5:1) of acetates 6 and 7 and diol 15 with the desired 6 predominating. These were readily freed from the necessary ozonolysis—reduction product 1-nonanol (16) by vacuum distillation. Replacing NaBH4 by KBH4 increased the content of 6 (ratio 6:7:15, 6:1.1:1). The best results (ratio 6:7:15, 8.6:1.6:1) were obtained for ozonolysis in CH2Cl2 in the presence of two equivalents of AcOH per double bond and with the mild reducing agent NaBH(OAc)3. Institute of Organic Chemistry, Ufa Scientific Center, Russian Academy of Sciences, 450054, Ufa, pr. Oktyabrya, 71, fax (3472) 35 60 66, e-mail: [email protected]. Translated from Khimiya Prirodnykh Soedinenii, No. 6, pp. 529-534, NovemberDecember, 2005. Original article submitted April 28, 2005. 0009-3130/05/4106-0643 ©2005 Springer Science+Business Media, Inc.

643

OAc

OH

OH

+

OH

OAc

6

+

CH3(CH 2)5

7

OH

+

R3O

CH 2

R3O

CH CH 2

R3O

15

18

R2 O

CH 2

R2O

CH

R2 O

CH 2

R1O

1. Ac2O

CH2

R1O

CH

R1O

CH2

17

OH

1

1. Bu2AlH

OH

LiAlH4

CH3(CH2) 5

(CH2) 8OT s

2. TsCl, Et3N

CH3(CH 2)5

11

9

BnBr NaH

Ac2O ∆

OAc CH 3(CH 2) 5

(CH2) 7CH3

12

OBn CH3(CH 2)5

(CH2) 7CH 3

(CH2) 7CH3

14

13 OBn

6 + 7 + 15 +

CH3(CH2) 8OH 16

CH 3(CH2) 5

OH

+

CH 3(CH 2) 8OH

19

16

OH 20 OH

9: R1 = CH3(CH2)5

OAc

O

17: R2 =

(CH2)7C

CH3(CH2)5

O

O (CH2)7C

18: R3 =

HO(CH2)8C

In order to extend the results to castor oil, we ozonolyzed 17, which furthermore enabled the side product of highly polar triglyceride 18 to be easily removed. The highest yield of 6 (ratio 6:7:15, 17.7:1.5:1) was obtained, like for 13, for ozonolysis in CH2Cl2 in the presence of two equivalents of AcOH per one double bond and the use of NaBH(OAc)3 as reductant. Benzyl ether 14 was ozonolyzed without any complications to give the other key hydroxyether 19 in high yield. The precursor of alkylnitrate 4, secondary alcohol 20, was prepared from diol 15 by deoxygenation of the corresponding primary tosylate. The Barton reaction of nitrites 2-4, which were prepared from alcohols 6, 19, and 20 by the standard method [2], was carried out in an inert atmosphere and an aprotonic medium (benzene). Photolysis of nitrites 2 and 21, synthesized from a mixture (14:1) of hydroxyacetates 6 and 7, produced the starting 6 and 7 in addition to diol 15, which were isolated from the reaction mixture and identified. 1-Acetoxy-3-nonanone (22), the product of disproportionation or a "cage radical" reaction between the alkoxyradical and NO [3] that were formed from nitrite 21, was also observed. O

ONO

6+7

NaNO2 H2SO4

2 +

CH3(CH2)5

hv OAc

6 + 7 + 15 + CH3(CH2)5

21

OAc 22

Because the secondary acetate in 6 was labile, we used the corresponding benzyl derivative 19. Photolysis of its primary nitrite 3 led to an unusual result. The usual product [4] of 1,5-cleavage was missing among the isolated and identified reaction products.

19 H SO 2 4

OBn

OBn

NaNO2

hv CH3(CH2)5

ONO 3

15 + 19 +

CH3(CH2)5

O

+ O

23 CH3(CH2)5

O 24

The principal product was 2-phenyl-4R-hexyl-1,3-dioxane (24) resulting from 1,6-cleavage in addition to impurities of 19, 15, and aldehyde 23. This was confirmed by spectral data. The PMR spectrum had signals at chemical shifts for a phenyl

644

ring (δ 7.25-7.55 ppm) and a singlet for an acetal proton (δ 5.52 ppm). The 13C NMR spectrum had a doublet (δ 101.09 ppm) for C-2 and other characteristic signals. Conversely, photolysis of nitrite 4, prepared from optically pure secondary alcohol 20, followed by chromatography of the reaction mixture isolated a compound with a modified alkyl chain of the starting material, 6-nitroso-3S-nonanol, existing primarily [5] as the more stable dimer 26, and 20 and 25 [3]. OH OH

20

O

ONO NaNO2 H2SO4

hv CH3(CH2)5

N

20 + CH3(CH2)5

+

O

O

N

25

4

OH ∆

26

OH

N OH

27

Thermolysis of 26 gave quantitatively the -hydroxyoxime 27 as an equal mixture of the syn- and anti-isomers. Twodimensional CH CORR and double resonance were used in order to assign unambiguously signals in the PMR and 13C NMR spectra of the stereoisomers of 27. The signals for C-5 [23.46 (syn-), 29.76 (anti-)] and C-7 [35.88 (syn-), 30.45 (anti-)] and their protons H-7 [2.35 (anti-), 2.15 (syn-)] and Ha-5 [2.40 (syn-), 2.70 (anti-)] were nonequivalent because of different shielding of the corresponding atoms in this pair of compounds and agreed well with the literature [6] (see Experimental).

EXPERIMENTAL IR spectra were recorded on a Specord M-82 instrument as thin layers. NMR spectra were obtained on a Bruker AMX300 spectrometer (working frequency 300.13 MHz for 1H and 75.47 MHz for 13C) in CDCl3 with an internal standard of the CDCl3 signals (1H δ 7.27 ppm, 13C δ 77.00 ppm). Chromatography was carried out in a Chrom-5 instrument [column length 1.2 m, stationary phase silicone SE-30 (5%) on Chromaton N-AW-DMCS (1.16-0.20 mm), working temperature 50-300°C, He carrier gas] and in a Shimadzu GC-9A chromatograph (quartz capillary column, 25 m, stationary phase OV-101, working temperature 80-260°C). Optical rotation was measured in a Perkin—Elmer 241 MC polarimeter. Photolysis was performed using a mercury lamp OKN-14, TU 64-1-1618-77 (220 V, 950 VA, 1000 W, 50 Hz). The Barton reaction was performed in a thermostatted Pyrex glass cell ( > 320 nm, 80 mL). TLC monitoring used Sorbfil SiO2 (Russia). Column chromatography was carried out over SiO2 (70-230, Lancaster, England) with elution by petroleum ether (40-70°C). The ozonator production was 38 mmol O3/h. Elemental analyses agreed with those calculated. Castor Oil Acetate (17). Castor oil (9, 20.00 g) and Ac2O (40.29 g, 394.9 mmol) were refluxed for 5 h, washed with hot water (3 × 100 mL), diluted with CH2Cl2 (300 mL), dried over Na2SO4, and evaporated to afford 17 (22.24 g, 98%). IR spectrum (ν, cm-1): 3016 (C=C), 1740 (C=O), 1240, 1164 (C–O–C), 736 (C=C). R-Octadec-9Z-en-1,12-diol (10). A solution of 9 (10.9 g) in absolute THF (270 mL) and Et2O (150 mL) (Ar, -10°C) was treated dropwise with a solution of i-Bu2AlH (23.0 mL, 102.4 mmol, 73%). After the entire amount of i-Bu2AlH was added, the reaction mixture was stirred for 1.5 h at 0°C, treated dropwise with H2O (22 mL, 0-5°C), warmed to room temperature, and stirred for another 2 h. The resulting solid was filtered off and washed with Et2O. The filtrate was dried over Na2SO4 and evaporated to afford the product (8.74 g), column chromatography of which (SiO2, PE:CH2Cl2, 2:1) isolated 10 (7.70 g, 94%), [α]D18 +2.24° (c 0.03, CHCl3). IR spectrum (ν, cm-1): 3304 (OH), 3010, 1660, (C=C), 1110, 1054, (C–O), 724 (C=C). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.89 (3H, t, J = 6.8, CH3), 1.20-1.45 (18H, m, 9CH2), 1.49 (2H, m, H-13), 1.53 (2H, m, H-2), 1.64 (2H, br.s, 2OH), 2.04 (2H, q, J = 6.7, H-8), 2.22 (2H, t, J = 7.2, H-11), 3.54 (1H, quintet, J = 5.8, H-12), 3.63 (2H, t, J = 6.6, H-1), 5.40 (1H, dt, J = 10.8, 7.4, H-10), 5.55 (1H, dt, J = 10.8, 7.1, H-9).

645

R-Octadec-9Z-en-7-ol (12). A solution of 10 (8.80 g, 30.9 mmol) in dry Et3N (7 mL, 128.5 mmol) was stirred (Ar, 0°C), treated in portions over 1 h with TsCl (7.05 g, 36.7 mmol), left overnight in a refrigerator, diluted with t-BuOMe (200 mL), washed successively with H2O, aqueous HCl (10%), and saturated NaHCO3 and NaCl solutions, and dried over Na2SO4 to give 11 (13.56 g), which was used without further purification. IR spectrum (ν, cm-1): 3346 (OH), 3010, 1650 (C=C), 1605 (Ar), 1378, 1185, (S=O), 1110, 1048, (C–O), 724 (C=C). A solution of 11 (13.56 g, 30.9 mmol) in absolute THF (80 mL) and absolute Et2O (80 mL) (Ar, 0°C) was treated with LiAlH4 (2.65 g, 71.0 mmol), stirred for 3 h at room temperature, cooled to 0(C, diluted with H2O (3.3 mL), warmed to room temperature, and stirred for another 3 h. The resulting solid was filtered off and washed with t-BuOMe. The filtrate was dried over Na 2SO4 and evaporated to give a product (7.02 g), column chromatography of which (SiO2, PE:CH2Cl2, 2:1, Rf 0.3) isolated 12 (4.12 g, 50%), [α]D20 +2.6° (c 0.03, CH2Cl2). IR spectrum (ν, cm-1): 3346 (OH), 3010, 1650 (C=C), 1110, 1048 (C–O), 724 (C=C). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.88 (6H, t, J = 6.5, 2CH3), 1.20-1.45 (20H, m, 10CH2), 1.47-1.50 (2H, m, H-6), 1.58 (1H, br.s, OH), 2.05 (2H, q, J = 6.8, H-11), 2.22 (2H, t, J = 6.6, H-8), 3.62 (1H, quintet, J = 5.9, H-7), 5.35-5.65 (2H, m, H-9, H-10). 13C NMR spectrum (CDCl , δ, ppm): 14.01 (both q, C-1, C-18), 22.60 (both t, C-2, C-17), 25.68 (t, C-5), 27.40 (t, 3 C-11), 29.27, 29.30, 29.34 (C-12, C-13, C-14), 29.47 (t, C-4), 29.66 (t, C-15), 31.82 (both t, C-3, C-16), 35.32 (t, C-8), 36.82 (t, C-6), 71.47 (d, C-7), 125.10 (d, C-9), 133.37 (d, C-10). R-7-Acetoxyoctadec-9Z-ene (13). A mixture of 12 (2.50 g, 9.3 mmol) and Ac2O (20.13 mL, 57.0 mmol) was refluxed for 2 h, washed with hot H2O (3 × 20 mL), diluted with CH2Cl2, dried over Na2SO4, and evaporated to afford 13 (2.87 g, 99%), [α]D18 +25.2° (c 0.02, CHCl3). IR spectrum (ν, cm-1): 1738 (C=O), 3010, 1654 (C=C), 1110, 1048 (C–O), 724 (C=C). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.86 (6H, t, J = 6.7, 2CH3), 1.20-1.45 (20H, m, 10CH2), 1.50-1.62 (2H, m, H-6), 2.00-2.10 (4H, m, H-11, AcO), 2.32 (2H, dd, J = 6.7, J = 6.2, H-8), 4.88 (1H, quintet, J = 6.2, H-7), 5.35 (1H, dt, J = 10.8, J = 7.2, H-9), 5.48 (1H, dt, J = 10.8, J = 7.2, H-10). R-7-Benzyloxyoctadec-9Z-ene (14). A suspension of NaH (0.56 g, 14.6 mmol, 60% in mineral oil) in DMF (5 mL) was treated dropwise (Ar, 0°C) with 12 (3.5 g, 13.0 mmol) in DMF (5.5 mL), stirred at 0°C for 1 h, treated with BnBr (2.3 mL, 18.3 mmol), stirred for 6 h at room temperature, left overnight, diluted with t-BuOMe (200 mL), washed with water and saturated NaCl solution, dried over Na2SO4, and evaporated. The solid (5.23 g) was chromatographed (SiO2, PE) to afford 14 (4.06 g, 87%), [α]D20 +18.1° (c 0.02, CH2Cl2). IR spectrum (ν, cm-1): 3010, 1662 (C=C), 1625 (Ar), 1110, 1048 (C–O), 755 (C=C). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.90 (6H, t, J = 6.7, 2CH3), 1.20-1.55 (20H, m, 10CH2), 1.50-1.62 (2H, m, H-6), 2.06 (2H, q, J = 6.3, H-11), 2.25-2.45 (2H, m, H-8), 3.43 (1H, quintet, J = 5.8, H-7), 4.51 (1H, d, J = 11.7, PhCHa), 4.60 (1H, d, J = 11.7, PhCHb), 5.35-5.60 (2H, m, H-9, H-10), 7.25-7.40 (5H, m, Ph). Ozonolysis and reduction of peroxide ozonolysis products: a) An ozone—oxygen mixture was bubbled through a solution of 13 (1.80 g, 8.8 mmol) in CH2Cl2 (18 mL) and absolute MeOH (0.7 mL, 17.6 mmol) at 0°C at an O3:13 mole ratio of 1:1. The reaction mixture was purged with Ar, diluted with CH2Cl2 (20 mL), stirred (10°C), treated with NaBH4 (0.44 g, 11.6 mmol), stirred at room temperature for 3 h, treated with AcOH (0.14 mL) and H2O (1.4 mL) (10°C), stirred at room temperature for 0.5 h, and washed with saturated NaCl solution. The organic layer was dried over Na2SO4 and evaporated to give a product (0.95 g) that contained 6, 7, and 15 in a ratio of 9:5:1, respectively, according to GC. b) The peroxide ozonolysis product obtained by method a) was reduced with KBH4 (0.63 g, 11.6 mmol), stirred at room temperature for 3 h, treated with AcOH (0.14 mL) and H2O (1.4 mL) (10°C), stirred at room temperature for 0.5 h, and washed with saturated NaCl solution. The organic layer was dried over Na2SO4 and evaporated to give a product (0.90 g) that contained 6, 7, and 15 in a ratio of 6:1.1:1 according to GC. c) An ozone—oxygen mixture was bubbled through a solution of 13 (0.90 g, 4.4 mmol) in CH2Cl2 (9 mL) and glacial AcOH (0.35 mL, 8.8 mmol) at 0°C at an O3:13 mole ratio of 1:1. The reaction mixture was purged with Ar, diluted with CH2Cl2 (20 mL), stirred (10°C), treated wth a previously prepared suspension of NaBH(OAc)3 [addition of glacial AcOH (3.4 mL, 60.5 mmol) in CH2Cl2 (6 mL) to a suspension of NaBH4 (0.76 g, 20.17 mmol) in CH2Cl2 (30 mL) with stirring for 2 h], warmed to room temperature, stirred for 3 h, cooled to 10°C, treated with AcOH (0.24 mL) and H2O (2.4 mL), stirred at room temperature for 0.5 h, and washed with saturated NaCl solution. The organic layer was dried over Na2SO4 and evaporated to give a product (1.03 g) that contained 6, 7, and 15 in a 8.6:1.6:1 ratio according to GC.

646

d) An ozone—oxygen mixture was bubled through a solution of 17 (5.0 g, 4.7 mmol) in CH2Cl2 (50 mL) and MeOH (1.1 mL, 28.3 mmol) at 0°C at an O3:17 mole ratio of 3:1. The reaction mixture was purged with Ar, diluted with CH2Cl2 (30 mL), stirred (10°C), treated with NaBH4 (1.4 g, 37.2 mmol), stirred at room temperature for 3 h, treated with AcOH (0.44 mL) and H2O (4.4 mL) (10°C), stirred at room temperature for 0.5 h, and washed with saturated NaCl solution. The organic layer was dried over Na2SO4 and evaporated to give a product (5.46 g) that contained 6 and 7 in a 1.6:1 ratio according to GC. e) The peroxide ozonolysis product obtained by method d) was reduced with KBH4 (2.0 g, 37.2 mmol), stirred at room temperature for 3 h, treated with AcOH (0.44 mL) and H2O (4.4 mL) (10°C), stirred at room temperature for 0.5 h, and washed with saturated NaCl solution. The organic layer was dried over Na2SO4 and evaporated to give a product (4.9 g) that contained 6, 7, and 15 in a 8.7:1.6:1 ratio according to GC. f) An ozone—oxygen mixture was bubbled through a solution of 17 (5.00 g, 4.7 mmol) in CH2Cl2 (50 mL) and glacial AcOH (1.6 mL, 28.3 mmol) at 0°C at an O3:17 mole ratio of 3:1. The reaction mixture was purged with Ar, diluted with CH2Cl2 (20 mL), stirred (10°C), treated with a previously prepared suspension of NaBH(OAc)3 [glacial AcOH (11.2 mL, 194.7 mmol) and NaBH4 (2.46 g, 64.9 mmol) in CH2Cl2 (120 mL)], warmed to room temperature, stired for 3 h, cooled to 10°C, treated with glacial AcOH (0.86 mL) and H2O (8.6 mL), stirred at room temperature for 0.5 h, and washed with saturated NaCl solution. The organic layer was dried over Na2SO4 and evaporated to give a product (4.97 g) that contained 6, 7, and 15 in a 17.7:1.5:1 ratio according to GC. Column chromatography isolated a mixture of 6 and 7 (1.14 g) in a ratio of 14:1, respectively. The IR and NMR spectra of 6, 7, and 15 were practically identical to those in the literature [1]. 3R-Benzyloxynonan-1-ol (19). Ozonolysis of 14 (4.00 g, 11.2 mmol) by method a) followed by vacuum distillation at 1 mm Hg and 100°C produced 19 (2.66 g, 95%), [α]D20 -44.07° (c 0.003, CHCl3). IR spectrum (ν, cm-1): 3420 (OH), 1660 (Ar), 1110, 1048 (C–O). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.91 (3H, t, J = 7.0, CH3), 1.20-1.40 (6H, m, 3CH2), 1.48-1.60 (2H, m, H-5), 1.60-1.73 (2H, m, H-4), 1.70-1.80 (2H, m, H-2), 2.50 (1H, br.s, OH), 3.60-3.65 (1H, m, H-3), 3.65-3.85 (2H, m, H-1), 4.48 (1H, d, J = 11.5, PhCHa), 4.51 (1H, d, J = 11.5, PhCHb), 7.20-7.50 (5H, m, Ph). 3S-Nonanol (20). A solution of 15 (5.38 g, 33.6 mmol) in Py (11 mL) at 0°C was stirred, treated in portions with TsCl (6.90 g, 36.7 mmol), left overnight in a refrigerator, diluted with t-BuOMe (200 mL), washed successively with HCl (10%) and saturated NaHCO3 and NaCl solutions, dried over Na2SO4, and evaporated to afford the tosylate (8.83 g, 80%) [IR spectrum (ν, cm-1): 1605 (Ar), 1375, 1180 (S=O), 1112 (C–O)], which was dissolved in absolute t-BuOMe (150 mL). The resulting solution was cooled to 0°C, treated with LiAlH4 (2.38 g, 62.7 mmol), warmed to room temperature, stirred for 3 h, cooled to 0°C, diluted with H2O (5 mL), warmed to room temperature, and stirred for 3 h. The solid was filtered off and washed with t-BuOMe. The filtrate was dried over Na2SO4 and evaporated to give a product (4 g), column chromatography of which (SiO2, PE:EtOAc, 3:1) gave 20 (3.0 g, 75%), [α]D20 +8.3° (c 0.05, CH3Cl). IR spectrum (ν, cm-1): 3400 (OH), 1150, 1078 (C–O). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.88 (3H, t, J = 7.0, CH3), 0.94 (3H, t, J = 7.3, CH3), 1.25-1.37 (7H, m, 3CH2, OH), 1.30-1.60 (6H, m, H-2, H-4, H-5), 3.45-3.55 (1H, m, H-3). 13C NMR spectrum (CDCl , δ, ppm): 9.79 (q, C-1), 13.98 (q, C-9), 22.56 (t, C-8), 25.56 (t, C-5), 29.32 (t, C-6), 30.00 3 (t, C-2), 31.78 (t, C-7), 36.85 (t, C-2), 73.20 (d, C-3). Nitrite Syntheses. 1. A 3-necked flask equipped with a thermometer, stirrer, and dropping funnel reaching to the bottom of the flask was charged with NaNO2 (0.42 g, 6.1 mmol) and H2O (1.66 mL) and cooled to 0°C. A cold (0°C) solution of a mixture of 6 and 7 (14:1, 1.12 g, 5.5 mmol) prepared as above in H2SO4 (0.28 g, 2.8 mmol) and H2O (0.11 mL) was added through the dropping funnel with stirring at a rate so that gas was practically not evolved and the temperature did not rise. After the addition the reaction mixture was diluted with t-BuOMe (50 mL), washed with saturated NaCl solution, dried over Na2SO4, and evaporated to give a mixture of 2 and 21 (1.11 g) that was used without further purification. IR spectrum (ν, cm-1): 1750, 1650, 1620 (N=O), 790, 780, 620 (O–N). 2. Analogously 19 (2.58 g, 10.3 mmol) afforded 3 (2.4 g) that was used without further purification. IR spectrum (ν, cm-1): 1650 (N=O), 1605 (Ar), 1080, 820, 750 (O–N). 3. Analogously 20 (2.20 g, 15.3 mmol) afforded 4 (1.61 g) that was used without further purification. IR spectrum (ν, cm-1): 1645 (N=O), 1085, 820 (O–N).

647

Photolysis of Nitrites. 1. A solution of a mixture of 2 and 21 (0.40 g) in benzene (70 mL) was placed in a thermostatted Pyrex glass cell and irradiated (Ar, 27°C) for 1.5 h. The solvent was evaporated. The solid (0.36 g) was chromatographed (PE:t-BuOMe, 15:1) to isolate and identify a mixture of 6 and 7 (0.08 g), 15 (0.20 g), and 22 (0.01 g). 1-Acetoxy-3-nonanone (22). IR spectrum (ν, cm-1): 1740 (OC=O), 1714 (C=O). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.90 (3H, t, J = 6.8, CH3), 1.20-1.40 (6H, m, 3CH2), 1.50-1.65 (2H, m, H-5), 2.05 (3H, s, OAc), 2.42 (2H, t, J = 7.4, H-4), 2.72 (2H, t, J = 6.3, H-2), 4.32 (2H, t, J = 6.3, H-1). 13C NMR spectrum (CDCl , δ, ppm): 13.98 (q, C-9), 20.85 (q, C–CO), 22.44 (t, C-8), 23.53 (t, C-5), 29.00 (t, C-6), 3 31.54 (t, C-7), 41.20 (t, C-2), 43.22 (t, C-4), 59.38 (t, C-1), 170.90 (s, CO2), 208.08 (d, C-3). 2. Photolysis of 3 (0.40 g) by the method described above gave a mixture of products (0.40 g), column chromatography of which isolated and identified 19 (0.13 g), 23 (0.06 g), 15 (0.02 g), and 24 (0.16 g). 3R-Benzyloxynonanal (23). IR spectrum (ν, cm-1): 1720 (C=O), 1650 (Ar). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.90 (3H, t, J = 6.7, CH3), 1.20-1.45 (6H, m, 3CH2), 1.50-1.65 (2H, m, H-5), 1.65-1.75 (2H, m, H-4), 2.60 (1H, ddd, J = 16.5, J = 4.9, J = 2.0, Ha-2), 2.67 (1H, ddd, J = 16.5, J = 7.2, J = 2.6, Hb-2), 3.95-4.00 (1H, m, H-3), 4.45-4.60 (2H, m, CH2-Ph), 7.25-7.38 (5H, m, Ph), 9.81 (1H, t, J = 2.2, CHO). 2-Phenyl-4R-hexyl-1,3-dioxane (24), [α]D20 +15.7° (c 0.02, CHCl3). IR spectrum (ν, cm-1): 1640 (Ar), 1155, 1125, 1040 (O–C–O),. PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.87 (3H, t, J = 7.02, CH3), 1.25-1.40 (6H, m, 3CH2), 1.40-1.75 (5H, m, H-1′, H-2′, He-5), 1.82 (1H, dtd, J = 14.0, J = 11.4, J = 4.8, Ha-5), 3.83 (1H, dddd, J = 11.4, J= 7.1, J = 4.8, J = 2.4, Ha-4), 3.95 (1H, td, J = 11.4, J = 2.3, Ha-6), 4.28 (1H, ddd, J = 11.4, J = 4.8, J = 1.2, He-6), 5.52 (1H, s, H-2), 7.25-7.55 (5H, m, Ph). 13C NMR spectrum (CDCl , δ, ppm): 14.04 (q, C-6′), 22.58 (t, C-5′), 24.92 (t, C-2′), 29.23 (t, C-3′), 31.34 (t, C-4′), 3 31.77 (t, C-1′), 36.01 (t, C-5), 67.09 (d, C-6), 77.41 (t, C-4), 101.09 (d, C-2), 125.99, 128.13, 128.56, 138.93 (m, Ph). 3. Photolysis of 4 (0.40 g) by the method descibed above gave a mixture of products (0.27 g), column chromatography of which isolated and identified 25 (0.02 g), 20 (0.08 g), and 26 (0.15 g). 3-Nonanone (25). IR spectrum (ν, cm-1): 1714 (C=O). PMR spectrum (CDCl3, δ, ppm, J/Hz): 0.90 (3H, t, J = 6.6, H-9), 1.15 (3H, t, J = 7.5, H-1), 1.20-1.35 (6H, m, 3CH2), 1.59 (2H, quintet, J = 7.3, H-5), 2.39 (2H, t, J = 7.4, H-2), 2.42 (2H, q, J = 7.3, H-4). 13C NMR spectrum (CDCl , δ, ppm): 7.77 (q, C-1), 13.95 (q, C-9), 22.56 (t, C-8), 24.87 (t, C-5), 29.64 (t, C-6), 31.82 3 (t, C-3), 35.76 (t, C-2), 42.37 (t, C-4), 211.85 (s, C-3). Dimer of 6-nitroso-3S-nonanol (26), [α]D20 +5.5° (c 0.01, CH2Cl2). IR spectrum (ν, cm-1): 3400 (OH), 1378, 1240, (N=O), 1174 (C–O). PMR spectrum (CDCl3, δ, ppm): 0.80-1.00 (6H, m, 2CH3), 1.20-2.00 (11H, m, 5CH2, OH), 3.40-3.55 (1H, m, H-3), 5.35-5.50 (1H, m, H-6). 13C NMR spectrum (CDCl , δ, ppm): 9.85 (q, C-1), 13.60 (q, C-9), 19.08 (t, C-8), 28.17 (t, C-5), 29.96 (t, C-2), 32.74 3 (t, C-4), 34.14 (t, C-7), 66.59 (d, C-6), 72.22 (d, C-3). 6-Oximino-3S-nonanol (27). Dimer 26 was heated at 60°C for 48 h to produce in quantitative yield an oily product that was a chromatographically inseparable equal mixture of the syn- and anti-isomers of 27, [α]D20 -6.2° (c 0.02, CH2Cl2). IR spectrum (ν, cm-1): 3500 (OH), 1660 (C=N), 1120 (C–O), 960 (N–O). PMR spectrum (CDCl3, δ, ppm, J/Hz): syn: 0.90-1.00 (6H, m, H-1,9), 1.45-1.60 (4H, m, H-2,8), 3.53 (1H, tdd, 3J = 6.3, 3J = 3,5, 3J = 9.5, H-3), 1.60-1.70 (2H, m, H-4), 2.25 (1H, ddd, 3J = 5.3, 3J = 7.5, 2J = 13.0, H -5), 2.70 (1H, td, a 3J = 8.6, 2J = 13.0, H -5), 2.15 (2H, t, 3J = 7.4, H-7), 1.20-1.70 (2H, br.s, OH). b anti: 0.90-1.00 (6H, m, H-1,9), 1.45-1.60 (4H, m, H-2,8), 3.42 (1H, tdd, 3J = 6.3, 3J = 3.5, 3J = 9.5, H-3), 1.60-1.70 (2H, m, H-4), 2.25 (1H, ddd, 3J = 5.3, 3J = 7.5, 2J = 13.0, Ha-5), 2.40 (1H, td, 3J = 8.6, 2J = 13.0, Hb-5), 2.35 (2H, t, 3J = 7.7, H-7), 1.20-1.70 (2H, br.s, OH). 13C NMR spectrum (CDCl , δ, ppm): syn: 10.04 (C-1), 30.06 (C-2), 72.10 (C-3), 33.04 (C-4), 23.46 (C-5), 161.85 3 (C-6), 35.88 (C-7), 19.04 (C-8), 14.30 (C-9). anti: 10.11 (C-1), 29.70 (C-2), 71.99 (C-3), 32.68 (C-4), 29.76 (C-5), 161.85 (C-6), 30.45 (C-7), 19.71 (C-8), 13.79 (C-9).

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REFERENCES 1. 2. 3. 4. 5. 6.

J. Kula, T. B. Quang, and M. Sikora, Tetrahedron: Asymmetry, 11, 943 (2000). B. A. Kazanskii, ed., in: Organic Synthesis New York (1946). D. H. R. Barton, R. H. Hesse, M. M. Pechet, and L. C. Smith, J. Chem. Soc., Perkin Trans. 1, 1159 (1979). P. Kabasakalian and E. Townley, J. Am. Chem. Soc., 84, 2711 (1962). N. K. Kochetkov and L. V. Bakinovskii, eds., Comprehensive Organic Chemistry, Vol. 3, Pergamon Press, Oxford (1979). E. Pretsch, T. Clerc, J. Seibl, and W. Simon, Tables of Spectral Data for Structure Determination of Organic Compounds, Springer-Verlag, Berlin (1983).

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Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

CHEMICAL CONSTITUENTS OF BROWN RICE GRAIN (Oryza sativa)

I. M. Chung,1 Mohd Ali,2 A. Ahmad,1 C. Y. Yu,3 K. H. Ma,4 J. G. Gwag,4 and Y. J. Park5

UDC 547.915

One new compound 3,7,11,15,19-pentamethyl-9α,10α,11α,17α,18α-pentahydroxy-n-tetracosan-1-oxy-phydroxycaffeoate (oryzaterpenyl caffeoate) (1), together with three known fatty acids linoleic acid, stearic acid and myristic acid were isolated and identified from the rice grain of Oryza sativa. The structure of the 1 1 1 13 new compound was elucidated by 1D and 2D NMR spectroscopic techniques ( H– HCOSY, H– C HETCOR) aided by EI-MS, and IR spectra. Key words: Oryza sativa, Poaceae, rice grains, 3,7,11,15,19-pentamethyl-9α,10α,11α,17α,18α-pentahydroxy-ntetracosan-1-oxy-p-hydroxycaffeoate, fatty acids. Rice (Oryza sativa) is the principal cereal food in Asia, the major staple food of the majority of the population, and is generally of two types; white hulled and colored hulls, but the most common is white (85%). On the variety front around the globe three major varieties are produced, of which Javanica, a medium grain variety, is grown only in Indonesia, and another two varieties; a long grain variety known as Indica, best suited for warm climate, is cultivated throughout South and Southeast Asia, and Central and South America. The other major variety the round grain Japonica; is well suited for cold the climate of East and Northeast Asia as well as some parts of North America like California. Color hulled rice, though not used by many, is still popular in China and Japan. One of the colored rice brown or red is believed to be rich in vitamin B and also has high fibre, and carbohydrate and protein contents, thus classifying it as a healthy food. The absence of gluten in this rice also makes it nonallergic to masses. This healthy rice is reported to have a marked effect on reducing the risk of coronary heart disease in both men and women, where a 36% reduction in heart attack is observed in men consuming such rice. Prolamin and glutelin compounds from mature and developing rice grains [1–2], which are polymeric procyanidins that are radical-scavenging components [3], have been reported. Several lipids from brown rice have been reported in the literature [4]. The antioxidant activity of brown rice, measured by DPPH and TBA methods, have also been described [5]. The literature does not contain much information even on the chemical constituents of its grain except for four reports [1–4]. This paper deals with the isolation and structural elucidation of a new compound 1 based on 1H NMR, 13C NMR, 1H–1H COSY, 1H–13C HETCOR aided by EIMS and IR spectra and three known fatty acids: linoleic, stearic, and myristic. Oryzaterpenyl Caffeoate (1). Oryzaterpenyl caffeoate 1 was obtained as a yellow semi-solid in minor quantity from the aqueous methanol (80%) extract of the rice grains. Its IR spectrum exhibited characteristeric absorption bands for hydroxy (3465 cm-1) and ester (1730 cm-1) groups. The mass spectrum of 1 displayed a molecular ion peak at m/z 65 corresponding to an acyclic nor-triterpene esterified with hydroxycaffeoic acid moiety, C38H68O8. The prominent ion peaks generated at m/z 149 [HO-C6H4-CH2CH2CO]+, 503 [M-149]+ and 165 [HO-C6H4CH2CH2COO]+ supported the esterification of the hydroxycaffeoyl moiety with the triterpene alcohol. The prominent ion peaks at m/z 71, 581 [C19–C20 fission]+, 99 [C18–C19 fission]+, 129 [C17–C18 fission]+, 159, 493 [C16–C17 fission]+, 479 [C15–C16 fission]+, 201, 451[C14–C15 fission]+, 215 [C13–C14 fission]+, 1) Department of Applied Life Science, Konkuk University, Seoul 143-701, South Korea, tel: +82 2 450 3730, fax 82 2 446 7856, e-mail: [email protected]; 2) Faculty of Pharmacy, Hamdard University, New Delhi-110062, India; 3) Bioherb Research Center, Division of Bioresources Technology, Kangwon National University, Chunchon 200-701, Korea; 4) Bio-Resource Research, National Institute of Agricultural Biotechnology, Suhdun-dong, Suwon 441-707, Korea; 5) International Plant Genetic Resources Institute, APO office, Serdang, Malaysia. Published in Khimiya Prirodnykh Soedinenii, No. 6, pp. 535-537, November-December, 2005. Original article submitted November 22, 2004. 650

0009-3130/05/4106-0650 ©2005 Springer Science+Business Media, Inc.

and 229 [C12–C13 fission]+ suggested the location of the two hydroxyl groups at C-18 and C-17. The other intensified ion peaks appearing at m/z 243 [C11–C12 fission]+, 287 [C10–C11 fission]+, 317 [C9–C10 fission]+, 347 [C8–C9 fission]+, 361 [C7–C8 fission]+, 389 [C6–C7 fission]+, 403 [C5–C6 fission]+, 417 [C4–C5 fission]+ and 431 [C3–C4 fission]+ indicated the location of the remaining three hydroxyl groups at C11, C10 and C9. The 1H NMR spectrum of 1 contains four one-proton multiplets at δ 7.71, 7.69, 7.53, and 7.51 assigned to aromatic H-6′, H-8′, H-5′ and H-9′ respectively. A one-proton multiplet at δ 4.32 with a half -width of 6.9 Hz was ascribed to the β-oriented H-9 carbinol proton. Another one-proton multiplet at δ 4.26 with halfwidth of 6.8 Hz was attributed to the β-oriented H-18 carbinol proton. Two one-proton doublets at δ 4.25 (J = 5.9 Hz) and 4.23 (J = 5.9 Hz) were associated with C-1′ oxygenated methylene protons. Another one-proton doublet at δ 4.29 (J = 3.0 Hz) was attributed to the β-oriented H-10 hydroxymethine proton. m/z 581

m/z 165 m/z 493 m/z 451 OH HO

OH 20

24 21

29 HO m/z 71 m/z 99 m/z 129 m/z 159 m/z 201

15 28

5' 8

12

17

m/z 149 6

4

2

1' 1

3'

O CO CH2 CH2

7'

4'

OH

11 27

OH

9' 26

m/z 389

m/z 431

25

m/z 503

m/z 215 m/z 229

Fig. 1. Fragmentation pattern of 1. A one-proton double doublet at δ 4.21 with coupling interactions of 2.3, 6.3 and 4.6 was assigned to carbinol H-17β. A three-proton broad signal at δ 1.25 was due to C-27 tertiary methyl protons attached to the carbinol carbon. Four three-proton doublets at δ 0.92 (J = 7.55 Hz), 0.88 (J = 6.7 Hz), 0.84 (J = 6.5 Hz), and 0.82 (J = 8.0 Hz) were attributed to C-29, C-28, C-26, and C-25 secondary methyl protons, respectively. A three-proton triplet at δ 0.77 (J = 7.6 Hz) was ascribed to C-24 primary methyl protons. The remaining methine and methylene protons appeared between δ 1.25–0.77 suggesting their attachment at the saturated carbon atoms. The 13C NMR spectrum of 1 exhibited signals for the ester carbon δ 167.96, benzene carbon signals between δ 167.91–129.02, and oxygenated carbons at δ 64.50 (C-1), 66.10 (C-9), 66.41 (C-10), 70.98 (C-11), 64.61 (C-18), and 68.37 (C-17). The methyl carbon signals appeared at δ 11.17 (C-24), 11.59 (C-25), 14.31 (C-26), 25.69 (C-27), 19.77 (C-28), and 22.87 (C-29). The remaining methylene and methine carbon signals resonated in the range δ 23.97 - 33.31. The 13 C–1H HETCOR spectrum of 1 showed correlation of C-7′ with H-6′, H-5′, H-8′, and H-9′; C-9 with H-8, H-10, and H-7; C-10 with H-9 and H-8; C-17 with H-15, H-16, H-18, and H-19; C-18 with H-16, H-17, H-19, and H-20. The 1H–1H COSY spectrum of 1 showed 1H–1H correlation of H-5′ with H-6′ and H-8′ with H-9′; H2-1 with H2-2, H-1, and H2-2′. On the basis of the foregoing account the structure of 1 has been formulated as 3,7,11,15,19-pentamethyl-9α,10α,11α,17α,18α-pentahydroxy-ntetracosan-1-oxy-p-hydroxyl caffeoate. The spectroscopic and physical data of known fatty acids are described in the experimental section.

EXPERIMENTAL Melting points were determined on an Electrochemical Eng. melting point apparatus and are uncorrected. Optical rotation was measured on an AA-10 model polarimeter. IR spectra were recorded on a Thermo Mattson 60-AR spectrophotometer. UV spectra were recorded using a UV-vis spectrometer TU-180PC.1H NMR (500 MHz) and 13C NMR (125 MHz) spectra were obtained on a Brucker Avance (DRX-500) using CDCl3 as solvent. EI-Mass spectra were recorded on a JEOL JMS-SX 102 A spectrometer. Column chromatography was performed over silica gel 70–230 (Merck). TLC analyses were performed on precoated silica gel glass plates 60 F254 (Merck) and visualized under UV light and by spraying with a vanillin 1 g/sulfuric acid 5 ml/ethanol 94 ml solution followed by heating (100–110°C). Plant Material. The grains of O. sativa were collected from Konkuk University (experimental farm) Seoul, South Korea in October 2002.

651

Extraction and Isolation of Compounds. The dried grains of O. sativa (2 kg) were finely ground to powder form and extracted with 80% methanol at room temperature. The solvent was evaporated under reduced pressure, the extract was freeze dried, and a powder extract was obtained (26.1 g). The whole extract was subjected to normal phase silica gel (600 g) column chromatography and gave the following fractions. Fractions 1–2 in hexane, frs. 3–4 in hexane–EtOAc (9:1), frs. 5–6, hexane–EtOAc (8:2), frs. 7–10 hexane–EtOAc (7:3), frs. 11–12 in hexane–EtOAc (3:2), frs. 13–14 in hexane–EtOAc (1:1), frs. 15–16 in hexane–EtOAc (2:3), frs. 17–18 in hexane–EtOAc (3:7), frs. 19–20 in hexane–EtOAc (2:8), frs. 21–22 in hexane–EtOAc (1:9), frs. 23–24 in EtOAc, frs. 25–26 in EtOAc–MeOH (9:1), frs. 27–28 in EtOAc–MeOH (7.5:2.5), frs. 29–30 in EtOAc–MeOH (1:1), frs. 31 and 32 in EtOAc–MeOH (2.5:7.5), and frs. 33–34 in MeOH. Fraction 7 after CC and preparative TLC afforded a new compound in minor quantity (1.3 mg) and other known fatty acids (100 mg). Fractions 8–11 after TLC, and after mixing and further CC over silica gel column chromatography also afforded known fatty acids (130 mg). The identify of the compounds was confirmed by comparison with an authentic sample from Sigma. The other polar fraction contain only sugar molecules on the basis of NMR. 3,7,11,15,19-Pentamethyl-9α,10α,11α,17α,18α-pentahydroxy-n-tetracosan-1-oxy-p-hydroxycaffeoate (1). Yellow semi-solid; Rf 0.34 (Hexane–EtOAc, 7:3); UV λmax(CHCl3) 238 nm; [α]D22 + 29.8° (c 0.8, CHCl3); IR (neat): νmax 3465, 2950, 2855, 1730, 1470, 1360, 1235, 1150,1080, 725 cm-1. PMR spectrum (δ, ppm, CDCl3, TMS, J/Hz): 7.71 (1H, m, H-6′), 7.69 (1H, m, H-8′), 7.53 (1H, m, H-5′), 7.51(1H, m, H-9′), 4.32 (1H, m, W1/2 = 6.9, H-9β), 4.29 (1H, d, J = 3.0, H-10β), 4.26 (1H, m, W1/2 = 6.8, H-18β), 4.25 (1H, d, J = 5.9, H2-1′α), 4.23 (1H, d, J = 5.9, H2-1′β), 4.21 (1H, ddd, J = 2.3, J = 6.3, J = 4.6, H-17β), 1.72 (1H, ddd, J = 7.15, J = 4.5, J = 7.0, H-19α), 1.69 (1H, ddd, J = 7.1, J = 6.0, J = 6.0, H-15α), 1.42 (1H, m, H-7α), 1.39 (2H, m, H2-16), 1.37 (2H, m, H2-12), 1.36 (1H, m, H2-8), 1.35 (2H, m, H2-2), 1.33 (1H, m, H-3α), 1.31 (10 H, br s, 5 × CH2), 1.30 (8H, br s, 4 × CH2), 1.25 (3H, br. s, Me-27), 0.92 (3H, d, J = 7.55, Me-29), 0.88 (3H, d, J = 6.7, Me-28), 0.84 (3H, d, J = 6.5, Me-26), 0.82 (3H, d, J = 8.0, Me-25), 0.77 (3H, t, J = 7.6, Me-24). 13 C NMR (CDCl3): δ 64.50 (C-1), 30.58 (C-2), 33.31(C-3), 26.35 (C-4), 26.07 (C-5), 23.97 (C-6), 36.69 (C-7), 34.56 (C-8 ), 66.10 (C-9), 66.41(C-10), 70.98 (C-11), 32.98 (C-12), 29.31 (C-13), 29.14 (C-14), 36.79 (C-15), 32.71 (C-16), 68.37 (C-17), 64.61(C-18), 38.96 (C-19), 29.70 (C-20), 28.99 (C-21), 28.81(C-22), 26.98 (C-23), 11.17(C-24), 11.59 (C-25), 14.31 (C-26), 25.69 (C-27), 19.77 (C-28), 22.87 (C-29), 167.96 (C-1′), 26.95 (C-2′), 26.53(C-3′), 132.69 (C-4′), 131.08 (C-5′), 132.56 (C-6′), 167.91 (C-7′), 131.11 (C-8′), 129.02 (C-9′). EIMS m/z (rel. int.): 652 [M]+ (C38H68O8) (6.1), 596 (34.6), 581 (34.2), 565 (82.6), 550 (45.3), 535 (46.7), 507 (24.2), 503 (12.5), 493 (12.6), 451 (5.6), 431 (20.9), 417 (16.1), 403 (19.7), 389 (18.5), 361 (18.7), 347 (17.4), 317 (53.6), 289 (22.1), 287 (14.6), 249 (10.7), 243 (70.1), 229 (18.1), 215 (27.3), 201 (6.3), 165 (100), 159 (7.3), 149 (47.8), 129 (12.3), 99 (12.3), 71 (60.5), 57 (100). Fatty Acids. Colorless oil; Rf 0.32–0.41 (Hexane–EtOAc, 7:3). Linoleic Acid (2). Colorless solid; Rf 0.32. EIMS m/z (rel.int.): M+ 280 (C18H32O2, linoleic acid) (28.0), 284 (C18H36O2, stearic acid) (5.2), 228 (C14H28O2, myristic acid) (5.6), 256 (100), 238, 213, 185, 171, 129, 79, 55. Stearic Acid (3). colorless solid; Rf 0.41, mp 69–70°C. EIMS m/z (rel. int.) 284 [M]+ (C18H36O2)(5.2), 256 (C16H32O6) (100), 238 (12), 213 (39), 185 (22), 129 (48), 57 (76). Myristic Acid (4). Colorless solid; Rf 0.38; mp 58–59°C.

ACKNOWLEDGEMENT The author (A. Ahmad) is thankful to the Korean Federation of Science and Technologies Societies (KOFST) for the award of a fellowship as senior researcher in Konkuk University, Seoul, South Korea. This study was supported by technology development program for Agriculture and Forestry, Ministry of Agriculture and Forestry, Rep. of Korea.

652

REFERENCES 1. 2. 3. 4. 5.

B. E. Mandac and B. O. Juliano, Phytochemistry, 17, 611 (1978). R. M. Villareal and B. O. Juliano, Phytochemistry, 17, 177 (1978). T. Oki, M. Masuda, M. Kobayashii, Y. Nishiba, S. Furuta, I. Suda, and T. Sato, J. Agric. Food Chem., 50, 7524 (2002). N. H. Choudhury and B. O. Juliano, Phytochemistry, 19, 1063 (1980). I. M. Chung, K. H. Kim, J. K. Ahn, and J. O. Lee, Korean J. Crop Sci., 45, 261 (2000).

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Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

DETERMINATION OF NATURAL VITAMIN E FROM ITALIAN HAZELNUT LEAVES G. Sivakumar and L. Bacchetta

UDC 543.867+577.16

A new potential source of natural vitamin E from thirteen samples of Corylus avellana L. leaves was screened: the major Italian cultivar – Tonda romana (collected from Latium and Sardinia localities); ten local genotypes from Sardinia – Moro seme, Suconcale, Moro, Sarda grossa, Sarda grossa seme, Sarda schiacciata, Coccoredda, Sarda lunga, Sarda piccola and Sarda tardiva; wild genotype – Selvatico from Latium. The determination was performed after optimizing the high-efficiency pressurized liquid extraction (PLE) conditions of α-tocopherol from Italian hazelnut tree green leaves. Moro from Sardinia showed the highest content of α-tocopherol (237.4±0.3 µg/g d.w). Leaves of this genotype may be considered as a potential new source for natural α-tocopherol. Key words: antioxidant, α-tocopherol, hazelnut, nutrceutical.

α-Tocopherol (vitamin E) occurs naturally in hazelnut tree leaves. Hazelnut plants can tolerate some environmental stress because of their antioxidative system which includes antioxidant such as a-tocopherol and biophenolic compounds [1]. These can interact with free radicals preventing the initiation of potentially lethal process, such as lipid peroxidation and subsequent membrane disorganization [2]. The tocopherol molecule, which is composed of a phytyl chain and a chromanone ring, may be incorporated into biological membranes, thus contributing to their physical stability [3]. The pharmacological action of α-tocopherol is not only as a natural antioxidant but also as a preventive against cancer [4]. Moreover, vitamin E is also used in cosmetology [5]. The nutraceutical demand for natural α-tocopherol continues to increase because it is presumed to be more bioactive than synthetic α-tocopherol. The chloroplast of plants contain significant amounts of α-tocopherol, which is the most biologically active isomer. So far, high-proficiency extraction of leaf α-tocopherol in the extract has not been reported. Therefore, the objective of this work was the optimization of high - efficiency pressurized liquid extraction (PLE) conditions of α-tocopherol from green leaves and the development of a high-performance liquid chromatography (HPLC) method for accurate value measurement. Our main goal was to identify new potential natural sources of antioxidants; thus, a preliminary screening was performed considering different Italian hazelnut tree leaves. This method will allow us to predict which of the Italian hazelnut local genotypes could represent an interesting source of natural α-tocopherol for nutraceutical, pharmaceutical, and cosmetic applications. Tocopherols are easily oxidized and oxidation losses can be incremented by heat, light, alkaline pH, and by the presence of free radicals [6]. This is especially critical in samples in which vitamin E can be oxidized during the extraction process and/or in the extract until its final analysis. To separate the α-tocopherol fraction, the lyophilized hazelnut leaf powder was mixed with Hydromatrix in the extraction cell and extracted with dehydrated hexane containing 0.01% of butylated hydroxytoluene in an accelerated solvent extraction. Recently, Lagouri and Boskou [7] found that tocopherols in oregano species were largely responsible for the protective effect of hexane extracts in stored cottonseed oil. Lyophilization gave the opportunity to avoid denaturation of α -tocopherol caused by heating, and maintaining samples frozen throughout drying led to completeness of rehydration. Pure nitrogen was used as a carrier gas because it allowed us to avoid denaturation of α-tocopherol during extraction in an ASE and better resolution of chromatographic peaks. Vitamin E is not chemically bound to proteins, lipids, or carbohydrates [8] and using harsh reagents and conditions to free it up (e.g., strong saponification) can destroy the vitamin. But, PLE results on α-tocopherol extraction efficiency showed that 60°C was a suitable temperature for better extraction in an ASE [1] at higher α-tocopherol contents without oxidation. BIOTEC–GEN, ENEA, Casaccia, Rome, Italy, e-mail: [email protected] Published in Khimiya Prirodnykh Soedinenii, No. 6, pp. 538-540, November-December, 2005. Original article submitted April 6, 2005. 654

0009-3130/05/4106-0654 ©2005 Springer Science+Business Media, Inc.

TABLE 1. α-Tocopherol Content from Italian Hazelnut Tree Leaves Major Italian hazelnut cultivar from different agro-climatic areas of Italy Tonda romana (Latinum) Tonda romana (Sardinia) Sardinia local genotypes Moro Sarda grossa seme Sarda grossa Moro seme Sarda schiacciata Coccoredda Sarda lunga Suconcale Sarda piccola Sarda tardiva Italian wild genotype Selvatico (Latium)

(µg/g d.w)* 149.3±0.4 147.6±0.7 237.4±0.3 225.0±0.3 187.5±0.5 132.9±0.9 89.7±0.5 77.2±0.3 58.3±0.8 54.6±0.3 49.5±0.4 34.6±0.4 150.5±0.6

______ *Mean±standard error. This method was evaluated through repetitive analysis of a standard and hazelnut leaf extracts. PLE can be used to isolate naturally occurring tocopherols from plant sources with the advantages inherent in this technique such as small amount of solvent, speed, and routine analysis [9]. These results show that the α-tocopherol extraction and quantification can be considered accurate. Table 1 shows the α-tocopherol content (mg/g d.w.) of hazelnut leaf samples. Moro leaves from Sardinia local genotypes showed the highest content of α-tocopherol (237.4±0.3); Sarda grossa seme, Sarda grossa, and Moro seme (225.0±0.3, 187.5±0.5, 132.9±0.9, respectively) and Sarda schiacciata, Coccoredda, Sarda lunga, Suconcale, Sarda piccola, and Sarda tardiva (89.7±0.5, 77.2±0.3, 58.3±0.8, 54.6±0.3, 49.5±0.4, 34.6±0.4, respectively) recorded the lowest. One of the major Italian varieties, Tonda romana (a traditional cultivar from Central Italy) collected from different places such as Latium (149.3±0.4) and Sardinia (147.6±0.7) showed a minor content when compared to Moro, Sarda grossa seme,and Sarda grossa, but a higher content than Moro seme and other local Sardinia genotypes. No significant differences were found between the same varieties from different localities. Regarding the α-tocopherol content in wild genotype, Selvatico from Latium showed a similar value (150.5±0.6) when compared to Tonda romana, but in Moro, Sarda grossa seme, and Sarda grossa the opposite was observed. With respect to leaf developmental stage and environmental conditions, there was a tendency for varying accumulation of α-tocopherol content. This was also noted by other workers [10, 11]. This was understood as being due to the synthesis of the α-tocopherol in the early stages of development of the leaves and some portion transferred to nuts as maturation progressed. Because of its hydrophobic nature, almost all cellular α-tocopherol is in the membrane fraction of cells. In that location it can readily donate an electron to a fatty acid hydroperoxyl radical, thus breaking the chain reaction associated with lipid peroxidation [12]. However, wide variations in α-tocopherol levels have been attributed to differences in hazelnut cultivars, maturity, growing practices, climates, etc. [13]. In conclusion, PLE shows several advantages when compared to other extraction methods such as higher extraction efficiency, small amount of solvent, time consumption, etc. Our HPLC screening data from hazelnut leaves could allow natural manipulation of ruminant diets to increase uptake of the natural antioxidant α-tocopherol by meat-producing cattles, thereby enhancing the color and oxidative stability of fresh meat [11]. It is well established from vitamin E deficiency studies that adequate amounts of α-tocopherol are necessary to prevent mitochondrial disfunction. Thus, not only is α-tocopherol required for mitochondrial stability, but it may also prevent radical release. Furthermore, the selection of Italian hazelnut varieties and genotypes could represent an interesting tool to find natural sources of α-tocopherol for nutraceutical, pharmaceutical, and cosmetic applications.

655

EXPERIMENTAL

Sampling and Sample Preparation. Thirteen samples of leaves from Corylus avellana L., collected during June 2004 in two different geographical conditions of Italy [major cultivar – Tonda romana from Latium and Sardinia]; ten local genotypes from Sardinia – Moro seme, Suconcale, Moro, Sarda grossa, Sarda grossa seme, Sarda schiacciata, Coccoredda, Sarda lunga, Sarda piccola, and Sarda tardiva; and wild genotype – Selvatico from Latium were analyzed. Three representative samples were collected from each cultivar, immediately frozen in liquid nitrogen, and milled to a fine powder by a Waring blender unit for 20 s. Then fine leaf powder samples were dehydrated on lyophilizer (Edwards Pirani 501) shelves to –60°C in two days. During all the steps that followed, care was taken to protect the samples from light and atmospheric oxygen. Pressurized Liquid Extraction (PLE). The extraction was performed on ASE 100 (Dionex) using extraction cells of 10 mL, with cellulose filters. The lyophilized leafs powder (1g) was mixed homogeneously with a drying agent (Hydromatrix, VARIAN) in the extraction cell. After placing the cell in the ASE, the selected assay conditions were applied. α-Tocopherol was extracted with dehydrated hexane containing 0.01% of butylated hydroxytoluene (BHT 99%, Sigma-Aldrich, which was added to inhibit the oxidative degradation of α-tocopherol during extraction) in an ASE. The assay cycle on the ASE was temperature 60°, pressure 1500 psi, flush volume 60%, purge time 100 s, number of static cycle 1, and total assay time 15 min. This solution (20 µL) was injected into the HPLC system. The extraction unit was wrapped in aluminum foil. High-performance Liquid Chromatography (HPLC). The normal-phase HPLC system (VARIAN, Prostar) consisted of a VARIAN Prostar 210 pump equipped with a VARIAN Lichrosorb Si 60-5 column (15 × 4mm, 5µm) and a variable wavelength UV-visible detector (VARIAN, Prostar 325), the wavelength of which was set at 292 nm. The isocratic mobile phase contained 0.5% isopropanol in dehydrated hexane and the flow rate was 1 mL/min. The mobile phase was degassed with helium. Pure α-Tocopherol standard was purchased from Sigma. Stock standard solution was prepared in extraction solvent and stored at –20°C in dark bottles. The α-tocopherol peak was identified by adding the standard to the samples before PLE extraction. The α-tocopherol was quantified using the external standard method. The relationship between the concentration of α-tocopherol and peak height value was established by the calibration curve and expressed in µg/g dry weight of α-tocopherol.

ACKNOWLEDGEMENT We thank Dr. Eugenio Benvenuto for helpful suggestions during the course of this project and Giovanna Zappa for HPLC facilities. This work was supported by ENEA, Rome, Italy.

REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

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G. Sivakumar, L. Bacchetta, R.Gatti, and G. Zappa, J. Plant Physiol., 162, 1280 (2005). M.S. Eiberger and G. Noga, Sci. Hortic., 91, 153 (2001). N. V. Gorbunov, V. K. Kagan, S. M. Alekseev, and A. N. Erin, Bull. Exp. Biol. Med., 112, 946, (1988). P. Jha, M. Flather, E. Lonn, M. Farkouh, and S. Yusuf, Ann. Intern. Med., 123, 860 (1995). E. P. Sheppard and M. J. Stutsman, J. Soc. Cosmet. Chem., 28, 115 (1977). D.J.M. Gomez-Coronado, E. Ibanez, F.J. Ruperez, and C. Barbas, J. Chromatogr. A, 1054, 227 (2004). V. Lagouri and D. Boskou, Int. J. Food Sci. Nutr., 47, 439 (1996). E. Herrera, Nutr. Obes., 3, 4 (2000). M. M. D. Zamarreno, M. B. Rangel, A. S. Perez, and R. C. Martinez, J. Chromatogr. A, 1056, 249 (2004). L. R. McDowell, S. N. Williams, N. Hidiroglou, C. A. Njeru, G. M. Hill, L. Ochoa, and N.S Wilkinson, Anim. Feed Sci. Technol, 60, 273 (1996). A. Lynch, J. P. Kerry, D. J. Buckley, P. A. Morrissey, and C. Lopez-Bote, Food Chemistry, 72, 521 (2001). X. Li and J. M. May, Mitochondrion, 3, 29 (2003). M. Ozdemir, F. Ackurt, M. Kaplan, M. Yildiz, M. Loker, T. Gurcan, G. Biringen, A. Okay, and F. G. Seyhan, Food Chemistry, 73, 411 (2001).

Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

PLANT COUMARINS. 2. BECKMANN REARRANGEMENT OF OREOSELONE E- AND Z-OXIMES

I. Yu. Bagryanskaya, Yu. V. Gatilov, S. A. Osadchii, A. A. Martynov, M. M. Shakirov, E. E. Shul′ts, and G. A. Tolstikov

UDC 547.587.51+548.737

Oximation of oreoselone to produce a mixture the E- and Z-oximes was investigated. The crystal and molecular structures of oreoselone Z-oxime and the Beckmann rearrangement product of oreoselone E- or Z-oximes and PCl5, 7-(1-chloro-2-methylpropoxy)-2-oxo-2H-1-benzopyran-6-carbonitrile, were established by XSA. Hydrolysis of the latter produced 7-hydroxy-2-oxo-2H-1-benzopyran-6-carbonitrile. Key words: coumarins, oreoselone, oxime, 7-(1-chloro-2-methylpropoxy)-2-oxo-2H-1-benzopyran-6-carbonitrile, 7-hydroxy-2-oxo-2H-1-benzopyran-6-carbonitrile, XSA, NMR spectroscopy. In the previous report [1] we prepared a series of peucedanin derivatives by modifying the furan ring. In order to expand the number of derivatives, which are potentially promising as synthons for preparing antiviral (anti-HIV) [2] and antitumor [3] agents, we attempted to prepare derivatives from racemic oximes of oreoselone (1), which is easily prepared by hydrolysis of peucedanin. The synthesis of racemic oreoselone oxime {3-hydroxyimino-2-(1-methylethyl)-7H-furo[3,2-g][1]benzopyran-7-one} of undetermined configuration with mp 200-202°C (dec., from EtOH) was previously reported [4]. Classical oximation [5] of 1 by a solution of the free base produced by alkaline neutralization of an aqueous alcohol solution of hydroxylamine hydrochloride (the excess of base was 3.2 mole relative to oreoselone) was used. We found by PMR that the product produced this way [4] was a mixture of the two isomeric oreoselone oximes, 3Z- and 3E-hydroxyimino-2-(1-methylethyl)-7H-furo[3,2g][1]-benzopyran-7-one [(Z)-2 and (E)-2]. In order to avoid the large excess of hydroxylamine, we investigated oximation of racemic 1 by boiling with an alcoholic pyridine solution of hydroxylamine hydrochloride (mole ratio 1:hydroxylamine hydrochloride:pyridine 1.00:1.05:2.00). According to PMR spectra, the reaction produced in 52% yield a crystalline mixture of (Z)-2 and (E)-2 with an E:Z ratio of about 1:0.7. The low yield of the mixture of oxime isomers under these conditions is apparently due to the formation of side products via reaction of hydroxylamine with the lactone ring of 1 and (or) its hydroxyimino derivatives (cf. the reaction of coumarin and hydroxylamine [6, 7]). O 3

H (CH3) 2HC

O

O

O

1 NH2OH ⋅ HCl HO H (CH3) 2HC 1'

Py, EtOH OH

N

N 3a

4a

9a

8a

H

3 1

O

(Z)-2

O

O

(CH3) 2HC

O

O

O

(E)-2

N. N. Vorozhtsov Novosibirsk Institute of Organic Chemistry, Siberian Division, Russian Academy of Sciences, 630090, Novosibirsk, pr. Lavrent′eva, 9, fax (3832) 330 97 52, e-mail: [email protected]. Translated from Khimiya Prirodnykh Soedinenii, No. 6, pp. 541-545, November-December, 2005. Original article submitted September 27, 2005. 0009-3130/05/4106-0657 ©2005 Springer Science+Business Media, Inc.

657

S O2 H O2

S

N1 C3

C 11

C5

C 3A

C 10 O1

H6 H6

C4 C 4A

C2

O3

O3 H

C6 S

C 8A

C 9a

R

C7

O2

C9

C 12

O4 O3

R

Fig. 1

Fig. 2

Fig. 1. Molecular structure of (Z)-2 and selected bond lengths from the XSA: O(1)–C(9A) 1.353(2), O(1)–C(2) 1.467(2), C(3)–N(1) 1.290(3), O(2)–N(1) 1.403(2), C(7)–O(3) 1.208(2), C(7)–O(4) 1.375(2), O(4)–C(8A) 1.377(2) Å. Fig. 2. Crystal packing of (Z)-2 [configurations of the C2 chiral centers (R or S) are shown]. Fractional crystallization of the mixture of oximes prepared in the alcoholic pyridine solution isolated both isomers. The Z-configuration was established by an x-ray structure analysis (XSA) for a single crystal of the pure compound with mp 242-244°C (dec.). Figure 1 shows the molecular structure of (Z)-2 and selected bond lengths from the XSA. All atoms of the framework of (Z)-2 lie in a single plane [mean-square deviation 0.016 Å; maximum deviation from the plane, 0.045(2) Å for C2]. The oxime group also lies practically in this same plane. Atoms N1 and O2 deviate from the plane by 0.114 and 0.149 Å, respectively. The torsion angle C2C3N1O2 is 0.3(3)°. Thus, the oxime group has the E-configuration. The bond lengths and angles in (Z)-2 are within 3σ of the statistical average values [8]. It should be noted only that the O1–C9A bond is shorter [1.353(2) Å] than the statistical average value of 1.374(13) Å. Among the 40 structures with a linear furocoumarin framework in the Cambridge crystallographic database [9], none were found with an exocyclic double bond in the 3-position. Figure 2 shows a portion of the crystal packing of (Z)-2. The molecules form infinite chains through H-bonds. The coordinates of the OH hydrogen shifted upon refinement rather significantly toward the ketone in the neighboring molecule: O(2)–H 1.28(5), O(2)...O(3) 2.715(2) Å, H...O(3) 1.46(5) Å, and O(2)–H...O(3) 164(4)°. These chains, in turn, are bonded to each other through short intermolecular O...H contacts: O(3)...H(6) 2.47(2) Å. Pairs of neighboring molecules can be found in the crystal packing of (Z)-2 that form head—tail dimers through π-stacking interactions of the aryl rings (interplanar distance 3.49, distance between aryl-ring centers 3.86 Å). These dimers bind the double chains pairwise to each other to form a onedimensional supramolecular motif in the crystal. It should be noted that weak nonbonded interactions are at present of great interest owing to their important role in the regulation of the packing of organic molecules in crystalline solids. The structure of the isomeric oxime (E)-2 [mp 193-195°C (dec.)] was established by comparing the PMR and 13C NMR spectra of the isomers and by elemental analysis. We attempted to perform a Beckmann rearrangement (BR) on the prepared oximes using in the first step PCl5. The predominant crystalline product (70-75% yield) from reaction of (E)-2 or (Z)-2 with PCl5 was 7-(1-chloro-2-methylpropoxy)-2oxo-2H-1-benzopyran-6-carbonitrile (3), the formation of which can be represented as a type II BR [10], which assumes cleavage of the C2–C3 bond in the intermediates formed from (E)-2 or (Z)-2.

(E)-2 (Z)-2

658

PCl5

NC Cl

PCl5

(CH3)2HC

H2O (EtOH)

H C O

O 3

O

NC

+ HO

O 4

O

(CH3)2HCCHO

C4 N1

C 13

b

C 4A

C6 O3

C3

C5 C 8A C7

C8

C2 O1

O2

C l1

C 12

Z

C9

c



C 10 C 11

Fig. 3

Fig. 4

Fig. 3. Molecular structure of 3 and selected bond lengths (Å) and torsion angles (°) from the XSA: O(1)–C(8A) 1.371(3), O(1)–C(2) 1.386(3), C(2)–O(2) 1.202(3), C(7)–O(3) 1.356(3), O(3)–C(9) 1.410(3), Cl(1)–C(9) 1.808(2), C(6)–C(13) 1.441(4), C(13)–N(1) 1.133(3), C(7)O(3)C(9)Cl(1) 72.4(2), C(7)O(3)C(9)C(10) -166.1(2), O(3)C(9)C(10)C(12) -57.0(3). Fig. 4. Crystal packing of 3 along the a axis. The structure of 3 was established by a single-crystal XSA. Figure 3 shows the molecular structure of 3 and selected bond lengths from the XSA. All atoms of the bicyclic framework of 3 lie in a single plane with a mean-square deviation of 0.008 Å. Atoms O(2), O(3), and the cyano group C(13)≡N(1) are practically in this same plane with deviations from the plane of 0.026(3), 0.035(2), 0.020(3), and 0.069(4) Å, respectively. The chlorobutyl group eclipses C(8) with a C(9)O(3)C(7)C(8) torsion angle of 5.5(3)°. The ethyl group in 7-ethoxycoumarin has a similar orientation [11]. The bond lengths and angles in 3 are within 3σ of the statistical average values [12]. The crystal packing of 3 (Fig. 4) shows centrosymmetric pairs of molecules that form head—tail dimers through π-stacking interactions of the aryl rings [interplanar distance 3.55; distance between centers, 3.861(1) Å]. Brief (0.5 h) boiling in aqueous alcohol cleaves 3 to form 7-hydroxy-2-oxo-2H-1-benzopyran-6-carbonitrile (4) and isobutyraldehyde. Thus, the E- and Z-oximes of 1 were prepared. The crystal and molecular structures of the Z-oxime and of the BR product of the E- or Z-oxime and PCl5 -[7-(1-chloro-2-methylpropoxy)-2-oxo-2H-1-benzopyran-6-carbonitrile] were established by XSA. Hydrolysis of the latter gave 7-hydroxy-2-oxo-2H-1-benzopyran-6-carbonitrile.

EXPERIMENTAL We used freshly distilled solvents and pure-grade reagents. Recemic 1 was prepared by hydrolysis of peucedanin by the literature method [13] with a modification [1]. Melting points were determined on a Kofler apparatus. IR spectra were recorded on a Vector 22 spectrometer in KBr disks; UV spectra, on a Specord UV—Vis spectrophotometer in ethanol (c 10-4 M). Molecular weights and elemental composition of new compounds were determined using a high-resolution mass spectrometer (Finnigan MAT, model 8200, EI, 70 eV). NMR spectra were obtained on Bruker AC 200 (working frequency 1H 200.13 MHz, 13C 50.32 MHz), Bruker AM-400 (1H 400.13 MHz, 13C 100.61 MHz), and Bruker DRX-500 (1H 500.13 MHz, 13C 125.76 MHz) instruments using solutions (10%) at 25°C and resonances stabilized on the solvent D signal (CDCl or 3 DMSO-d6). Chemical shifts (ppm) were measured relative to those of internal standards CHCl3 (δH 7.24 ppm and δC 76.90 ppm) and DMSO (δH 2.50 ppm and δC 39.50 ppm). The multiplicity of signals in the 13C NMR spectra was determined by standard methods with J-modulation (JMOD) and with off-resonance irradiation of protons. Signals in the PMR and

659

13C NMR of (Z)-2 and (E)-2 were assigned using data for the model compound 1 [1, 14].

Signals in 13C NMR spectra of 3 were

Two-dimensional 13C—1H spectra (COSY 125 Hz, COLOC 7-10 Hz) were

assigned using various types of C—H correlation. recorded on a Bruker DRX-500 instrument using standard Bruker programs. Analytical TLC was performed on Sorbfil UV 254 plates (ZAO Sorbpolimer, Krasnodar, RF) with AcOEt:hexane (1:2 by vol) eluent. Mobilities (Rf) for 1, (E)-2 oxime, and (Z)-2 oxime were 0.65, 0.55, and 0.30, respectively, with Rf of (3) > Rf of (E)-2 oxime. X-ray structure analyses were carried out on a Bruker P4 diffractometer (Mo Kα-radiation, graphite monochromator, 2θ/θ-scanning, 2θ < 52°). A light yellow crystalline prism of (Z)-2 of dimensions 0.50 × 0.30 × 0.17 mm was selected. The crystal was triclinic and racemic, a = 5.9047(7), b = 10.015(1), c = 11.039(1) Å, α = 87.25(1)°, β = 78.777(8)°, γ = 81.500(9)°, V = 633.2(1) Å3, space group P1, Z = 2, C14H13NO4, dcalc = 1.360 g/cm3, µ = 0.101 mm-1. Intensities of 2212 independent reflections were measured. Absorption corrections were applied using the crystal faces (transmission 0.97-0.98). For 3, a colorless crystalline plate of dimensions 0.60 × 0.30 × 0.1 mm was selected. The crystal was monoclinic, a = 12.762(2), b = 9.697(1), c = 11.331(2) Å, β = 105.27(1)°, V = 1352.7(4) Å3, space group P21/c, Z = 4, C14H12ClNO3, dcalc = 1.364 g/cm3, µ = 0.285 mm-1. Intensities of 2650 independent reflections were measured. Absorption corrections were not applied. The structures were solved by direct methods using the SHELXS-97 program [15]. Structure factors were refined by full-matrix anisotropic least-squares techniques using the SHELXL-97 program [15]. Factors for H atoms were calculated in each refinement cycle using coordinates of the corresponding C atoms. Final refinement of 172 parameters for the structure of (Z)-2 over all F2 gave wR2 = 0.1321 and S = 1.002 (R = 0.0463 for 1645 F > 4σ). Final refinement of 225 parameters for the structure of 3 over all F2 gave wR2 = 0.1349 and S = 1.051 (R = 0.0443 for 1693 F > 4σ). Coordinates and temperature factors for the atoms were deposited in the Cambridge Crystallographic Database (registration numbers CCDC 271987 and CCDC 271988 for (Z)-2 and 3, respectively). Oximation of 1. A stirred mixture of hydroxylamine hydrochloride (1.80 g, 25.9 mmol) and absolutee pyridine (4.0 mL) was treated with absolute alcohol (44 mL). The resulting solution was treated with 1 (6.00 g, 24.6 mmol), stirred, and refluxed for 5 h. 1 dissolved completely to give a green solution after 30 min from the start of refluxing. Then solvent was removed from the mixture by heating on a bath. Residual solvent was removed at 80°C/30 torr. The resinous solid was treated with a mixture of CHCl3 (60 mL) and water (10 mL). The aqueous layer was removed. The organic layer was washed with additional water (10 mL) and evaporated. The resinous solid was dried at 80°C/30 torr and dissolved in ether (15 mL). The crystalline solid that formed on standing was filtered off, washed with ether, and dried to afford a product (3.31 g, 52%) that was a mixture of the Z- and E-oximes of 1 (E:Z = 1:0.7) according to PMR spectra. The Z-oxime was isolated by treating the mixture with boiling CHCl3 (2 × 10 mL). The insoluble solid was recrystallized from boiling alcohol to afford crystalline prisms of (Z)-2, mp 242-244°C (dec.). PMR spectrum (200.13 MHz, DMSO-d6, δ, ppm, J/Hz): 0.64 and 1.08 [both d, 3H each, J = 6.8, (CH3)2C], 2.75 (septet of doublets, 1H, J = 6.8, J = 2.4, H-1′), 5.44 (d, 1H, J = 2.4, H-2), 6.29 (d, 1H, J = 9.6, H-6), 6.98 (s, 1H, H-9), 7.81 (s, 1H, H-4), 7.99 (d, 1H, J = 9.6, H-5), 11.50 (s, 1H, NOH). 13C NMR spectrum of the same solution (50.32 MHz, δ , ppm): 14.8 and 19.3 [both q, (CH ) C], 28.5 (d, C-1′), 88.3 C 3 2 (d, C-2), 98.4 (d, C-9), 112.8 (d, C-6), 113.4 (s, C-3a), 119.4 (s, C-4a), 120.6 (d, C-4), 144.6 (d, C-5), 154.1 (s, C-9a), 157.2 (s, C-8a), 159.9 (s, C-3), 166.2 (s, C-7). Found, %: C 64.70, H 5.02, N 5.46. C14H13NO4. Calc., %: C 64.86, H 5.05, N 5.40. IR spectrum (ν, cm-1): 3310, 2969 (NO–H), 1716 (C=O), 1653, 1621, 1569, 1481, 1465, 1396, 1362, 1299, 1199, 1143, 1112, 1056, 948, 925, 890, 836. UV spectrum (λmax, nm, log ε): 229 (4.15), 256 (4.43), 301 (3.83), 314 (3.86), 351 (4.28). 3E-Hydroxyimino-2-(1-methylethyl)-7H-furo[3,2-g][1]benzopyran-7-one [(E)-2] was isolated by fractional crystallization from CHCl3 of the solid obtained after separation of the Z-oxime, mp 193-195°C (dec.). PMR spectrum (200.13 MHz, DMSO-d6, δ, ppm, J/Hz): 0.83 and 1.03 [both d, 3H each, J = 6.8, (CH3)2C], 2.11 (septet of doublets, 1H, J = 6.8, J = 3.4, H-1′), 5.16 (d, 1H, J = 3.4, H-2), 6.27 (d, 1H, J = 9.6, H-6), 6.96 (s, 1H, H-9), 8.03 (d, 1H, J = 9.6, H-5), 8.39 (s, 1H, H-4), 11.71 (s, 1H, NOH). 13C NMR spectrum of the same solution (50.32 MHz, δ , ppm): 15.1 and 17.8 [both q, (CH ) C], 32.7 (d, C-1′), 87.7 C 3 2 (d, C-2), 98.4 (d, C-9), 112.4 (d, C-6), 112.9 (s, C-3a), 117.1 (s, C-4a), 128.2 (d, C-4), 144.8 (d, C-5), 152.2 (s, C-9a), 157.4 (s, C-8a), 159.8 (s, C-3), 166.5 (s, C-7). Found, %: C 64.61, H 5.02, N 5.37. C14H13NO4. Calc., %: C 64.86, H 5.05, N 5.40. IR spectrum (ν, cm-1): 3273, 2970, 2932 (NO–H), 1744, 1721, 1625, 1568, 1465, 1394, 1365, 1258, 1221, 1204, 1144, 1103, 973, 912, 824 (C=O).

660

UV spectrum (λmax, nm, log ε): 229 (4.15), 256 (4.43), 301 (3.83), 314 (3.86), 351 (4.28). (Z)-2 and (E)-2 were isolated analogously by oximation of 1 using another method [4]. 7-(1-Chloro-2-methylpropoxy)-2-oxo-2H-1-benzopyran-6-carbonitrile (3). PCl5 (574 mg, 2.76 mmol) was dissolved in refluxing dry benzene (11.3 mL). The solution was cooled to 20°C, treated in one portion with (Z)-2 (714 mg, 2.76 mmol), stirred until the solution turned yellow (slightly exothermic reaction), boiled for 20 min, cooled to 10°C, stirred, and treated with icewater (11 mL) and ether (22 mL). The ether layer was separated. The aqueous layer was extracted with ether (2 × 5 mL). The combined organic layers were dried over MgSO4. Solvent was removed. The solid was dried at 30°C/1 torr and treated with EtOH (4 mL) to afford 3 (547 mg, 71%), mp 150-151°C (hot aqueous 90% EtOH). High-resolution MS. Found, m/z: 277.05079. C14H12ClNO3. Calc., m/z: 277.05056. PMR spectrum (200.13 MHz, CDCl3, δ, ppm, J/Hz): 1.17 and 1.19 [both d, 3H each, J = 6.8, (CH3)2C], 2.43 (septet of doublets, 1H, J = 6.8, J = 4.2, H-2′), 5.90 (d, 1H, J = 4.2, H-1′), 6.39 (d, 1H, J = 9.6, H-3), 7.21 (s, 1H, H-8), 7.65 (d, 1H, J = 9.6, H-4), 7.76 (s, 1H, H-5). 13C NMR spectrum of the same solution (125.77 MHz, δ , ppm*): 16.97 and 17.01 [both q, (CH ) C], 36.00 (d, C-2′), C 3 2 95.86 (d, C-1′), 100.07 (s, C-6)a, 103.32 (d, C-8), 114.03 (s, C-4a)a, 114.08 (s, CN), 115.84 (d, C-3), 133.36 (d, C-5), 141.64 (d, C-4), 157.60 (s, C-8a)b, 158.79 (s, C-7)b, 158.97 (s, C-2)b. IR spectrum (ν, cm-1): 3058, 2977, 2883, 2232 (C≡N), 1739 (C=O), 1625, 1495, 1386, 1334, 1285, 1204, 1164, 1128, 1031, 919, 831, 737. UV spectrum (EtOH, λmax, nm, log ε): 210 (4.37), 237 (4.35), 294 (3.93), 320 (3.99). Compound 3 was prepared analogously from (E)-2 in 70% yield. 7-Hydroxy-2-oxo-2H-1-benzopyran-6-carbonitrile (4). A solution of 3 (115 mg) in aqueous alcohol (1.0 mL, 90% EtOH by vol) was refluxed for 0.5 h. The needlelike crystals that formed on cooling were separated, washed with CHCl3, and dried to afford 4 (71 mg, 92%), mp 268-270°C (dec.). High-resolution MS. Found, m/z: 187.02706. C10H5NO3. Calc., m/z: 187.02694. PMR spectrum (400.13 MHz, DMSO-d6, δ, ppm, J/Hz): 6.29 (d, 1H, J = 9.6, H-3), 6.80 (s, 1H, H-8), 7.86 (d, 1H, J = 9.6, H-4), 8.00 (s, 1H, H-5). 13C NMR spectrum of the same solution (100.61 MHz, δ , ppm*): 96.8 (s, C-6), 102.9 (d, C-8), 112.0 (s, C-4a), 113.4 C (d, C-3), 115.8 (s, CN), 134.3 (d, C-5), 143.2 (d, C-4), 157.6 (s, C-8a)a, 159.3 (s, C-7)a, 162.6 (s, C-2). *Assignments denoted with the same superscript should possibly be switched. IR spectrum (ν, cm-1): 3487 and 3434 (OH), 3088, 3057, 2761, 2669, 2242 (C≡N), 1726 (C=O), 1707, 1627, 1515, 1446, 1405, 1329, 1295, 1272, 1142, 1100, 910, 856, 835, 672, 573, 522, 456. UV spectrum (EtOH, λmax, nm, log ε): 210 (4.37), 237 (4.35), 294 (3.93), 320 (3.99). Isobutyraldehyde was identified using the PMR spectrum in the mother liquor from 4.

ACKNOWLEDGMENT The work was supported financially by the RFFR (project No. 03-03-33093) and through an integrated project of the SD and FED of the RAS No. 43. We thank the RFFR (project 02-07-90322) for assistance in purchasing the license for the Cambridge Crystallographic Database. REFERENCES 1. 2. 3. 4. 5.

S. A. Osadchii, E. E. Shults, M. M. Shakirov, and G. A. Tolstikov, "Some Transformations of Peucedanin," in: Abstracts Int. Conf. Nat. Prod. and Phys. Active Substances, Novosibirsk, Russia (2004), p. 95. H.-B. Yuan and A. L. Parrill, Bioorg. Med. Chem., 10, 4169 (2002). C. Ito, M. Itoigawa, Y. Mishina, V. C. Filho, F. Enjo, H. Tokuda, H. Hishino, and H. Furukawa, J. Nat. Prod., 66, 368 (2003). G. K. Nikonov, Zh. Obshch. Khim., 31, 305 (1961). H. Bauer and H. Moll, Die Organische Analyse, Akademische Verlagsgesellschaft, Leipzig (1960), p. 262, 265.

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6. 7. 8. 9. 10. 11. 12. 13. 14. 15.

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R. D. H. Murray, J. Mendez, and S. A. Brown, The Natural Coumarins. Occurrence, Chemistry and Biochemistry, John Wiley & Sons, Ltd., Chichester, New York, etc. (1982), pp. 56-61. G. A. Kuznetsova, Natural Coumarins and Furocoumarins [in Russian], Nauka, Leningrad (1967). Cambridge Crystallographic Data Centre, program MOGUL 1.0 (2004). F. H. Allen, Acta Crystallogr. Sect. B: Struct. Sci., 58, 380 (2002). K. V. Vatsuro and G. L. Mishchenko, Name Reactions in Organic Chemistry [in Russian], Khimiya, Moscow (1976). K. Ueno, Acta Crystallogr. Sect. C: Cryst. Struct. Commun., 41, 1786 (1985). F. H. Allen, O. Kennard, D. G. Watson, L. Brammer, A. G. Orpen, and R. Taylor, J. Chem. Soc. Perkin Trans. II, No. 12, S1 (1987). H. Schmid and A. Ebnoether, Helv. Chim. Acta, 34, 1982 (1951). E. E. Shul′ts, T. N. Petrova, M. M. Shakirov, E. I. Chernyak, L. M. Pokrovskii, S. A. Nekhoroshev, and G. A. Tolstikov, Khim. Interesakh Ustoich. Razvit., 683 (2003). G. M. Sheldrick, SHELX-97 release 97-2, University of Gottingen, Germany (1998).

Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

MODIFIED COUMARINS. 19. SYNTHESIS OF NEOFLAVONE D-GLYCOPYRANOSIDES

Ya. L. Garazd,1 M. M. Garazd,2 and V. P. Khilya1

UDC 547.814.5

Neoflavone β-D-glucopyranosides, β-D-galactopyranosides, β-D-xylopyranosides, and α-Darabinopyranosides were synthesized by Michael condensation of potassium salts of 7-hydroxy-4arylcoumarins with acetobromosugars followed by deacetylation of the resulting peracetates. Key words: coumarins, neoflavones, 4-arylcoumarins, glycosides, glycosylation. More than 130 neoflavones with the 4-phenylcoumarin structure have been isolated from natural sources [1, 2]. Among the natural neoflavones are O-glycosides with the carbohydrate at various positions of the coumarin. Examples of these are serratin 7-β-glucoside (1) isolated from Passiflora serratodigitata [3], 6-O-β-D-galactopyranosyldalbergin (2) isolated from Hesperethusa crenulata [4], and 5-O-(6′′-acetyl-β-D-glucopyranosyl)-7,3′,4′-trihydroxy-4-phenylcoumarin (3) produced by Hintonia latiflora [5]. The goal of our work was to synthesize O-D-glycopyranosides of 7-hydroxy-4-arylcoumarins. 7-Hydroxy-4-arylcoumarins 4-7 that were needed for further transformations were prepared in high yields by Pechmann condensation of ethylbenzoylacetate and ethyl-4-methoxybenzoylacetate with polyphenols (resorcinol and 2-methylresorcinol) in the presence of conc. H2SO4 [6, 7]. The O-glycopyranosides of the 7-hydroxy-4-arylcoumarins were synthesized using a method based on the reaction of a glycosyl donor and the potassium salts of the hydroxycoumarins in aqueous acetone with cooling (0°C) (modified Michael method [8, 9]) that was successfully used to synthesize a similar class of compounds [10]. The condensation of the potassium salts of 4-7 and acetobromosugars was carried out in concentrated aqueous acetone solution. Concentrated solutions of the phenolates of 4-7 were prepared using equivalent amounts of coumarins, KOH solution (10%), and twice (relative to the volume of base solution) the amount of acetone. The glycosyl donors in these syntheses were the D-acetobromoglycopyranoses α-acetobromoglucose (Ac4GlupBr), α-acetobromogalactose (Ac4GalpBr), β-acetobromoarabinose (Ac3ArapBr), and α-acetobromoxylose (Ac3XylpBr). The syntheses produced in 36-52% yields the O-peracetates of glucopyranosides 8-10, galactopyranosides 11-14, xylopyranoside 15, and arabinopyranoside 16, all of which have the sugar at the 7-position of the neoflavone. The structures of the prepared glycosides and the configurations of their anomeric centers were confirmed unambiguously by PMR spectroscopy. The PMR spectra of 8-16 contained signals for four (for glucosides and galactosides) or three (for arabinosides and xylosides) acetyl groups at 2.00-2.20 ppm and for the carbohydrates and aglycons. The PMR spectra of 8-10 exhibited a doublet at 5.12-5.18 ppm with SSCC J = 7.2-7.8 Hz for H-1 of the carbohydrate ring. A SSCC of this magnitude for H-1 and H-2 in carbohydrates is consistent with their trans-diaxial position in the ring [11]. Together with the chemical shift of H-1, this confirmed the β-configuration of the glucopyranosides. The CH2-6 methylene protons of the glucosides were chemically nonequivalent and formed together with H-5 a group of signals near 4 ppm. The methylene protons resonated as two doublets of doublets at 4.17-4.19 and 4.30-4.31 ppm with SSCC J = 12.0-12.4 and J = 2.02.4 and J = 12.0-12.4 and 5.6-6.0, respectively.

1) Taras Shevchenko Kiev National University, 01033, Ukraine, Kiev, ul. Vladimirskaya, 64; 2) Institute of Bioorganic Chemistry and Petrochemistry, National Academy of Sciences of Ukraine, 02094, Ukraine, Kiev, ul. Murmanskaya, 1, e-mail: [email protected]. Translated from Khimiya Prirodnykh Soedinenii, No. 6, pp. 546-550, November-December, 2005. Original article submitted October 3, 2005. 0009-3130/05/4106-0663 ©2005 Springer Science+Business Media, Inc.

663

R1 HO

R1

R1 O

AcO

O

O

O

HO

O

O

Ac4GlupBr, KOH

4-7

MeONa

OAc

H2O-CH3COCH3

Ac4GalpBr, KOH H2O-CH3COCH3

O

R

17 - 19

R1 O

OH

R

8 - 10

AcO AcO

HO

OAc

R

O

OH

MeOH

AcO

O

O

O

O

MeONa

O MeOH

OAc

R1

HO HO

O

O

O

O OH

OAc

OH

11 - 14

20 - 23

R

R

4, 8, 11, 17, 20: R = R1 = H; 5, 9, 12, 18, 21: R1 = Me, R = H; 6, 10, 13, 19, 22: R1 = H, R = OCH3 7, 14, 23: R1 = Me, R = OCH3 HO

O

O

O Ac3XylpBr, KOH AcO

H2O-CH3COCH3

AcO

O

OAc 16

OH

Ph

O

MeOH

24

O HO

MeONa O

Ph

HO

OAc 15

HO

O

OH

MeOH Ph

O

O

MeONa

O AcO

Ac3ArapBr, KOH

O

O

OAc

H2O-CH3COCH3

4

O

O

O

O

O

OH 25

Ph

PMR spectra of 11-14 contained a doublet for H-1 of the carbohydrate at 5.05-5.12 ppm with SSCC J = 7.8-8.1 Hz. This was consistent with the β-configuration of the prepared galactopyranosides. In contrast with the glucopyranosides, the CH2-6 methylene protons of the galactopyranosides appeared as a complicated multiplet at 4.05-4.25 ppm. A doublet for H-1 in the PMR spectrum of triacetylxyloside 15 at 5.26 ppm with SSCC J = 7.2 Hz is consistent with the β-configuration of the anomeric center. The CH2-5 methylene protons resonated as two doublet of doublets at 3.59 and 4.22 ppm with SSCC J = 12.0, J = 6.0 and J = 12.0, J = 2.1 Hz, respectively. The presence in the PMR spectrum of 16 of a doublet for H-1 at 5.20 ppm with SSCC J = 6.0 Hz confirmed that H-1 and H-2 were trans-diaxial. For a D-triacetylarabinoside, this occurs exclusively for the α-anomer, i.e., the arabinoside had the α-configuration. The CH2-5 methylene protons resonated as two doublets of doublets at 3.83 and 4.15 ppm with SSCC J = 12.0, J = 2.1 and J = 12.0, J = 6.0 Hz, respectively. The IR spectra of 8-16 contained two bands at 1690-1760 cm-1 that were characteristic of acetyl C=O and coumarin ring stretching vibrations, respectively. D-Glycopyranosides 17-25 with free hydroxyls were prepared in high yields by deacetylation of tetra- and tri-Oacetylglycopyranosides 8-16 using a modified Zemplen method (NaOMe in absolute MeOH). The PMR spectra of the synthesized glycosides contained signals for the carbohydrate and aglycon fragments and lacked signals for the acetyls in the starting peracetates. The presence in the PMR spectra of a doublet for anomeric H-1 with a typical SSCC confirmed that the 1,2-trans-diaxial orientation of H-1 and H-2 had been retained. The IR spectra of the glycopyranosides contained two bands at 3300-3400 and 1690-1720 cm-1 that were typical of hydroxyls and C=O of the coumarin ring, respectively.

664

EXPERIMENTAL The course of reactions and the purity of products were monitored by TLC on Merck 60 F254 plates with elution by CHCl3:CH3OH (9:1). Melting points were determined on a Kofler block. IR spectra were recorded on a Nicolet FTIR Nexus 475 spectrometer; PMR spectra, on Varian VXR-300 and Varian Mercury 400 spectrometers at 300 and 400 MHz, respectively, relative to TMS (internal standard). Elemental analyses agreed with those calculated. The syntheses of 7-hydroxy-4-arylcoumarins 4-7 have been published [6, 7]. Peracetylpyranosylbromides were prepared as before [12, 13]. 7-Peracetylglycopyranosyloxy-4-arylchromen-2-ones 8-16. A solution of 7-hydroxycoumarin (4-7, 10 mmol) in acetone (10 mL) and KOH solution (5.6 mL, 10%) was stirred vigorously and cooled (0°C) for 30 min, treated in portions with stirring over 1 h with the appropriate peracetylpyranosylbromide (10 mmol), stirred for 4 h with cooling (0°C), left overnight at room temperature, diluted with CHCl3 (50 mL), and worked up in a separatory funnel with KOH solution (1 N, 2 × 50 mL) and water (50 mL). Acidification of the combined alkaline extracts regenerated unreacted hydroxycoumarin. The organic layer was dried over anhydrous MgSO4 and evaporated in a rotary evaporator. The oily product was crystallized from propan-2-ol. 7-(2,3,4,6-Tetra-O-acetyl-β-D-glucopyranosyloxy)-4-phenylchromen-2-one (8), yield 42%, C29H28O12, mp 182183.5°C. IR spectrum (KBr, cm-1): 1756, 1696, 1612, 1568, 1444, 1376, 1220, 1072. PMR spectrum (400 MHz, CDCl3, δ, ppm, J/Hz): 2.05, 2.06, 2.07, 2.13 (12H, four s, four CH3COO), 3.93 (1H, m, H-5″), 4.19 (1H, dd, J = 12.4, J = 2.4, H-6″α), 4.31 (1H, dd, J = 12.4, J = 5.6, H-6″β), 5.17 (1H, d, J = 7.6, H-1″), 5.19 (1H, m, H-4″), 5.29-5.36 (2H, m, H-2″, H-3″), 6.28 (1H, s, H-3), 6.86 (1H, dd, J = 2.4, J = 8.8, H-6), 7.03 (1H, d, J = 2.4, H-8), 7.41 (1H, d, J = 8.8, H-5), 7.43 (2H, m, H-2′, H-6′), 7.53 (3H, m, H-3′, H-4′, H-5′). 7-(2,3,4,6-Tetra-O-acetyl-β-D-glucopyranosyloxy)-8-methyl-4-phenylchromen-2-one (9), yield 46%, C30H30O12, mp 168-169.5°C. IR spectrum (KBr, cm-1): 1756, 1692, 1606, 1452, 1374, 1248, 1040. PMR spectrum (300 MHz, CDCl3, δ, ppm, J/Hz): 2.05, 2.06, 2.08 (12H, three s, four CH3COO), 2.31 (3H, s, CH3-8), 3.87 (1H, m, H-5″), 4.17 (1H, dd, J = 12.0, J = 2.4, H-6″α), 4.30 (1H, dd, J = 12.0, J = 5.7, H-6″β), 5.12 (1H, d, J = 7.8, H-1″), 5.19 (1H, t, J = 9.6, H-4″), 5.30-5.40 (2H, m, H-2″, H-3″), 6.28 (1H, s, H-3), 6.91 (1H, dd, J = 8.7, H-6), 7.29 (1H, d, J = 8.7, H-5), 7.43 (2H, m, H-2′, H-6′), 7.50 (3H, m, H-3′, H-4′, H-5′). 7-(2,3,4,6-Tetra-O-acetyl-β-D-glucopyranosyloxy)-4-(4-methoxyphenyl)chromen-2-one (10), yield 49%, C30H30O13, mp 214.5-216°C. IR spectrum (KBr, cm-1): 1758, 1698, 1610, 1512, 1424, 1376, 1216, 1156, 1120, 1076. PMR spectrum (400 MHz, CDCl3, δ, ppm, J/Hz): 2.05, 2.07, 2.13 (12H, three s, four CH3COO), 3.89 (3H, s, CH3O-4′), 3.93 (1H, m, H-5″), 4.19 (1H, dd, J = 12.4, J = 2.0, H-6″α), 4.31 (1H, dd, J = 12.4, J = 6.0, H-6″β), 5.18 (1H, d, J = 7.2, H-1″), 5.19 (1H, m, H-4″), 5.29-5.36 (2H, m, H-2″, H-3″), 6.25 (1H, s, H-3), 6.86 (1H, dd, J = 2.4, J = 8.8, H-6), 7.01 (1H, d, J = 2.4, H-8), 7.04 (2H, d, J = 8.4, H-3′, H-5′), 7.39 (2H, d, J = 8.4, H-2′, H-6′), 7.48 (1H, d, J = 8.8, H-5). 7-(2,3,4,6-Tetra-O-acetyl-β-D-galactopyranosyloxy)-4-phenylchromen-2-one (11), yield 36%, C29H28O12, light yellow oil. PMR spectrum (300 MHz, CDCl3, δ, ppm, J/Hz): 2.06, 2.12, 2.18 (12H, three s, four CH3COO), 4.00-4.28 (3H, m, H-5″, CH2-6″), 5.05 (1H, d, J = 8.1, H-1″), 5.15 (1H, dd, J = 3.3, J = 8.1, H-3″), 5.45 (1H, d, J = 3.3, H-4″), 5.57 (1H, dd, J = 8.1, J = 8.1, H-2″), 6.28 (1H, s, H-3), 6.86 (1H, dd, J = 2.4, J = 8.7, H-6), 7.03 (1H, d, J = 2.4, H-8), 7.41 (1H, d, J = 8.7, H-5), 7.43 (2H, m, H-2′, H-6′), 7.53 (3H, m, H-3′, H-4′, H-5′). 7-(2,3,4,6-Tetra-O-acetyl-β-D-galactopyranosyloxy)-8-methyl-4-phenylchromen-2-one (12), yield 46%, C30H30O12, mp 187-188.5°C. IR spectrum (KBr, cm-1): 1758, 1696, 1604, 1444, 1370, 1222, 1070. PMR spectrum (300 MHz, CDCl3, δ, ppm, J/Hz): 2.04, 2.09, 2.21 (12H, three s, four CH3COO), 2.32 (3H, s, CH3-8), 4.05-4.25 (3H, m, H-5″, CH2-6″), 5.09 (1H, d, J = 8.1, H-1″), 5.15 (1H, dd, J = 3.3, J = 8.1, H-3″), 5.48 (1H, d, J = 3.3, H-4″), 5.60 (1H, dd, J = 8.1, J = 8.1, H-2″), 6.28 (1H, s, H-3), 6.92 (1H, d, J = 8.7, H-6), 7.29 (1H, d, J = 8.7, H-5), 7.43 (2H, m, H-2′, H-6′), 7.52 (3H, m, H-3′, H-4′, H-5′).

665

7-(2,3,4,6-Tetra-O-acetyl-β-D-galactopyranosyloxy)-4-(4-methoxyphenyl)chromen-2-one (13), yield 39%, C30H30O13, light yellow oil. PMR spectrum (300 MHz, CDCl3, δ, ppm, J/Hz): 2.04, 2.10, 2.21 (12H, three s, four CH3COO), 3.88 (3H, s, CH3O-4′), 4.00-4.25 (3H, m, H-5″, CH2-6″), 5.05 (1H, d, J = 8.1, H-1″), 5.15 (1H, dd, J = 3.3, J = 8.1, H-3″), 5.51 (1H, d, J = 3.6, H-4″), 5.62 (1H, dd, J = 8.1, J = 8.1, H-2″), 6.26 (1H, s, H-3), 6.86 (1H, dd, J = 2.4, J = 8.7, H-6), 7.01 (1H, d, J = 2.4, H-8), 7.04 (2H, d, J = 8.4, H-3′, H-5′), 7.39 (2H, d, J = 8.4, H-2′, H-6′), 7.48 (1H, d, J = 8.7, H-5). 7-(2,3,4,6-Tetra-O-acetyl-β-D-galactopyranosyloxy)-8-methyl-4-(4-methoxyphenyl)chromen-2-one (14), yield 52%, C31H32O13, mp 128-129.5°C. IR spectrum (KBr, cm-1): 1758, 1698, 1604, 1514, 1370, 1248, 1080. PMR spectrum (400 MHz, CDCl3, δ, ppm, J/Hz): 2.03, 2.05, 2.09, 2.21 (12H, four s, four CH3COO), 2.32 (3H, s, CH3-8), 3.89 (3H, s, CH3O-4′), 4.05-4.25 (3H, m, H-5″, CH2-6″), 5.08 (1H, d, J = 8.1, H-1″), 5.13 (1H, dd, J = 3.3, J = 8.1, H-3″), 5.48 (1H, d, J = 3.6, H-4″), 5.60 (1H, dd, J = 8.1, J = 8.1, H-2″), 6.25 (1H, s, H-3), 6.93 (1H, d, J = 8.7, H-6), 7.03 (2H, d, J = 8.7, H-3′, H-5′), 7.35 (1H, d, J = 8.7, H-5), 7.38 (2H, d, J = 8.7, H-2′, H-6′). 7-(2,3,4-Tri-O-acetyl-β-D-xylopyranosyloxy)-4-phenylchromen-2-one (15), yield 41%, C26H24O10, mp 184-185°C. IR spectrum (KBr, cm-1): 1758, 1692, 1610, 1428, 1374, 1250, 1220, 1120, 1074. PMR spectrum (300 MHz, CDCl3, δ, ppm, J/Hz): 2.11 (9H, s, CH3COO-2″, CH3COO-3″, CH3COO-4″), 3.59 (1H, dd, J = 12.0, J = 6.0, H-5″α), 4.22 (1H, dd, J = 12.0, J = 2.1, H-5″β), 5.04 (1H, m, H-4″), 5.17-5.32 (2H, m, H-2″, H-3″), 5.26 (1H, d, J = 7.2, H-1″), 6.27 (1H, s, H-3), 6.86 (1H, dd, J = 2.1, J = 8.7, H-6), 7.04 (1H, d, J = 2.1, H-8), 7.41 (1H, d, J = 8.7, H-5), 7.44 (2H, m, H-2′, H-6′), 7.52 (3H, m, H-3′, H-4′, H-5′). 7-(2,3,4-Tri-O-acetyl-α-D-arabinopyranosyloxy)-4-pheylchromen-2-one (16), yield 44%, C26H24O10, mp 185186.5°C. IR spectrum (KBr, cm-1): 1744, 1696, 1612, 1374, 1248, 1236, 1218, 1090, 1048. PMR spectrum (300 MHz, CDCl3, δ, ppm, J/Hz): 2.07, 2.09, 2.16 (9H, three s, four CH3COO), 3.83 (1H, dd, J = 12.0, J = 2.1, H-5″α), 4.15 (1H, dd, J = 12.0, J = 6.0, H-5″β), 5.17 (1H, m, H-4″), 5.20 (1H, d, J = 6.0, H-1″), 5.36 (1H, m, H-3″), 5.46 (1H, dd, J = 9.6, J = 10.2, H-2″), 6.27 (1H, s, H-3), 6.89 (1H, dd, J = 2.4, J = 8.7, H-6), 7.06 (1H, d, J = 2.4, H-8), 7.41 (1H, d, J = 8.7, H-5), 7.44 (2H, m, H-2′, H-6′), 7.53 (3H, m, H-3′, H-4′, H-5′). 7-Glycopyranosyloxy-4-arylchromen-2-ones 17-25. A solution of peracetate 8-16 (4 mmol) in absolute MeOH (20 mL) was treated with NaOMe (20 mg). The reaction mixture was boiled for 10-30 min (completion of reaction determined by TLC). The solid that precipitated on cooling (0°C) was filtered and washed with cold MeOH. 7-(β-D-Glucopyranosyloxy)-4-phenylchromen-2-one (17), yield 87%, C21H20O8, mp 199.5-201°C. IR spectrum (KBr, cm-1): 3436, 1704, 1686, 1620, 1552, 1384, 1292, 1166, 1076. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 3.18 (1H, m, H-4′), 3.26 (2H, m , H-2′, H-3′), 3.44 (2H, m, H-5′, H-6′α), 3.71 (1H, dd, J = 11.4, J = 2.4, H-6′β), 4.52 (1H, t, J = 5.7, OH-6), 4.96 (1H, d, J = 7.5, H-1′), 4.99 (1H, d, J = 4.2, OH), 5.04 (1H, d, J = 3.9, OH), 5.31 (1H, d, J = 4.2, OH), 6.21 (1H, s, H-3), 6.97 (1H, dd, J = 2.4, J = 8.7, H-6), 7.12 (1H, d, J = 2.4, H-8), 7.37 (1H, d, J = 8.7, H-5), 7.50 (2H, m, H-2′, H-6′), 7.54 (3H, m, H-3′, H-4′, H-5′). 7-(β-D-Glucopyranosyloxy)-8-methyl-4-phenylchromen-2-one (18), yield 94%, C22H22O8, mp 219-220.5°C. IR spectrum (KBr, cm-1): 3456, 1708, 1690, 1604, 1562, 1448, 1380, 1274, 1074, 1042. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 2.33 (3H, s, CH3-8), 3.15-3.38 (4H, m, H-2′, H-3′, H-4′, H-5′), 3.45 (1H, dd, J = 11.4, J = 4.8, H-6′α), 3.65 (1H, dd, J = 11.4, J = 2.4, H-6′β), 4.35 (1H, t, J = 5.7, OH-6), 4.89 (1H, d, J = 7.2, H-1′), 5.01 (1H, d, J = 4.2, OH), 5.12 (1H, d, J = 4.2, OH), 5.29 (1H, d, J = 4.2, OH), 6.19 (1H, s, H-3), 7.08 (1H, dd, J = 9.0, H-6), 7.23 (1H, d, J = 9.0, H-5), 7.49 (2H, m, H-2′, H-6′), 7.54 (3H, m, H-3′, H-4′, H-5′). 7-(β-D-Glucopyranosyloxy)-4-(4-methoxyphenyl)-chromen-2-one (19), yield 86%, C22H22O9, mp 195-196.5°C. IR spectrum (KBr, cm-1): 3420, 2940, 1722, 1706, 1692, 1606, 1510, 1464, 1376, 1254, 1072. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 3.17 (1H, m, H-4′), 3.27 (2H, m, H-2′, H-3′), 3.40 (1H, m, H-5′), 3.44 (1H, dd, J = 11.4, J = 4.8, H-6′α), 3.72 (1H, dd, J = 11.4, J = 2.4, H-6′β), 3.85 (3H, s, CH3O-4′), 4.47 (1H, t, J = 5.7, OH-6), 4.96 (1H, d, J = 7.5, H-1′), 4.98 (1H, d, J = 4.2, OH), 5.05 (1H, d, J = 3.9, OH), 5.31 (1H, d, J = 4.2, OH), 6.16 (1H, s, H-3), 6.97 (1H, dd, J = 2.4, J = 8.7, H-6), 7.08 (2H, d, J = 8.7, H-3′, H-5′), 7.10 (1H, d, J = 2.4, H-8), 7.44 (2H, d, J = 8.7, H-2′, H-6′), 7.46 (1H, d, J = 8.7, H-5).

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7-(β-D-Galactopyranosyloxy)-4-phenylchromen-2-one (20), yield 82%, C21H20O8, mp 233-234.5°C. IR spectrum (KBr, cm-1): 3380, 1704, 1690, 1616, 1380, 1294, 1166, 1080. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 3.40-3.75 (6H, m, H-2″, H-3″, H-4″, H-5″, CH2-6″), 4.49 (1H, d, J = 4.5, OH), 4.65 (1H, t, J = 5.7, OH-6), 4.90 (1H, d, J = 4.5, OH), 5.03 (1H, d, J = 7.5, H-1′), 5.30 (1H, d, J = 4.2, OH), 6.28 (1H, s, H-3), 7.00 (1H, dd, J = 2.1, J = 8.7, H-6), 7.16 (1H, d, J = 2.1, H-8), 7.37 (1H, d, J = 8.7, H-5), 7.50-7.60 (5H, m, H-2′, H-3′, H-4′, H-5′, H-6′). 7-(β-D-Galactopyranosyloxy)-8-methyl-4-phenylchromen-2-one (21), yield 89%, C22H22O8, mp 252-253.5°C. IR spectrum (KBr, cm-1): 3364, 1706, 1692, 1604, 1376, 1276, 1140, 1112, 1076. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 2.33 (3H, s, CH3-8), 3.35-3.75 (6H, m, H-2″, H-3″, H-4″, H-5″, CH2-6″), 4.40 (1H, d, J = 4.5, OH), 4.61 (1H, t, J = 5.7, OH-6), 4.68 (1H, d, J = 4.5, OH), 4.82 (1H, d, J = 7.5, H-1′), 5.15 (1H, d, J = 4.2, OH), 6.18 (1H, s, H-3), 7.08 (1H, dd, J = 9.0, H-6), 7.22 (1H, d, J = 9.0, H-5), 7.49 (2H, m, H-2′, H-6′), 7.54 (3H, m, H-3′, H-4′, H-5′). 7-(β-D-Galactopyranosyloxy)-4-(4-methoxyphenyl)chromen-2-one (22), yield 91%, C22H22O9, mp 235-236.5°C. IR spectrum (KBr, cm-1): 3396, 2912, 1706, 1690, 1608, 1548, 1510, 1380, 1250, 1212, 1088, 1032. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 3.38-3.74 (6H, m, H-2″, H-3″, H-4″, H-5″, CH2-6″), 3.85 (3H, s, CH3O-4′), 4.45 (1H, d, J = 4.5, OH), 4.58 (1H, t, J = 5.7, OH-6), 4.79 (1H, d, J = 4.5, OH), 4.93 (1H, d, J = 7.5, H-1′), 5.16 (1H, d, J = 4.2, OH), 6.15 (1H, s, H-3), 6.96 (1H, dd, J = 2.1, J = 8.7, H-6), 7.07 (2H, d, J = 8.7, H-3′, H-5′), 7.10 (1H, d, J = 2.1, H-8), 7.45 (3H, d, J = 8.7, H-5, H-2′, H-6′). 7-(β-D-Galactopyranosyloxy)-8-methyl-4-(4-methoxyphenyl)chromen-2-one (23), yield 93%, C23H24O9, mp 245246.5°C. IR spectrum (KBr, cm-1): 3420, 1704, 1690, 1608, 1510, 1372, 1284, 1254, 1072. PMR spectrum (300 MHz, DMSO-d6, δ, ppm, J/Hz): 2.33 (3H, s, CH3-8), 3.38-3.75 (6H, m, H-2″, H-3″, H-4″, H-5″, CH2-6″), 3.85 (3H, s, CH3O-4′), 4.45 (1H, d, J = 4.5, OH), 4.53 (1H, t, J = 5.7, OH-6), 4.75 (1H, d, J = 4.5, OH), 4.83 (1H, d, J = 7.8, H-1′), 5.18 (1H, d, J = 4.2, OH), 6.14 (1H, s, H-3), 7.07 (3H, d, J = 8.7, H-6, H-3′, H-5′), 7.30 (1H, d, J = 8.7, H-5), 7.44 (2H, d, J = 8.7, H-2′, H-6′). 7-(β-D-Xylopyranosyloxy)-4-phenylchromen-2-one (24), yield 87%, C20H18O7, mp 194-195.5°C. IR spectrum (KBr, cm-1): 3452, 1712, 1612, 1380, 1278, 1160, 1112, 1078. PMR spectrum (400 MHz, DMSO-d6, δ, ppm, J/Hz): 3.10-3.40 (4H, m, H-2′, H-3′, H-4′, H-5′a), 3.76 (1H, m, H-5′e), 4.95 (1H, d, J = 6.9, H-1′), 5.03 (1H, d, J = 4.5, OH), 5.08 (1H, d, J = 3.9, OH), 5.34 (1H, d, J = 4.8, OH), 6.28 (1H, s, H-3), 7.00 (1H, dd, J = 2.1, J = 8.7, H-6), 7.16 (1H, d, J = 2.1, H-8), 7.37 (1H, d, J = 8.7, H-5), 7.50-7.60 (5H, m, H-2′, H-3′, H-4′, H-5′, H-6′). 7-(α-D-Arabinopyranosyloxy)-4-phenylchromen-2-one (25), yield 82%, C20H18O7, mp 126-127.5°C. IR spectrum (KBr, cm-1): 3408, 1712, 1610, 1376, 1208, 1152, 1074. PMR spectrum (400 MHz, DMSO-d6, δ, ppm, J/Hz): 3.48 (1H, m, H-4′), 3.60-3.80 (4H, H-2′, H-3′, CH2-5′), 4.59 (1H, d, J = 4.2, OH), 4.77 (1H, d, J = 5.4, OH), 4.98 (1H, d, J = 6.9, H-1′), 5.22 (1H, d, J = 4.8, OH), 6.25 (1H, s, H-3), 7.01 (1H, dd, J = 2.1, J = 8.7, H-6), 7.15 (1H, d, J = 2.1, H-8), 7.38 (1H, d, J = 8.7, H-5), 7.50-7.60 (5H, m, H-2′, H-3′, H-4′, H-5′, H-6′).

ACKNOWLEDGMENT We thank OAO "Eksimed" (Kiev, Ukraine) for assistance with the work.

REFERENCES 1. 2. 3. 4.

R. D. H. Murray, The Naturally Occurring Coumarins, Springer, Vienna-New York (2002). M. M. Garazd, Ya. L. Garazd, and V. P. Khilya, Khim. Prir. Soedin., 47 (2003). A. Ulubelen, R. R. Kerr, and T. J. Mabry, Phytochemistry, 21, 1145 (1982). D. Kumar and D. K. Mukharya, Acta Cienc. Indica, Chem., 16C(4), 411 (1990); Chem. Abstr., 116, 170107w (1992).

667

5. 6. 7. 8. 9. 10. 11. 12. 13.

668

R. Mata, M. del R. Camacho, S. Mendoza, and M. del C. Cruz, Phytochemistry, 31, 3199 (1992). L. L. Woods and J. Sapp, J. Org. Chem., 27, 3703 (1962). M. M. Garazd, Ya. L. Garazd, A. S. Ogorodniichuk, and V. P. Khilya, Khim. Prir. Soedin., 428 (2002). V. G. Pivovarenko, V. P. Khilya, V. N. Kovalev, and S. A. Vasil′ev, Khim. Prir. Soedin., 511 (1988). V. G. Pivovarenko, V. P. Khilya, V. N. Kovalev, and S. A. Vasil′ev, Khim. Prir. Soedin., 519 (1988). M. M. Garazd, Ya. L. Garazd, and V. P. Khilya, Khim. Prir. Soedin., 7 (2004). J. F. Stoddart, Stereochemistry of Carbohydrates, Interscience, New York (1971). C. E. Redemann and C. Niemann, Organic Syntheses, Vol. 22, John Wiley & Sons, New York (1942), p. 1. N. K. Kochetkov, Methods of Carbohydrate Chemistry [in Russian], Mir, Moscow (1967).

Chemistry of Natural Compounds, Vol. 41, No. 6, 2005

METHYL JASMONATE INDUCE ENHANCED PRODUCTION OF SOLUBLE BIOPHENOLS IN PANAX GINSENG ADVENTITIOUS ROOTS FROM COMMERCIAL SCALE BIOREACTORS

G. Sivakumar1 and K. Y. Paek2

UDC 547.972

The contents of soluble biophenols such as protocatechuic, gentisic, vanillic, caffeic, syringic, p-coumaric, ferulic, salicylic, and cinnamic acids were screened and quantified from the adventitious roots of Panax ginseng by HPLC-MS. Control adventitious roots which showed the greatest accumulation of ferulic acid (0.09 mg/g DW) were observed in our experiments. An increase in the total soluble biophenol content of adventitious roots was observed 5 days after treatment with elicitor 200 µM/L methyl jasmonate (MeJa) i.e., 35 to 40 days of inoculation. Among the biophenols investigated, the salicylic acid content was higher (0.44 mg/g DW) in MeJa treated adventitious root culture in the bioreactor. MeJa might apparently increase soluble biophenols production in adventitious roots by enhancing the biosynthesis pathway from phenylalanine to salicylic acid and other simple biophenols. Key words: antioxidant, biophenols, bioreactors, ginseng, HPLC-MS. Panax ginseng C. A. Meyer (Araliaceae) has been used as a natural traditional medicine for thousands of years in Asian countries for its reputed anti-stress, and anti-aging properties and enhances memory power as well as sperm production in men. The cultivation of ginseng root is a long and laborious process and, as a result, it is an expensive commodity in the international market. Biophenols are present in three different forms: soluble, esterified, and insoluble, bound to the cell wall, in Korean ginseng [1]. Ginseng soluble biophenols have recently gained great attention for their antioxidant, anticancer, antidiabetic, anticarcinogenic, antimutagenic, and immunomodulating activity [2]. Due to these factors, pharmaceutical and healthcare professionals are increasingly considering complementary as well as alternative approaches for the commercial production of ginseng bioactive biophenols from disease-free and pesticide-residue-free roots. Modern bioreactor culture systems provide a more advanced technology to produce higher in vitro natural biophenols from P. ginseng root tissues using artificial nutrients with MeJa in an aseptic environment. The objective of the present study was to screen the soluble biophenolic profiles from bioreactor-derived ginseng adventitious roots and to increase production of soluble biophenols using MeJa. This is the first report on screening and enhanced production of biophenols using MeJa from bioreactor-derived P. ginseng adventitious roots. We previously reported the effects of various auxins and their combinations on callus induction and higher multiple adventitious root induction using the root explant of P. ginseng [3]. This paper presents an HPLC-MS screened soluble biophenol profile and enhanced soluble biophenol accumulation in P. ginseng adventitious roots using MeJa in bioreactor culture. By careful analysis of the chromatograms at different wavelengths in the scale of 200–800 nm, it was found that the chromatograms at 240 nm together with 280 nm could well represent the profile of the constituents. By comparing the chromatogram of control P. ginseng adventitious roots with those of its MeJa treatment extracts, the root derivation of each peak was confirmed in terms of the retention time and spectra achieved from the PDA detection. The identified soluble biophenol compounds are listed in Table 2. The presence of acetic acid in the mobile phases suppressed the dissociation of the biophenolic compounds and enhanced the selectivity of the elution system. Comparisons of chromatograms generated at different detection wavelengths support the use of 200–800 nm as a compromise for the detection of biophenols [4]. 1) Biotech Genomics, ENEA, Casaccia, Rome, Italy, e-mail: [email protected]; 2) Research Center for the Development of Advanced Horticultural Technology, Chungbuk National University, Cheongju 361-763, South Korea. Published in Khimiya Prirodnykh Soedinenii, No. 6, pp. 551-554, November-December, 2005. Original article submitted March 16, 2005. 0009-3130/05/4106-0669 ©2005 Springer Science+Business Media, Inc.

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TABLE 1. Effects of Methyl Jasmonate on Adventitious Root Growth and Biophenol Production of P. ginseng MeJa Con., µM/L

Biomass* fresh wt., g

Dry wt., g

% of dry wt.

Growth rate**

Total soluble biophenols*

Control 100 200 300 400

489.6 a 476.2 b 470.6 c 420.1 d 332.0 e

42.1 b 42.9 b 45.9 a 38.4 d 34.6 c

5.05 b 5.29 c 5.73 a

24.76 25.23 27.00 22.58 20.35

0.22 d 0.71 b 1.19 a 0.45 c 0.39 c

5.37 ab 6.10 d

______ Data was collected after 40 days of culture using a 5 L balloon type bioreactor having 4 L 3/4 MS medium. *Mean followed by different letters within a column are significantly different at P < 0.05 by Duncan’s multiple range test. **The values are the quotients of the dry weight after cultivation and the dry weight of the inoculum. Similar experiments were done 3 times.

TABLE 2. Soluble Biophenolic Profile of P. ginseng, Methyl Jasmonate (200 µM/L) Treated Adventitious Roots Biophenol profile (acid) Gallic Protocatechuic Gentisic Vanillic Caffeic Syringic p-Coumaric Ferulic Salicylic Cinnamic Total soluble biophenols

Retention time 3.15 5.34 8.90 10.21 11.16 12.10 12.21 15.33 17.51 23.74

Control adventitious root biophenols, mg/g DW

MeJa treated adventitious root biophenols, mg/g DW

Internal standard 0.01±0.01* 0.03±0.01 0.01±0.01 0.01±0.01 0.01±0.01 0.01±0.02 0.09±0.02 0.03±0.02 0.02±0.01 0.190

Internal standard 0.03±0.06* 0.15±0.01 0.11±0.05 0.04±0.02 0.07±0.02 0.02±0.02 0.27±0.09 0.44±0.09 0.09±0.05 1.21

______ Data was collected after 40 days of culture using a 5 L balloon type bioreactor having 4 L of 3/4 MS liquid medium. *Mean ± standard error of 3 replicates. A great advantage of mass spectrometry in terms of biophenols is that it provides prominent peaks for the molecular ions. The mass spectra of each soluble biophenol peak from P. ginseng adventitious roots show the presence of abundant [M–H]– ions at the appropriate m/z values, along with a unique retention time for each of the soluble biophenols. These chromatograms at m/z 170, 154, 154, 168, 180, 164, 194, 138, and 148 show the corresponding mass numbers of gallic acid (internal standard), protocatechuic, gentisic, vanillic, caffeic, syringic, p-coumaric, ferulic, salicylic, and cinnamic acids, respectively. We carefully studied the mass spectra of these compounds and compared them with standards and reference data [5]. ESI overcomes the lack of analyte volatility by the direct formation or emission of ions from the surface of the condensed phase. Deprotonated molecular ions represented the base peak in the negative ion spectra of all ten biophenolic species. Different concentrations of MeJa (100–400 µM/L) were added to adventitious root cultures of P. ginseng at culture day 35 after inoculation. Chaudry et al. [6] reported that after 48 h of jasmonate treatment the general decrease in protein synthesis is caused by the ribosome-inactivating protein JIP 60. These results were similar to our present report after treatment of MeJa resulted in inhibition of adventitious root growth but a higher accumulation of soluble biophenols (Table 1). In order to prevent a decreasing biomass of adventitious roots, the authors added MeJa 5 days before harvesting i.e., after 35 days of inoculation. The soluble biophenolic profile of MeJa-treated adventitious roots was dominated by salicylic, ferulic, and gentisic acids, with 670

lesser amounts of vanillic, cinnamic, syringic, caffeic and protocatechuic acids (Table 2). Most plant phenolic acids are derivatives of benzoic (C6–C1) or cinnamic (C6–C3) acids, and phenolic acids of low molecular weight occur widely in decomposing plant residues and in roots [7]. The production of soluble biophenols increased significantly compared to control (without MeJa), when up to 400 µM/L of MeJa was added. The optimum concentration of MeJa for the higher production of soluble biophenols was found to be 200 µM/L (Table 1), yielding a total soluble biophenol content of 1.21 mg/g DW. The maximum yield of salicylic acid was 0.44 mg/g DW when treated with 200 µM/L MeJa on the 40th day of inoculation (Table 2). The production of soluble biophenols did not change when MeJa was added at the zero day of inoculation but it inhibit the adventitious root biomass (data not shown). The time of addition of the MeJa had a significant effect on biophenol production compared to the control. These results also indicate that addition of MeJa at the early phase of P. ginseng adventitious root growth is not favored. Plata et al. [8] reported that the highest anthocyanin content was obtained after adding MeJa to sweet potato cell suspension cultures. Therefore it can be assumed that the presence of MeJa might be beneficial in triggering the expression of biophenol biosynthetic genes in the ginseng adventitious roots, which results in an increase in soluble biophenol production. Huang et al. [9] reported that ferulic acid was converted to vanillic acid and subsequently to protocatechuic acid by R. rubra under aerobic conditions. However, in the present study, salicylic acid accumulation was higher in MeJa-treated adventitious roots. Recent advances in root elicitor signalling pathway research have shown that roots are capable of differentially activating distinct defense pathways [10]. Depending on the type of elicitor encountered, the root appears to be capable of switching on the appropriate pathway or combination of pathways. The root signalling molecule salicylic acid plays an important role in this signalling network: blocking the response to either of these signals can render elicitor-treated roots more susceptible to stress [11]. We assume that higher accumulations of biophenols are associated with the elicitor of MeJa within ginseng adventitious root, which can enhance the activity of shikimic acid pathway enzymes. The authors observed the decreased biomass of ginseng adventitious roots when MeJa was added even after 35 days of inoculation. It may be, that MeJa can trigger the activation of proteinase inhibitor genes in P. ginseng. Kim et al. [12] identified a MeJa-responsive element in the promoter of a proteinase inhibitor II gene, indicating that this gene is a transcription ally regulated in response to MeJa. In conclusion, although it has been recognized that MeJa is an excellent elicitor for increasing secondary metabolite production, the mechanism of MeJa-induced biophenols biosynthesis is still not well understood. The results of MeJa induction on soluble biophenol production may be useful for exploring new strategies to improve salicylic acid production. The main finding derived from this work is that P. ginseng adventitious root can supply bioactive biophenols for pharmaceuticals. In addition, its use in Western medicine has also been growing rapidly due to the growing interest in alternative remedies using natural in vitro herbal medicines. Furthermore, the model system presented here seems to be a good soluble biophenol screening method for P. ginseng adventitious roots.

EXPERIMENTAL Induction of Callus. Panax ginseng C.A. Meyer was collected from the Mountain of KeumSan province in South Korea and the roots were selected as explants. They were surface-disinfected according to the Sivakumar and Krishnamurthy [13] method. Calluses were induced from root using Murashige and Skoog (MS) medium [14] supplemented with indolebutyric acid (IBA) in the range of 2.46–9.84 µM/L, 0.46–4.64 µM/L kinetin (Kin), and 3% sucrose, and solidified with 0.2% gelrite; subcultures were grown every 4 weeks. Adventitious roots were induced from 4-week-old well-grown, morphogenic callus cultures in 3/4 MS medium supplemented with 24.60 µM/L IBA and 0.2% gelrite under dark conditions. Adventitious roots were grown at a temperature of 20±2°C. The cultures were subcultured every 3 weeks. Bioreactor Culture of Adventitious Roots. The pH of the medium was adjusted to 6.0 before autoclaving at 121°C and 1.2 kg.cm-2 pressure for 40 min. The volume of input air in bioreactor was adjusted to 0.1 vvm (air volume/min). Bioreactors were maintained at 22±2°C in a dark room until harvest. 20 g fresh weight of adventitious roots was inoculated into 5 L balloon type bioreactors containing 4 L of the culture medium. Elicitor Treatment. After 35 days culture, filter sterilized methyl jasmonate (MeJa), 100–400 µM/L, was added to the 3/4 MS medium. After 5 days, harvested adventitious roots were used for biomass and soluble biophenol analysis.

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Determination of Root Biomass. Adventitious roots were separated from the medium by passing through a stainless steel sieve. Fresh weight was obtained after the roots were rinsed with tap water and the surface water blotted away. Dry weight was recorded after the roots were dried to constant weight at 50°C for two days. The adventitious root growth rate was calculated as: Growth rate = harvest dry weight (g)/inoculated dry weight (g). Extraction and Characterization of Soluble Biophenol Fraction. Freeze-dried P. ginseng adventitious roots, each sample containing 20 g, were separately pitted by blending and homogenized in methanol–acetone (1:1. 80 ml) saturated with sodium metadisulfite at medium speed in an Ultraturrax homogenizer (Janke & Kunkel, IKA–Labortechnik, Germany) at 0°C for 3 min and centrifuged at 5000 g for 20 min at 4°C [15]. The supernatant was separated and the pellet resuspended four times in the same solvent until a colorless solution was obtained. The combined supernatants were used for soluble biophenol (SBP) analysis. The combined supernatants were evaporated under vacuum at 35°C. The dry residue was resuspended with pH 2 water solution and centrifuged to separate a cloudy precipitate. The clear supernatant was extracted five times with hexane at a hexane to water ratio of 1:1 to remove free fatty acids and other lipidic contaminants. The SBPs were then extracted with ether/ethyl acetate (1:1) six times at a solvent to water ratio of 1:1. The ether–ethyl acetate extracts were dehydrated with anhydrous sodium sulfate, filtered, and evaporated to dryness under vacuum at 30°C. After evaporation to dryness, the residue was redissolved in LC-MS CHROMASOLV grade methanol (2 mL) in vials containing an internal standard (IS) of known amount, i.e., gallic acid, and filtered through a 0.45 µm millipore filter. HPLC. The soluble biophenol fraction was analyzed using a HPLC system (Waters 2690 separation module with autoinjector; Waters 996 photodiode array detector; Waters millennium 2010 chromatography manager, USA) on a C18 (25 cm × 4.6 mm i.d.) column; the eluates detected range from 200 to 800 nm. The extracted BP fractions (20 µL) were injected into HPLC column. Elution was run at a 0.2 ml/min flow rate using the following mobile phases: methanol (solvent A) and water/acetic acid, pH 3.3 (0.1%) (solvent B). The selected gradient started with 10% A 90% B; then solvent A was raised to 90% in 60 min and acquisition was stopped. The column was washed 10 min with 100% A and then kept at the initial conditions another 10 min. HPLC eluate was introduced into the ESI-MS interface. MS-ESI. MS detection was carried out using a Waters Micromass ZQ, USA. The system was monitored by a Compaq P720 Power PC computer equipped with MASSLYNXTM 4.0 software for instrument control and data acquisition, data reprocessing, and solute quantification. Nitrogen was employed as the nebulizing gas at a pressure of 60 p.s.i. The temperatures of the electrospray source and desolvation gas were 100 and 200°C, respectively. ESI mass spectra were acquired in the negative mode by scanning over the m/z range 100–1000 with unit mass resolution. Ions generated in the ion source were sampled into the mass analyzer by passing through a 25 µm i.d. orifice (voltage from 0 to 30 V) at the rear end of the atmospheric chamber. To prevent solvent vapors and contaminants from entering the vacuum chamber, we used a flow rate of 0.2 mL/min during all experiments, and 0.1 ml/min when the instrument was set in overnight standby.

ACKNOWLEDGMENT This work is financially supported by the Ministry of Education and Human Resources Development (MOE), the Ministry of Commerce, Industry and Energy (MOCIE) and the Ministry of Labor (MOLAB) through the fostering project of the Lab of Excellency as well as BioGreen 21 project RDA through Research Center for the Development of Advanced Horticultural Technology at Chungbuk National University, South Korea.

REFERENCES 1. 2. 3. 4. 5.

672

M. Y. Jung, B. S. Jeon, and J. Y Bock, Food Chem., 79, 105 (2003). D. D. Kitts, A. N. Wijewickreme, and C. Hu, Mol. Cell. Biochem., 203, 1 (2000). G. Sivakumar, K. W. Yu, and K. Y. Paek, Eng. Life. Sci., 5 (4), 333 (2005). S. Tian, K. Nakamura, T. Cui, and H. Kayahara, J. Chromatog. A, 1063, 121 (2005). J. Zhang, M. Cui, Y. He, H. Yu, and D. Guo, J. Pharm. Biomed. Anal., 36, 1029 (2005).

6. 7. 8. 9. 10. 11. 12. 13. 14. 15.

B. Chaudry, F. Muller-Uri, V. Cameron-Mills, S. Gough, D. Simpson, K. Skriver, and J. Mundy, Plant J., 6, 815 (1994). U. Blum, S. R. Shafer, and M. E. Lehman, Crit. Rev. Plant Sci., 18, 673 (1999). N. Plata, I. Konczak-Islam, S. Jayram, K. McClelland, T. Woolford, and P. Franks, Biochem. Eng. J., 14, 171 (2003). Z. Haung, L. Dostal, and J. P. N. Rosazza, Appl. Environ. Microbiol., 59, 2244 (1993). M. J. C. Pieterse and L. C. van Loon, Trends Plant Sci., 4, 52 (1999). T. Gaffney, L. Friedrich, B. Vernooij, D. Negrotto, G. Nye, S. Uknes, E. Ward, H. Kessmann, and J. Ryals, Science, 261, 754 (1993). S. R. Kim, J. L. Choi, M. A. Costa, and G. An, Plant Physiol., 99, 627 (1992). G. Sivakumar and K. V. Krishnamuthry, Russ. J. Plant. Physiol., 51, 713 (2004). T. Murashige and F. Skoog, Physiol. Plant., 15, 23 (1962). K. Krygier, F. Sosulski, and L. Hogge, J. Agr. Food Chem., 30, 330 (1982).

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