Ultrasound in fatty acid chemistry: synthesis of a 1-pyrroline fatty acid ester isomer from methyl ricinoleate

Lipase-catalyzed hydrolysis of TG containing acetylenic FA

Hydrolysis of symmetrical acetylenic TG of type AAA [viz., glycerol tri-(4-decynoate), glycerol tri-(6-octadecynoate), glycerol tri-(9-octadecynoate), glycerol tri-(10-undecynoate), and glycerol tri-(13-docosynoate)] in the presence of eight microbial lipases was studied. Novozyme 435 (Candida antarctica), an efficient enzyme for esterification, showed a significant resistance in the hydrolysis of glycerol tri-(9-octadecynoate) and glycerol tri-(13-docosynoate). Hydrolysis of acetylenic TG with Lipolase 100T (Humicola lanuginosa) was rapidly accomplished. Lipase PS-D (Pseudomonas cepacia) showed a fair resistance toward the hydrolysis of glycerol tri-(6-octadecynoate) only, which reflected its ability to recognize the delta6 positional isomer of 18:1. Lipase CCL (Candida cylindracea, syn. C. rugosa) and AY-30 (C. rugosa) were able to catalyze the release of 10-undecynoic acid and 9-octadecynoic acid from the corresponding TG, but less readily the 13-docosynoic acid in the case of glycerol tri-(13-docosynoate). The two lipases CCL and AY-30 were able to distinguish the small difference in structure of fatty acyl moieties in the TG substrate. To confirm this trend, three regioisomers of mixed acetylenic TG of type ABC (containing one each of delta6, delta9, and delta13 acetylenic FA in various positions) were prepared and hydrolyzed with CCL and AY-40. The results reconfirmed the observation that AY-30 and CCL were able to distinguish the slight differences in the molecular structure (position of the acetylenic bond and chain length) of the acyl groups in the TG during the hydrolysis of such TG substrates.

Novel azido fatty acid ester derivatives from conjugated C(18) enynoate

Methyl octadec-11Z-en-9-ynoate (1) was epoxidized to give methyl 11,12-Z-epoxy-octadec-9-ynoate (2, 81%). Acid catalyzed ring opening of the epoxy ring of compound 2 gave methyl 11,12-dihydroxy-octadec-9-ynoate (3, 80%). The latter was treated with mesyl chloride to yield methyl 11,12-dimesyloxy-octadec-9-ynoate (4, 76%). Reaction of compound 4 with sodium azide furnished methyl 11-azido-12-mesyloxy-octadec-9-ynoate (5a, 49%) and methyl 11-azido-octadec-11E-en-9-ynoate (5b, 24%). Compound 2 was semi-hydrogenated over Lindlar catalyst to give methyl 11,12-Z-epoxy-octadec-9Z-enoate (6, 90%). This allylic epoxy fatty ester (6) was reacted with sodium azide to give a mixture of methyl 11-azido-12-hydroxy-octadec-9Z-enoate (7a) and methyl 9-azido-12-hydroxy-octadec-9E-enoate (7b), which could not be separated into individual components by silica chromatography. Chromic acid oxidation of the mixture of compounds 7a and 7b furnished methyl 9-azido-12-oxo-octadec-10E-enoate (8, 42% based on amount of compound 6 used) and an intractable mixture of polar compounds. The various products were characterized by NMR spectroscopic and mass spectral analyses.

Cyclodehydration reactions of methyl 9,10-; 10,12-; and 9,1 2-dioxostearates with 1,2-diaminoethane under ultrasonic irradiation

Reaction of methyl 9,10-dioxostearate (1) and 9,12-dioxostearate (2) with 1,2-diaminoethane under concomitant ultrasonic irradiation (10-15 min, 60 degrees C) in water furnished the corresponding 2,3-dihydropyrazine (4, 79%) and 1,2,3,4-tetrahydro-1,4-diazocine (5, 70%) derivatives, respectively. Reaction of methyl 10,12-dioxostearate (3) with 1,2-diaminoethane was successful only when glacial acetic acid was used instead of water under ultrasonic irradiation (4 x 10 min, 70 degrees C) to give a 2,3-dihydro-1H-1,4-diazepine (6, 95%) derivative. The structures of these novel six-membered (4), seven-membered (6), and eight-membered (5) N-heterocyclic fatty ester derivatives were confirmed by a combination of infrared, nuclear magnetic resonance spectroscopic and mass spectral analyses.

Fullerene lipids: synthesis of C60 fullerene derivatives bearing a long-chain saturated or unsaturated triester system

Tris(hydroxymethyl)aminomethane was successfully esterified with saturated and unsaturated long-chain fatty acids. The resulting amino-triester intermediates were successively reacted with chloroacetyl chloride, sodium azide, and C60 fullerene. Spectral evidence showed that the aziridine ring is joined to the junction of 16,6]-fused rings of the fullerene. The structures of the various C60 fullerene derivatives bearing a long-chain saturated or unsaturated triester system were characterized by spectroscopic and spectrometric methods.

Lipase specificity toward some acetylenic and olefinic alcohols in the esterification of pentanoic and stearic acids

The esterification of five medium- and long-chain acetylenic alcohols (2-nonyn-1-ol, 10-undecyn-1-ol, 6-octadecyn-1-ol, 9-octadecyn-1-ol, and 13-docosyn-1-ol), seven olefinic alcohols (cis-3-nonen-1-ol, 10-undecen-1-ol, cis-6-octadecen-1-ol, cis-9-octadecen-1-ol, trans-9-octadecen-1-ol, trans-9, trans-11-octadecadien-1-ol, cis-9,cis-12-octadecadien-1-ol), and four short-chain unsaturated alcohols (allyl alcohol, 3-butyn-1-ol, 3-pentyn-1-ol, and cis-2-penten-1-ol) with pentanoic or stearic acid in the presence of various lipase preparations was studied. With the exception of 2-nonyn-1-ol, where Lipase AY-30 (Candida rugosa) was used as the biocatalyst, the esterification of C11, C18, and C22 acetylenic alcohols with pentanoic acid appeared to be generally unaffected by the presence of an acetylenic bond in the alcohol as relatively high yields of the corresponding esters (78-97%) were obtained. However, medium- and long-chain olefinic alcohols were discriminated by Lipase AY-30, Lipolase 100T (Rhizomucor miehei), and especially by porcine pancreatic lipase (PPL), when esterification was conducted with pentanoic acid. Esterification of medium- and long-chain acetylenic or olefinic alcohols with a long-chain fatty acid, stearic acid, was very efficient except when Lipase AY-30 and Lipolase 100T were used. Short-chain unsaturated alcohols were much more readily discriminated. 3-Pentyn-1-ol and 3-butyn-1-ol were difficult (<5% yield) to esterify with pentanoic or stearic acid in the presence of Lipase AY-30 and PPL, respectively. Very low yields (<26%) of esters were produced when 3-butyn-1-ol and 3-pentyn-1-ol were reacted with pentanoic or stearic acid, when catalyzed by lipase from Candida cylindracea. No reaction took place between 3-butyn-1-ol and stearic acids in the presence of Lipase AY-30. Esterification of short-chain acetylenic and olefinic alcohols was most efficiently achieved with Lipolase 100T (Rhizomucor miehei), Lipozyme IM20 (Rh. miehei), or Novozyme 435 (Candida antarctica) as the biocatalyst.

Fullerene lipids: synthesis of dialkyl 1,2-[6,6]-methano-[60]-fullerene dicarboxylate derivatives

Reaction of C60 fullerene with dialkyl bromomalonate (where the alkyl groups consist of short-, medium-, and long-saturated chains or unsaturated long chains) in the presence of sodium hydride gives [6,6]-bridged mono-adducts of methanofullerene. The spectroscopic properties of such fullerenoid lipids are reported.

Studies of lipase-catalyzed esterification reactions of some acetylenic fatty acids

Esterification of five positional isomers of acetylenic fatty acids [viz. 9:1(2a), 11:1(10a), 18:1(6a), 18:1(9a) and 22:1(13a)] of different chain lengths with n-butanol in n-hexane in the presence of eight different lipases [Lipozyme IM 20 (Rhizomucor miehei), Lipolase 100T (R. miehei), Novozyme 435 (Candida antarctica), PPL (porcine pancreatic lipase), CCL (C. cylindracea), PS-D (Pseudomonas cepacia), Lipase A-12 (Aspergillus niger) and Lipase AY-30 (C. rugosa)] was studied. 2-Nonynoic acid was not esterified except when catalyzed by the lipase from C. antarctica (Novozyme 435) to give 42% butyl ester after 48 h. The lipases from A. niger (Lipase A-12) and C. rugosa (Lipase AY-30) showed poor biocatalytic behavior in the esterification of the acetylenic fatty acids studied. 10-Undecynoic acid gave the highest conversion rate of esterification with each kind of lipase used. 6-Octadecynoic acid showed a marked degree of resistance to esterification carried out in the presence of C. cylindracea (CCL), P. cepacia (PS-D), or porcine pancreatic (PPL) lipase but not significantly in the presence of the lipases of R. miehei (Lipozyme IM 20), R. miehei (Lipolase 100T), or Novozyme 435. 9-Octadecynoic acid and 13-docosynoic acid were not discriminated and were readily esterified by the remaining six lipases, but when compared to oleic acid the acetylenic fatty acids were comparatively much slower in conversion to the esters.

Ultrasound-assisted synthesis of santalbic acid and a study of triacylglycerol species in Santalum album (Linn.) seed oil

Methyl ricinoleate (1) was treated with bromine and the dibromo derivative (2) was reacted with ethanolic KOH under ultrasonic irradiation to give 12-hydroxy-octadec-9-ynoic acid upon acidification with dil. HCI. The latter compound was methylated with BF3/methanol to give methyl 12-hydroxy-octadec-9-ynoate (3). Compound 3 was treated with methanesulfonyl chloride in the presence of triethylamine in CH2Cl2 to give methyl 12-mesyloxy-octadec-9-ynoate (4). Reaction of methyl 12-mesyloxy-octadec-9-ynoate with aqueous KOH under ultrasonic irradiation (20 kHz) gave (11E)-octadecen-9-ynoic acid (5, santalbic acid, 40%) and (11Z)-octadecen-9-ynoic acid (6, 60%) on acidification with dil. HCI. These isomers were separated by urea fractionation. The 13C nuclear magnetic resonance (NMR) spectroscopic properties of the methyl ester and the triacylglycerol (TAG) esters of these enynoic fatty acid isomers were studied. The carbon shifts of the unsaturated carbon nuclei of the methyl ester of the E-isomer were unambiguously assigned as 88.547 (C-9), 79.287 (C-10), 109.760 (C-11), and 143.450 (C-12) ppm, while the unsaturated carbon shifts of the (Z)-enynoate isomer appeared at 94.277 (C-9), 77.561 (C-10), 109.297 (C-11), and 142.668 (C-12) ppm. In the 13C NMR spectral analysis of the TAG molecules of type AAA containing either the (Z)- or (E)-enyne fatty acid, the C-1 to C-6 carbon atoms on the alpha- and beta-acyl positions were differentiated. The unsaturated carbon atoms in the alpha- and beta-acyl chains were also resolved into two signals except that of the C-11 olefinic carbon. Sandal (Santalum album) wood seed oil (a source of santalbic acid) was separated by silica chromatography into three fractions. The least polar fraction (7.2 wt%) contained TAG which had a random distribution of saturated and unsaturated fatty acids, of which oleic acid (69%) was the predominant component. The second fraction (3.8 wt%) contained santalbic acid (58%) and oleic acid (28%) together with some other normal fatty acids. Santalbic acid in this fraction was found in both the alpha- and beta-acyl positions of the glycerol "backbone." The most polar fraction (89 wt%) consisted of TAG containing santalbic acid only. The distribution of the various fatty acids on the glycerol "backbone" was supported by the results from the 13C NMR spectroscopic analysis.

Chemical and enzymatic preparation of acylglycerols containing C18 furanoid fatty acids

C18 furanoid triacylglycerol [glycerol tri-(9,12-epoxy-9,11-octadecadienoate)] was prepared by chemical transformation of triricinolein isolated from castor oil. The procedure involved oxidation, epoxidation and cyclization of the epoxy-keto intermediate with sodium azide and ammonium chloride in aqueous ethanol. The furanoid triacylglycerol was also obtained by esterification of C18 furanoid fatty acid with glycerol using Novozyme 435 (Novo Nordisk A.S., Bagsvaerd, Denmark) as biocatalyst. When Lipozyme (Novo Nordisk A.S.) was used, a mixture of the furanoid 1(3)-rac-monoacylglycerol and 1,3-diacylglycerol was obtained. In order to obtain the C18 furanoid 1,2(2,3)-diacylglycerol, selective hydrolysis of the furanoid triacylglycerol was achieved using porcine pancreatic lipase in tris(hydroxymethyl) methylamine buffer. Interesterification of triolein with methyl C18 furanoid ester in the presence of Lipozyme showed maximum incorporation of 34% of furanoid fatty acid. Extension of the interesterification to vegetable oils (olive, peanut, sunflower, corn and palm oil) allowed a maximum of 24% furanoid acid incorporation to be achieved.