Table of Contents Author Guidelines Submit a Manuscript
Journal of Lipids
Volume 2012, Article ID 476595, 13 pages
Research Article

Nonsterol Triterpenoids as Major Constituents of Olea europaea

1Institut de Biologie Moléculaire des Plantes du Centre National de la Recherche Scientifique (UPR 2357), Université de Strasbourg, 28 rue Goethe, 67083 Strasbourg, France
2Institut für Molekulare Physiologie und Biotechnologie der Pflanzen (IMBIO), Universität Bonn, Kirschallee 1, 53115 Bonn, Germany

Received 5 May 2011; Accepted 20 October 2011

Academic Editor: Angel Catala

Copyright © 2012 Naïm Stiti and Marie-Andrée Hartmann. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


Plant triterpenoids represent a large and structurally diverse class of natural products. A growing interest has been focused on triterpenoids over the past decade due to their beneficial effects on human health. We show here that these bioactive compounds are major constituents of several aerial parts (floral bud, leaf bud, stem, and leaf) of olive tree, a crop exploited so far almost exclusively for its fruit and oil. O. europaea callus cultures were analyzed as well. Twenty sterols and twenty-nine nonsteroidal tetra- and pentacyclic triterpenoids belonging to seven types of carbon skeletons (oleanane, ursane, lupane, taraxerane, taraxastane, euphane, and lanostane) were identified and quantified by GC and GC-MS as free and esterified compounds. The oleanane-type compounds, oleanolic acid and maslinic acid, were largely predominant in all the organs tested, whereas they are practically absent in olive oil. In floral buds, they represented as much as 2.7% of dry matter. In callus cultures, lanostane-type compounds were the most abundant triterpenoids. In all the tissues analyzed, free and esterified triterpene alcohols exhibited different distribution patterns of their carbon skeletons. Taken together, these data provide new insights into largely unknown triterpene secondary metabolism of Olea europaea.

1. Introduction

Plant triterpenoids, which include sterols, steroids, and brassinosteroids, constitute a large and structurally diverse group of natural products, with over 100 different carbon skeletons [1, 2]. Oxidative modifications and glycosylations generate more chemical diversity [3]. Sterols and nonsterol triterpenoids are synthesized via the cytoplasmic acetate/mevalonate pathway and share common biosynthetic precursors up to (3S)-2,3-oxidosqualene (OS). The conversion of OS to cycloartenol by the cycloartenol cyclase (CAS, EC is the first committed step in sterol biosynthesis, but OS can be also cyclized by distinct OS cyclases (OSCs), also known as triterpene synthases, into a variety of triterpene skeletons including those of α- and β-amyrins, the most commonly occurring plant triterpenes. These nonsterol triterpenoids are then metabolized into multioxygenated compounds, the precursors of triterpene saponins [4]. Thus, OS cyclization by the various triterpene synthases is a major branch point in the regulation of the carbon flux toward either the sterol pathway (primary metabolism) or the nonsterol triterpenoid pathway (secondary metabolism).

During these last ten years, triterpenoids isolated from a large number of plant organs from different species have been reported to exhibit a variety of antimicrobial, antioxidant, anti-inflammatory, antiviral, or antitumor-promoting biological activities [57]. Whereas the biological roles of sterols and brassinosteroids are well known [8, 9], the functions of nonsterol triterpenoids in planta still remain poorly understood. Nevertheless, there is increasing evidence that these “secondary” metabolites do contribute to plant defense, as attested by the production of triterpenic phytoalexins [10] or saponins [11] in response to biotic and abiotic stresses. Triterpenoids are known to be constituents of surface waxes on leaves and fruit of various species [1215] and consequently affect cuticular structure and water permeability. These compounds play important roles in plant-insect interactions. As an example, Guhling et al. have mentioned the contribution of lupeol to the formation of epicuticular wax crystals at the surface of Ricinus communis stems, making them slippery [16]. These crystals serve as a physical barrier, hampering insect adhesion and preventing herbivores from climbing vertical plant organs. Triterpenoids have been also reported to be major components of the insect-trapping glue of Roridula species [17].

Olive (Olea europaea L.) tree cultivation started 6000 years ago on the Mediterranean shores from where it has continued to spread throughout many countries. Olive oil has become increasingly popular due to its advantageous nutritional and medicinal properties. However, while the beneficial effects of olive oil monounsaturated fatty acids and phenols are well recognized, little attention has been paid to minor compounds, including triterpenoids. In a previous work, we reported the composition of sterols and nonsteroidal triterpenoids as free and esterified conjugates in olive fruit, along with the changes that occurred throughout ripening [18]. In particular, we showed that mature olive fruit contained substantial amounts of oleanolic acid and maslinic acid, two triterpene acids known to protect humans against several diseases [19]. However, as these compounds are mainly located in the surface waxes of the olive skin [20], they occur only at low amounts in the oil. The present study provides a detailed investigation of the triterpenoid profile of other aerial organs (floral bud, leaf bud, stem, young and mature leaves) that were harvested from the same O. europaea tree as olive fruit used previously [18]. Evidence is given for the occurrence of a vast array of nonsterol triterpenoids in all the tissues analyzed, with a predominance of oleanane-type triterpenoids. The content of free and esterified sterols in the different tissues was also determined. Finally, attention was also paid to sterols and triterpenoids in callus cultures, a potential readily available source of bioactive compounds.

2. Material and Methods

2.1. Plant Material and Callus Tissue Cultures

The various organs (olive fruit, floral buds, leaf buds, stems, young and mature leaves) were all collected from the same olive tree (Olea europaea L., cv chemlali), during the spring of 2005. This tree was grown in the “El Intilaka” olive grove, located in the area of Beni Khalled (Le Cap Bon, Tunisia). The quantities (in fresh wt) of collected organs were the following: 92 floral buds (140.8 mg), 125 leaf buds (132.6 mg), 67 young leaves (5.3 g), 84 mature leaves (16.9 g), and stems (23.4 g). The term “stem” was used to designate a small branch (diameter less than 1 cm) from which leaves were removed. We designated young leaves those with a length between 1.5 and 2 cm and mature leaves those with a length higher than 3.0 cm.

Olive callus cultures were initiated as follows. Seeds from the same cultivar were provided by Prof. N. Drira (Faculté des Sciences, Sfax, Tunisia) and allowed to germinate in the dark on a 1/2 Murashige and Skoog (MS) medium containing agarose (8 g/L), gibberellin A3 (2 mg/L), and sucrose (20 g/L). After 10 to 15 days, germinated seeds were transferred to 1/2 MS medium with agarose (8 mg/L) and sucrose (40 g/L) and grown at 24°C under a photoperiod of 16 h light/8 h dark for 12 weeks. Callus cultures were generated from an axenic leaf collected from seedlings grown in vitro. The leaf was wounded slightly with a scalpel and put on a 1/2 agar MS medium supplemented with sucrose (20 g/L) and 2,4-dichlorophenoxyacetic acid (0.1 mg/L). After 2 to 3 weeks, induced callus tissues were transferred to 1/3 agar MS medium containing sucrose (20 g/L), 6-Υ-Υ-(dimethylallylamino)-purine (0.1 mg/L) and 1-naphtalene acetic acid (0.5 mg/L) and grown at 25°C in the dark. Callus tissues were subcultured every 2 weeks onto a fresh supplemented MS medium.

2.2. Extraction and Isolation of Triterpenoids

Fresh Olea europaea samples were grounded in liquid nitrogen before being lyophilized. Free and esterified sterols and nonsteroidal triterpenoids were isolated from the various tissues as reported previously [18]. Briefly, sterols and nonsteroidal triterpenoids were extracted from dry material by refluxing at 65°C with dichloromethane/methanol (2 : 1 by vol). Free and esterified sterols were separated by TLC (Merck 60F254, 0.25 mm) and analyzed according to [21]. Free pentacyclic and tetracyclic triterpenes were recovered with the fraction of 4,4-dimethylsterols and triterpene diols, as a polar band located below 4-demethylsterols. Sterols and triterpenoids released from ester conjugates by hydrolysis with 6% KOH in methanol were purified as above. All compounds were acetylated with acetic anhydride/pyridine. Mono- and diacetate derivatives of sterols and nonsterol triterpenoids were quantified by gas chromatography (GC) using a Varian model 8300 chromatograph equipped with a FID and a DB-1 or DB-5 capillary column (25 m × 0.32 mm i.d., 0.25 μm thickness, J&W Scientific). The temperature program included a fast rise from 60 to 230°C (30°C/min), a slow rise from 230 to 280°C (2°C/min), and a plateau at 280°C for 10 min. Free cholesterol was used as an internal standard. Triterpene acids were isolated from the total lipid extract as described previously [18], converted into methyl esters by treatment with ethereal diazomethane at 0°C for 1 h, then acetylated to give acetoxy methylesters. Mono- and diacetoxy methylesters of nonsterol triterpenoids were analyzed by GC under similar conditions as above, but with oleanolic methylester as internal standard and a slightly modified temperature program (from 60 to 240°C at 40°C/min, then from 240 to 300°C at 2°C/min and a plateau at 300°C for 25 min). The amount of each compound was calculated as the ratio of the respective GC peak area to the internal standard (cholesterol or oleanolic methylester) peak area. Quantitative determinations correspond to means of three replicates ±SD.

Both sterol and nonsterol triterpenoids were identified by GC-mass spectrometry (MS) carried out on a 6890 Agilent gas chromatograph equipped with an on-column injector and a DB-5 (J&W Scientific) capillary column (30 m × 0.32 mm, i.d., 0.25 μm thickness) coupled to an Agilent 5973 mass selective detector using electron impact at 70 eV. Compounds were identified by their relative retention time (RRt) in GC and MS fragmentation pattern in GC-MS.

RRts were calculated in reference to that of the internal standard. Mass spectra were compared to those of available authentic samples or to literature data [18, 22, 23]. The structures of nonsterol triterpenoids described in the paper are shown in Scheme 1 and their MS fragmentation patterns are given in Table 1.

Table 1: GC-MS data of nonsterol triterpenoids.
Scheme 1: Structures of nonsterol triterpenoids cited in the paper.

3. Results

3.1. Free and Esterified Nonsterol Triterpenoids in Olea europaea Organs
3.1.1. Identification of Free Nonsterol Triterpenoids

Analysis of free triterpenoids in the different aerial parts of the olive tree (Table 2) shows the occurrence of 18 pentacyclic triterpenoids that are distributed within 5 families of carbon skeletons: (i) oleanane type: β-amyrin 1 (5α-olean-12-en-3β-ol), 28-nor-β-amyrin 2, erythrodiol 3 (5α-olean-12-ene-3β,28-diol), oleanolic acid 4 (3β-hydroxyolean-12-en-28-oic acid), and maslinic acid 5 (2α,3β-dihydroxy-olean-12-en-28-oic acid); (ii) ursane type: α-amyrin 6 (5α-urs-12-en-3β-ol), 28-nor-α-amyrin 7, uvaol 8 (5α-urs-12-ene-3β,28-diol), ursolic acid 9 (3β-hydroxyurs-12-en-28-oic acid), pomolic acid 10 (3β,19α-dihydroxyurs-12-en-28-oic acid) and 2-hydroxyursolic acid 11 (2α,3β-dihydroxyurs-12-en-28-oic acid); (iii) lupane type: lupeol 12 (5α-lup-20(29)-en-3β-ol), 3-epi-betulin 14 (5α-lup-20(29)-ene-3α,28-diol), and 3-epi-betulinic acid 15 (3α-hydroxy-5α-lup-20(29)-en-28-oic acid); (iv) taraxerane type: taraxerol 16 (taraxer-14-en-3β-ol) and myricadiol 17 (taraxer-14-ene-3β,28-diol or 28- taraxerol); (v) taraxastane type: Ψ-taraxasterol 19 (taraxast-20-en-3β-ol) and taraxasterol 20 (taraxast-20(30)-en-3β-ol). The structures of all these nonsterol triterpenoids are shown in Scheme 1. Among these triterpenoids, the compounds 10, 11, 19, and 20 had not been detected previously in the olive fruit [18]. Compounds 19 and 20 were identified as taraxastane triterpenes by their RRt in GC and their MS fragmentation patterns [22, 23]. Compounds 10 and 11 appeared to be ursane-oxygenated derivatives. Acetylation of the methylester derivative of 10 yielded a monoacetate [24] that was recovered with the fraction of methoxy triterpene acid diacetates. The MS spectrum showed fragment peaks at m/z 528 instead of 570 (molecular peak) and 510 [M-H2O]+, suggesting the presence of a tertiary OH group. Retro-Diels-Alder (RDA) cleavage fragments from rings D and E gave prominent diagnostic peaks at m/z 260 [278-H2O]+, 219 [278-CO2Me]+, 201 [278-H2O–CO2Me]+, indicating the presence of a carboxyl methyl group on the C-28 and of an hydroxyl group located on the ring E. A peak at m/z 249, arising from RDA cleavage of rings A and B, was observed. This fragmentation pattern is consistent with MS data reported by Hidaka et al. [25] for ilexgenin A, a C-24 carboxylic derivative of pomolic acid, suggesting that compound 10 is 3β,19α-dihydroxy-12-urs-en-28-oic acid or pomolic acid. This triterpene acid was identified recently in O. europaea cell suspension cultures [26]. The MS fragmentation pattern of compound 11 was similar to that of methyl maslinic acid diacetate 5, with abundant ions at m/z 262 and 203. These ion fragments usually result from the typical RDA cleavage of ring C of ursan-12-enes and olean-12-enes with a C-17 methoxycarbonyl and no hydroxyl groups on rings D/E [22]. However, in contrast to 5, compound 11 exhibited the most prominent ion at m/z 262 (base peak) instead of m/z 203. Furthermore, the appearance of a signal at m/z 249 and 189, resulting from successive losses of acetate from the ion at m/z 309, indicated that the additional oxygen atom was present on rings A/B. Compound 11 was identified as 2-hydroxyursolic acid. The stereochemistry at C-2 remained undetermined, but by analogy with maslinic acid (5), the corresponding oleanane triterpene acid possessing a 2α-OH, the structure of compound 11 as 2α,3β-dihydroxy ursolic acid or corosolic acid can be reasonably proposed. Moreover, this compound was also recently found in O. europaea cell suspensions [26].

Table 2: Amounts of free nonsterol triterpenoids in various organs of Olea europaea tree.

The representative compound of each group of pentacyclic triterpenoids appeared to be converted into more highly oxygenated compounds such as diols, mono- and dihydroxylated triterpene acids. Through several oxidations steps, the C-28 methyl group is converted sequentially into a hydroxymethyl and a carboxylic group, but introduction of hydroxyl groups at other positions of the pentacyclic carbon skeleton also seems to take place. The enzymes involved in these oxidation reactions have not been characterized, but are likely cytochrome P-450 monooxygenases [27, 28]. The first identification of a P450 involved in the 24-hydroxylation of two pentacyclic triterpenes, β-amyrin and sophoradiol, has been reported recently [29], but the absence of hydroxylation at the C-17 of these compounds suggests that distinct P450 oxidases might be needed to hydroxylate different carbon positions.

No triterpene acids with a taraxerane or taraxastane skeleton were found.

More polar compounds with additional hydroxyl (i.e., triols) or carboxyl groups such as rotundic acid, tormentic acid, or 19α-hydroxyasiatic acid have been recently identified from O. europaea cell suspensions [26]. These compounds that were not looked for might occur in the olive tree organs as well.

3.1.2. Quantification of Free Nonsterol Triterpenoids

Table 2 shows the distribution of free triterpenoids, as well as their content within the various parts of the olive tree. Oleanane-, ursane-, lupane-, and taraxerane-type pentacyclic triterpenoids were represented in all the organs, but taraxastane-type compounds, Ψ-taraxasterol 19 and taraxasterol 20, occurred only in leaves. Oleanane-derived triterpenoids corresponding to β-amyrin 1 and its oxygenated metabolites were by far the major nonsterol triterpenoids found. This pattern holds true especially for floral buds and mature olive fruit, where oleanolic acid 4 and maslinic acid 5 constituted up to 98-99% of total triterpenoids. Ursane-type triterpenoids were the second most abundant triterpenoids in the majority of organs. For instance, in young leaves they amounted to 29% of total triterpenoids, with a substantial contribution of uvaol 8. However in stems and floral buds, lupane-type triterpenoids were slightly more abundant than ursane-type triterpenoids. Taraxerane-type and taraxastane-type triterpenoids were minor compounds (less than 1%).

Floral and leaf buds were clearly the richest parts of the olive tree in terms of free triterpenoids with as much as 26.8 mg and 10.2 mg per g dry matter (2.7 and 1%, resp.), amounts consistent with those found in other triterpene-rich plant parts such as the plane bark or rosemary leaves [30]. In comparison, triterpenoids were far less abundant in stems (0.04% of dry wt). Olive fruit and leaves contained intermediate amounts (0.2 to 0.4%).

The wealth of triterpenoids in olive floral buds is consistent with literature data on the wide array of pentacyclic triterpenoids detected in flowers from various plant families [5, 3133]. Floral buds also contained a substantial amount of 3-epi-betulinic acid 15, far more than that in other organs.

A different distribution pattern of free triterpenoids was observed in leaf buds (Table 2). In particular, high levels in α- and β-amyrins were found, but only β-amyrin appeared to be significantly converted into triterpene acids, oleanolic acid 4 and maslinic acid 5. In young and mature leaves as well as in young olive fruit, substantial amounts of 28-nor-α-amyrin 7 and 28-nor-β-amyrin 2 were detected. These compounds lack a methyl group at C-17. As previously discussed [18], these two triterpenes might arise from the decarbonylation in planta of the aldehydes, 3β-hydroxy-5α-urs-12-en-28 al and 3β-hydroxy-5α-olean-12-en-28 al, which are the likely biosynthetic intermediates involved during the conversion of the C-28 hydroxymethyl group into a C-28 carboxylic group. Such a decarbonylation reaction on these unstable aldehydes could be promoted by exposure of leaves to intensive summer sunlight [34]. Occurrence of high levels of oleanolic and maslinic acids in olive leaves is well known [35, 36], but that of the ursane-type triterpenoids, pomolic acid 10 and 2-hydroxyursolic acid 11, has not been reported yet. Taken together, these data confirm the occurrence of pentacyclic triterpene acids in olive leaves as well as in leaves from various other plants [30, 37, 38].

Stems contained a low amount of nonsterol triterpenoids (Table 2). Although oleananes were the major nonsterol triterpenoids, this part of the olive tree was found to have the highest relative level in lupane derivatives, 12.2% of total free triterpenoids versus 0.5 to 1.9% in other organs. The main lupane compound was 3-epi-betulin 14. Lupeol 12 but not its 3-epimer 13 was detected. These data are consistent with the occurrence of lupane-type triterpenoids in tree bark [39, 40] and the predominance of trinorlupeol in Arabidopsis stems [14].

3.1.3. Changes in Relative Content of Triterpenoid Classes throughout Leaf Development

An interesting observation concerns changes throughout leaf development in the ratio of oleanane- to ursane-type triterpenoids following stepwise oxidations (Table 3 and Supplementary Figure S1 available online at doi: 10.1155/2012/476595). At the level of precursors, the ratio β-amyrin 1 to α-amyrin 6 was comprised between 0.7 and 1.0 for all leaf tissues, indicating the presence of substantial amounts of α-amyrin besides β-amyrin. At the level of diols, the value of the ratio erythrodiol 3 to uvaol 8 was 8.6-fold higher for leaf buds, but remained unchanged for both young and mature leaves. At the level of triterpene acids, a dramatic increase in the ratio (oleanolic acid 4 + maslinic acid 5) to (ursolic acid 9 + pomolic acid 10 + 2-hydroxyursolic acid 11) was observed, especially in mature leaves (a 150-fold increase compared to triterpene precursors and diols). If we consider the amounts of 28-nor-β-amyrin 2 and 28-nor-α-amyrin 7 as an estimate of the content in leaf tissues of the corresponding oleanane- and ursane-type aldehydes, the value found for the ratio 2/7 is intermediate between the corresponding values for diols and acids. These data emphasize clearly the progressive metabolization of β-amyrin into maslinic acid and its accumulation during leaf development, but also address the question of the fate of uvaol arising from α-amyrin oxidation. Given the very low amount of ursolic acid in mature leaves (Table 2), the diol uvaol might serve as a substrate for a specific glycosyltransferase involved in saponin biosynthesis. In oleanane-type saponins, the largest class of triterpene saponins, the C-17 position of the triterpene carbon skeleton is frequently occupied by a hydroxymethyl group [4]. A similar situation is likely to occur for ursane-type saponins. The presence of uvaol 8 after acid hydrolysis of a butanolic extract from young olive fruit, a process allowing recovery of triterpene aglycones from saponins, would support this hypothesis (N. Stiti, unpublished results). Surprisingly, at the best of our knowledge, no information about O. europaea triterpenoid saponins was available, although in Tunisia olive oil pomaces have been used for various domestic tasks such as linen washing or hand care for a very long time, indicating clearly their wealth in saponins (N. Stiti, personal remark).

Table 3: Ratio of oleanane- to ursane-type triterpenoids following oxidation steps of the C-28 CH3 throughout olive tree leaf development.
3.1.4. Identification and Quantification of Esterified Nonsterol Triterpenoids

The distribution of esterified triterpenoids within the different olive tree organs is shown in Table 4. Only triterpene alcohols were analyzed after their release from esters. Because of the nonavailability of standards, we did not look for 3-monoesters of triterpene diols. In all tissues, the same pentacyclic triterpenes, belonging to the five classes of carbon skeletons described previously, were found, but their amounts were much lower than those of the free forms. Moreover, their relative distribution among the various organs was completely different. Leaf buds were the richest organs in esterified pentacyclic triterpenes, with almost equal amounts of α-amyrin 6, β-amyrin 1, and lupeol 12. A clear predominance of oleanane compounds was observed only in floral buds. In leaves, both 3α and 3β-epimers of lupeol were detected, with a predominance of the 3α-epimer 13.

Table 4: Amounts of esterified triterpene alcohols in various organs from Olea europaea tree.

Leaves also possessed substantial amounts (60 to 66% of total esterified triterpenes) of tetracyclic triterpenes with euphane (butyrospermol 23) and lanostane (parkeol 24 and 24-methylene-dihydroparkeol 25) skeletons, parkeol being the major compound. These tetracyclic triterpenes had been found previously in olive fruit, but only between 21 and 30 weeks after flowering (WAF) [18]. Low amounts (1–3%) of taraxerol, taraxasterol, and Ψ-taraxasterol were detected in several organs (Table 4).

3.2. Free and Esterified Nonsterol Triterpenoids in Callus Cultures

Composition of free nonsterol triterpenoids in Olea europaea callus tissue cultures is shown in Table 5. Compared to organs of the olive tree, pentacyclic triterpenoids from these undifferentiated cells were of the same five classes of carbon skeletons (i.e., oleanane-, ursane-, lupane-, taraxerane-, and taraxastane-type), but were distributed differently. Moreover, higher amounts of tetracyclic triterpenes were present. Several new compounds were identified for both classes of triterpenoids.

Table 5: Amounts of free and esterified nonsterol triterpenoids in Olea europaea callus cultures.

The predominant pentacyclic triterpenoids remained the oleanane- and ursane-type triterpenoids, which were found at nearly equal amounts (Table 5), indicating that O. europaea undifferentiated cells produce more ursane-type triterpenoids than the differentiated cells. Callus cultures contained the same oleanane-type triterpenoids as those occurring in the olive tree samples, that is, β-amyrin 1, erythrodiol 3, oleanolic acid 4, and maslinic 5 acid, but also a new compound 21 in low amount (1.6%). The MS of 21 showed a fragmentation pattern typical of a Δ18-oleanene, with a prominent ion at m/z 203 (base peak) and similar to that of the diacetate of 5α-olean-18-ene-3β,28-diol or moradiol 21 [41]. Moradiol is a metabolite of germanicol, which has been found in olive oil [42].

Among ursane-type triterpenoids, we recovered α-amyrin 6, 28-nor-α-amyrin 7, uvaol 8, ursolic acid 9, 2-hydroxyursolic acid 11, and pomolic acid 10 (4.5%). The triterpene acids 9, 10 and 11 were also detected in Olea europaea cell suspensions [26]. We found traces of a new triterpene alcohol, with a MS fragmentation pattern very similar to that of bauerenol [22], but its relatively early RRt in GC with a DB5 column (1.326), close to that of α-amyrin (RRt: 1.335), was indicative of a compound with a Δ8-double bond rather than a Δ7-double bond. Thus, this compound was tentatively identified as isobauerenol 22.

The lupane-type triterpenoids from callus tissue cultures were represented by 3-epi-lupeol 13, 3-epi-betulin 14, and 3-epi-betulinic acid 15. Two other compounds, with late RRts in GC with a DB5 column (1.478 and 1.493, resp.), were recovered in the fraction of diacetates. The MS fragmentation patterns of both compounds were very similar to that of 3-epi-betulin. One of those compounds might correspond to betulin and the other to 3β,30-dihydroxylup-20(29)-ene (hennadiol), but these structures need to be confirmed.

Besides taraxer-14-ene-3β,28-diol 17, we found a second compound exhibiting an MS with fragmentations characteristic of the Δ14-taraxerene series of triterpenes [22] and very alike that of 28-hydroxytaraxerol (myricadiol) diacetate [23]. The presence of peaks at m/z 269, 202, and 189 indicated no additional group on cycles A, B, or C. Prominent ion fragments were observed at m/z 344 (base peak), 329, 284, and 269, corresponding to RDA fragmentation with collapse of ring D and successive losses of a methyl group, acetic acid and both groups. At the same time, the intensities of fragments at m/z 189 [262-CH2OAc]+ and 202 m/z [262-AcOH]+, which are characteristic for rings C and E, were decreased significantly. As reported by Budzikiewicz et al. [22], similar relative intensity changes were observed between erythrodiol diacetate and 30-hydroxy-β-amyrin diacetate MS fragmentation patterns, when the CH2OAc group is shifted from C-17 to C-30 (compare Figures 3 and 4 of [22], resp.). Thus, the position of the second hydroxyl group in this new taraxerene derivative might be located at C-29 or C-30. According to NMR and X-ray diffraction data from [43], the equatorial C-29 position for the CH2OAc would be favored and consequently the structure of 18 was established as taraxer-14-ene-3β,29-diol, a rare taraxerol derivative identified in the root bark of Hippocratea excelsa [43].

In contrast to olive tree organs in which free nonsteroidal tetracyclic triterpenes were not detected (Table 2), callus cultures contained substantial amounts of lanostane-type triterpenoids (34% of total free triterpenoids). By GC-MS, we identified parkeol 24 and 24-methylene-24-dihydroparkeol (25) (Table 1), but also 24-methylene lanost-8-en-3β-ol 26 and (24Z)-24-ethylidene-lanost-8-en-3β-ol 27 (Table 1, [44]). Parkeol was particularly abundant (Table 4). Two other lanostane triterpenoids with very late RRts in GC (1.478 and 1.493, resp. with a DB5-column) were also found. The first compound gave an intensive molecular ion at m/z 484, while the second one, with a MW of 482, showed a prominent fragment at m/z 341, corresponding to the loss of an unsaturated side chain plus 2H (Table 1). For both compounds, the absence of fragments at m/z 287 [M+-side chain-D ring] and m/z 227 [M+-side chain-D ring-acetate] was consistent with the absence of a 14α-methyl group. The two compounds displayed MS fragmentation patterns very similar to those of 24-ethyl- and 24-ethylidenelophenol but with a 14 mass increment of all ions fragments, suggesting that these two lanostanes can be ascribed to 4,4-dimethylsterols with a Δ7-double bond, 4,4-dimethylstigmast-7-en-3β-ol (28), and 4,4-dimethylstigmasta-7, Z-24(241)-dien-3β-ol (29), respectively.

Besides lanostane-type triterpenoids, callus cultures were also found to contain butyrospermol 23, an euphane-type triterpene.

It should be noticed that in these callus cultures, butyrospermol and parkeol were present as free forms, while in all the olive tree organs, both compounds were present only as fatty acid conjugates.

Callus cultures also contained esterified triterpenoids (Table 5). As in the case of olive tree organs, only esterified triterpenoid precursors were analyzed. We identified at least 24 compounds distributed between six families of carbon skeletons: oleanane, ursane, lupane, taraxastane, euphane, and lanostane. The major constituents were parkeol 24 and butyrospermol 23, two tetracyclic triterpenes representing together 76% of total esterified triterpenes. The other triterpenoids found were β-amyrin 1, 3-epi-lupeol 13, and α-amyrin 6.

3.3. Free and Esterified Sterols in the Various Organs of the Olive Tree

In all the olive tree organs analyzed, free sterols were present as end products as well as usual biosynthetic precursors, 4,4-dimethyl- (24-methylenecycloartanol and traces of cycloartenol) and 4α-methylsterols (obtusifoliol, cycloeucalenol, 24-methylene- and 24-ethylidene lophenol) (Supplementary Table S1). Sterol end products were largely predominant (97 to 99% of total free sterols).

All organs contained the same sterols, with sitosterol by far the major component (87 to 95% of total free sterols). This sterol profile was similar to that of the olive fruit [18] and olive oil [42, 45]. Flower buds, with rapidly dividing cells, were particularly rich in free sterols. However, the sterol content of floral buds was lower than that of mature fruit. Conversely, leaf buds appeared to have the lowest amount of free sterols among the organs sampled. Floral buds contained relatively higher contents of 24-methylcholesterol (6.6%) than the other olive organs (1 to 2.3%), suggesting active brassinosteroid synthesis in this organ [46].

Analysis of sterols recovered from ester conjugates showed that the various parts of the olive tree contained the same esterified sterols as the free forms (Supplementary Table S2). Sterol end products and their precursors, 4,4-dimethylsterols (24-methylene cycloartanol) and 4α-methylsterols (cycloeucalenol and 24-ethylidene lophenol), were found. Sitosterol was the major sterol (between 84 and 93% of total sterol esters). In all the tissues except leaf buds, esterified sterols were found in lower amounts than the free forms (Supplementary Tables S1 and S2).

3.4. Free and Esterified Sterols from Callus Cultures

Supplementary Table S3 indicates that undifferentiated olive cells synthesize the same sterol molecules as the olive tree organs. This similarity held true for both free and esterified sterols. Sitosterol remained the main sterol in callus. In the case of free sterols, substantial amounts of 4,4-dimethyl- and 4α-methylsterols were found, with sterol end products representing only 66% of total sterols. Such an accumulation of biosynthetic precursors may indicate a slow sterol metabolism in the callus tissue cultures compared to differentiated cells, although the total amount of sterols in these cultures is largely predominant compared to the sterol content of any olive tree organ (Supplementary Tables S1, S2 and S3). Moreover, in contrast to the situation in olive tree organs, nonsteroidal triterpenoids were not the major products of triterpene metabolism in callus cultures (Table 4 and supplementary Table S3).

4. Discussion

Both sterol and nonsteroidal pathways share common biosynthetic precursors up to oxidosqualene OS. Thus, the OS cyclization step appears as the site for a complex regulatory process between primary and secondary triterpene metabolisms. In Olea europaea fruit, we observed previously a preferential channeling of OS molecules towards one or the other pathway, depending on the stage of olive fruit development [18]. As expected, the present data indicate that all the parts of the olive tree contain significant amounts of free sterols, metabolites needed to sustain membrane biogenesis and plant growth [8, 9]. Besides sterols, O. europaea actively produces a vast array of nonsterol triterpenoids belonging to seven families of carbon skeletons, but oleanane-type triterpenoids predominate as in the majority of higher plants. Together with ursane-type compounds, oleanane-type triterpenoids comprised more than 97% of nonsterol triterpenoids in aerial parts of the O. europaea tree. In all the tissues analyzed, nonsterol triterpenoids were found at higher amounts than free sterols, suggesting that a large part of OS was dedicated to the nonsteroidal pathway, at least at the development stages of the various olive tree tissues we harvested. An opposite situation occurs in olive callus cultures, in which sterols were found to be 3-fold more abundant than nonsterol triterpenoids, represented mainly by lanostane-type and not oleanane-type triterpenoids (Table 4 and Supplementary Table S3). In that context, it would be interesting to investigate whether carbon flux toward these nonsterol triterpenoids could be upregulated following addition of an elicitor.

Arising from OS cyclization by the various OSCs, pentacyclic triterpenes are oxidized stepwise into triterpene diols and acids that accumulate. In olive fruit throughout ripening [18], we have shown that the main pathway leading to nonsterol triterpenoids starts with oxidation of the C-28 methyl group of α- and β-amyrine to give successively the corresponding dialcohols, uvaol and erythrodiol, and acids, ursolic acid and oleanolic acid, whereas introduction of additional hydroxyl groups at C-2 takes place later (Supplementary Figure S1). We assume that similar steps occur in other olive tree organs. O. europaea, oleanane-type triterpenoids are metabolized preferentially into triterpene acids compared to ursane-type compounds. Triterpene acids with a carboxyl group at C-28 might serve further as substrates for specific glycosyltransferases [28, 47, 48] to form triterpenoid saponins. As already pointed out, the biosynthetic pathway leading to triterpene saponins in O. europaea remains to be deciphered, but our data constitute a basis for further studies.

The occurrence of a vast array of nonsteroidal triterpenoids in O. europaea raises questions on the origin of such structural diversity. Many OSCs involved in OS cyclization are able to form simultaneously a large number of products, resulting from the stabilization of specific carbocationic intermediates. Other OSCs catalyze the formation of only one product. Thus, OSCs are often categorized as either “multifunctional” or accurate, but that concept has been questioned recently [49]. Arabidopsis can form as many as 35 triterpenes [49]. Although only two cyclases, CAS1 and LUP2, are sufficient to produce all the C30 triterpene alcohols that have been detected in Arabidopsis [50, 51], its genome encodes 13 homologs of triterpene synthases [50, 52]. Baruol synthase (BARS1) itself generates as many as 23 compounds, with baruol the predominant product [49].

Up to now, three Olea europaea OSCs have been isolated and characterized: OEW, a lupeol synthase [53], OEX, a cycloartenol synthase, and OEA, an amyrin synthase [26]. OEX catalyzes the first committed step involved in the sterol pathway, the cyclization of OS into cycloartenol, and is likely responsible for the constitutive synthesis of sterols expressed in all the parts of the olive tree. OEW is an accurate enzyme, able to form only lupeol, the precursor of lupane triterpenoids that were detected in all the olive tree organs, but were relatively more abundant in the stems (Table 2). OEA has been isolated from O. europaea cell suspension cultures initiated from leaf stalks [26]. When expressed in a yeast cell-free system, OEA was shown to be able to form not only α-amyrin 6, but also β-amyrin 1, butyrospermol 23, and Ψ-taraxasterol 19. The major product was by far α-amyrin (60%). This multifunctional OSC might be responsible for the synthesis of these four pentacyclic triterpenes in olive tree leaves. The three isolated OSCs might not be sufficient to form all the triterpenoids detected in O. europaea. The large predominance of oleanane-type triterpenoids suggests that a β-amyrin synthase might be involved. We isolated two full-length cDNAs encoding two proteins comprising 760 amino acids and sharing 91% identity. These two proteins exhibited 86% identity with other plant β-amyrin synthases. We are characterizing the function of both genes in a lanosterol synthase-deficient Saccharomyces cerevisiae strain. The occurrence of a substantial amount of lanostane-type triterpenoids in callus tissue cultures also suggests the potential involvement of a lanosterol synthase in O. europaea, as already found in other plants [51, 54].

In conclusion, evidence is given here for the occurrence in all the parts of the Olea europaea tree analyzed of significant amounts of oleanane-type triterpenoids, among which oleanolic and maslinic acids are largely predominant. Although further work is needed to investigate their roles in planta, these two compounds already appear to be promising for their valuable effects on glucose and lipid metabolism as well as for their antimicrobial, antiviral, and antioxidant activities [19, 55]. In this context, because of their high content in both these triterpenic acids, waste olive pomaces as well as leaves collected with mechanically harvested fruit constitute potential sources for isolating these bioactive compounds and thus might contribute to better valorize the Olea europaea tree.


CAS:Cycloartenol synthase
OSC:Oxidosqualene cyclase
WAF:Weeks after flowering.


The authors thank Professor Noureddine Drira and Dr. Mohamed Maâlej (Laboratoire des Biotechnologies Végétales Appliquées à l’Amélioration des Cultures, Sfax, Tunisia) for providing them the Olea europaea seeds and protocols to generate callus cultures. They are also grateful to Annie Hoeft for assistance with GC-MS analyses. Professor J. H. Weil and Professor Saida Triki are acknowledged for their encouragements to this work, which is a part of N. Stiti’s Ph.D. thesis. This work was supported by the Centre National de la Recherche Scientifique (CNRS) and a Wood-Whelan Research Fellowship from the International Union of Biochemistry and Molecular Biology (IUBMB).


  1. S. B. Mahato, A. K. Nandy, and G. Roy, “Triterpenoids,” Phytochemistry, vol. 31, no. 7, pp. 2199–2249, 1992. View at Google Scholar · View at Scopus
  2. R. Xu, G. C. Fazio, and S. P. Matsuda, “On the origins of triterpenoid skeletal diversity,” Phytochemistry, vol. 65, no. 3, pp. 261–291, 2004. View at Publisher · View at Google Scholar · View at Scopus
  3. J. D. Connolly and R. A. Hill, “Triterpenoids,” Natural Product Reports, vol. 27, no. 1, pp. 79–132, 2010. View at Google Scholar · View at Scopus
  4. J. P. Vincken, L. Heng, A. de Groot, and H. Gruppen, “Saponins, classification and occurrence in the plant kingdom,” Phytochemistry, vol. 68, no. 3, pp. 275–297, 2007. View at Publisher · View at Google Scholar · View at Scopus
  5. T. Akihisa, K. Yasukawa, and Y. Kasahara, “Triterpenoids from the flowers of Compositae and their anti-inflammatory effects,” Current Topics in Phytochemistry, vol. 1, pp. 137–144, 1997. View at Google Scholar
  6. G. Topçu, “Bioactive triterpenoids from Salvia species,” Journal of Natural Products, vol. 69, no. 3, pp. 482–487, 2006. View at Publisher · View at Google Scholar · View at Scopus
  7. P. Dzubak, M. Hajduch, D. Vydra et al., “Pharmacological activities of natural triterpenoids and their therapeutic implications,” Natural Product Reports, vol. 23, no. 3, pp. 394–411, 2006. View at Publisher · View at Google Scholar · View at Scopus
  8. M. A. Hartmann, “Plant sterols and the membrane environment,” Trends in Plant Science, vol. 3, no. 5, pp. 170–175, 1998. View at Publisher · View at Google Scholar · View at Scopus
  9. H. Schaller, “The role of sterols in plant growth and development,” Progress in Lipid Research, vol. 42, no. 3, pp. 163–175, 2003. View at Publisher · View at Google Scholar · View at Scopus
  10. R. Van der Heijden, D. R. Threlfall, R. Verpoorte, and I. M. Whitehead, “Regulation and enzymology of pentacyclic triterpenoid phytoalexin biosynthesis in cell suspension cultures of Tabernaemontana divaricata,” Phytochemistry, vol. 28, no. 11, pp. 2981–2988, 1989. View at Google Scholar · View at Scopus
  11. K. Papadopoulou, R. E. Melton, M. Leggett, M. J. Daniels, and A. E. Osbourn, “Compromised disease resistance in saponin-deficient plants,” Proceedings of the National Academy of Sciences of the United States of America, vol. 96, no. 22, pp. 12923–12928, 1999. View at Publisher · View at Google Scholar · View at Scopus
  12. S. Bauer, E. Schulte, and H. P. Thier, “Composition of the surface wax from tomatoes,” European Food Research and Technology, vol. 219, no. 3, pp. 223–228, 2004. View at Google Scholar
  13. S. Mintz-Oron, T. Mandel, I. Rogachev et al., “Gene expression and metabolism in tomato fruit surface tissues,” Plant Physiology, vol. 147, no. 2, pp. 823–851, 2008. View at Publisher · View at Google Scholar · View at Scopus
  14. H. Shan, W. K. Wilson, D. R. Phillips, B. Bartel, and S. P. Matsuda, “Trinorlupeol: a major nonsterol triterpenoid in Arabidopsis,” Organic Letters, vol. 10, no. 10, pp. 1897–1900, 2008. View at Publisher · View at Google Scholar · View at Scopus
  15. C. Van Maarseveen and R. Jetter, “Composition of the epicuticular and intracuticular wax layers on Kalanchoe daigremontiana (Hamet et Perr. de la Bathie) leaves,” Phytochemistry, vol. 70, no. 7, pp. 899–906, 2009. View at Publisher · View at Google Scholar · View at Scopus
  16. O. Guhling, B. Hobl, T. Yeats, and R. Jetter, “Cloning and characterization of a lupeol synthase involved in the synthesis of epicuticular wax crystals on stem and hypocotyl surfaces of Ricinus communis,” Archives of Biochemistry and Biophysics, vol. 448, no. 1-2, pp. 60–72, 2006. View at Publisher · View at Google Scholar · View at Scopus
  17. B. R. Simoneit, P. M. Medeiros, and E. Wollenweber, “Triterpenoids as major components of the insect-trapping glue of Roridula species,” Zeitschrift für Naturforschung C, vol. 63, no. 9-10, pp. 625–630, 2008. View at Google Scholar · View at Scopus
  18. N. Stiti, S. Triki, and M. A. Hartmann, “Formation of triterpenoids throughout Olea europaea fruit ontogeny,” Lipids, vol. 42, no. 1, pp. 55–67, 2007. View at Publisher · View at Google Scholar · View at Scopus
  19. J. Liu, R. Rajendram, and L. Zhang, “Effects of oleanolic acid and maslinic acid on glucose and lipid metabolism: implications for the beneficial effects of olive oil on health,” in Olives and Olive Oil in Health and Disease Prevention, V. R. Preedy and R. R. Watson, Eds., pp. 1423–1429, Oxford Academic, Oxford, UK, 2010. View at Google Scholar
  20. G. Bianchi, C. Murelli, and G. Vlahov, “Surface waxes from olive fruits,” Phytochemistry, vol. 31, no. 10, pp. 3503–3506, 1992. View at Google Scholar · View at Scopus
  21. M. A. Hartmann and P. Benveniste, “Plant membrane sterols: isolation, identification, and biosynthesis,” Methods in Enzymology, vol. 148, pp. 632–650, 1987. View at Google Scholar
  22. H. Budzikiewicz, J. M. Wilson, and C. Djerassi, “Mass spectrometry in structural and stereochemical problems. XXXII. Pentacyclic triterpenes,” Journal of the American Chemical Society, vol. 85, no. 22, pp. 3688–3699, 1963. View at Google Scholar · View at Scopus
  23. K. Shoijima, Y. Arai, K. masuda, Y. Takase, T. Ageta, and H. Ageta, “Mass spectra of pentacyclic triterpenoids,” Chemical and Pharmaceutical Bulletin, vol. 40, no. 7, pp. 1683–1690, 1992. View at Google Scholar · View at Scopus
  24. S. Ngouela, B. Nyasse, E. Tsamo, B. L. Sondengam, and J. D. Connolly, “Spathodic acid: a triterpene acid from the stem bark of Spathodea campanulata,” Phytochemistry, vol. 29, no. 12, pp. 3959–3961, 1990. View at Google Scholar · View at Scopus
  25. K. Hidaka, M. Ito, Y. Matsuda, H. Kohda, K. Yamasaki, and J. Yamahara, “A triterpene and saponin from roots of Ilex pubescens,” Phytochemistry, vol. 26, no. 7, pp. 2023–2027, 1987. View at Google Scholar · View at Scopus
  26. H. Saimaru, Y. Orihara, P. Tansakul, Y. H. Kang, M. Shibuya, and Y. Ebizuka, “Production of triterpene acids by cell suspension cultures of Olea europaea,” Chemical and Pharmaceutical Bulletin, vol. 55, no. 5, pp. 784–788, 2007. View at Publisher · View at Google Scholar · View at Scopus
  27. J. Ehlting, V. Sauveplane, A. Olry, J. F. Ginglinger, N. J. Provart, and D. Werck-Reichhart, “An extensive (co-)expression analysis tool for the cytochrome P450 superfamily in Arabidopsis thaliana,” BMC Plant Biology, vol. 8, pp. 47–65, 2008. View at Google Scholar
  28. M. A. Naoumkina, L. V. Modolo, D. V. Huhman et al., “Genomic and coexpression analyses predict multiple genes involved in triterpene saponin biosynthesis in Medicago truncatula,” The Plant Cell, vol. 22, no. 3, pp. 850–866, 2010. View at Publisher · View at Google Scholar · View at Scopus
  29. M. Shibuya, M. Hoshino, Y. Katsube, H. Hayashi, T. Kushiro, and Y. Ebizuka, “Identification of β-amyrin and sophoradiol 24-hydroxylase by expressed sequence tag mining and functional expression assay,” FEBS Journal, vol. 273, no. 5, pp. 948–959, 2006. View at Publisher · View at Google Scholar · View at Scopus
  30. S. Jäger, H. Trojan, T. Kopp, M. N. Laszczyk, and A. Scheffler, “Pentacyclic triterpene distribution in various plants—rich sources for a new group of multi-potent plant extracts,” Molecules, vol. 14, no. 6, pp. 2016–2031, 2009. View at Publisher · View at Google Scholar · View at Scopus
  31. T. Akihisa, K. Yasukawa, H. Oinuma et al., “Triterpene alcohols from the flowers of compositae and their anti- inflammatory effects,” Phytochemistry, vol. 43, no. 6, pp. 1255–1260, 1996. View at Publisher · View at Google Scholar · View at Scopus
  32. M. Ukiya, T. Akihisa, K. Yasukawa et al., “Constituents of compositae plants. 2. Triterpene diols, triols, and their 3-O-fatty acid esters from edible chrysanthemum flower extract and their anti-inflammatory effects,” Journal of Agricultural and Food Chemistry, vol. 49, no. 7, pp. 3187–3197, 2001. View at Publisher · View at Google Scholar · View at Scopus
  33. T. Akihisa, S. G. Franzblau, M. Ukiya et al., “Antitubercular activity of triterpenoids from asteraceae flowers,” Biological and Pharmaceutical Bulletin, vol. 28, no. 1, pp. 158–160, 2005. View at Publisher · View at Google Scholar · View at Scopus
  34. R. K. Hota and M. Bapuji, “Triterpenoids from the resin of Shorea Robusta,” Phytochemistry, vol. 35, no. 4, pp. 1073–1074, 1994. View at Publisher · View at Google Scholar · View at Scopus
  35. G. Bianchi, G. Vlahov, C. Anglani, and C. Murelli, “Epicuticular wax of olive leaves,” Phytochemistry, vol. 32, no. 1, pp. 49–52, 1993. View at Google Scholar
  36. N. Sánchez-Ávila, F. Priego-Capote, J. Ruiz-Jiménez, and M. D. Luque de Castro, “Fast and selective determination of triterpenic compounds in olive leaves by liquid chromatography-tandem mass spectrometry with multiple reaction monitoring after microwave-assisted extraction,” Talanta, vol. 78, no. 1, pp. 40–48, 2009. View at Publisher · View at Google Scholar · View at Scopus
  37. N. Banno, T. Akihisa, H. Tokuda et al., “Triterpene acids from the leaves of Perilla frutescens and their anti-inflammatory and antitumor-promoting effects,” Bioscience, Biotechnology and Biochemistry, vol. 68, no. 1, pp. 85–90, 2004. View at Publisher · View at Google Scholar · View at Scopus
  38. N. Banno, T. Akihisa, H. Tokuda et al., “Anti-inflammatory and antitumor-promoting effects of the triterpene acids from the leaves of Eriobotrya japonica,” Biological and Pharmaceutical Bulletin, vol. 28, no. 10, pp. 1995–1999, 2005. View at Publisher · View at Google Scholar · View at Scopus
  39. B. J. W. Cole, M. D. Bentley, and Y. Hua, “Triterpenoid extractives in the outer bark of Betula lenta (Black birch),” Holzforschung, vol. 45, no. 4, pp. 265–268, 1991. View at Google Scholar
  40. I. Habiyaremye, T. Stevanovic-Janezic, B. Riedl, F. X. Garneau, and F. I. Jean, “Pentacyclic triterpene constituents of yellow birch bark from Quebec,” Journal of Wood Chemistry and Technology, vol. 22, no. 2-3, pp. 83–91, 2002. View at Publisher · View at Google Scholar · View at Scopus
  41. D. Abramson, L. J. Goad, and T. W. Goodwin, “Triterpenes and sterols of Buxus sempervirens and local variations in their levels,” Phytochemistry, vol. 12, no. 9, pp. 2211–2216, 1973. View at Google Scholar · View at Scopus
  42. T. Itoh, K. Yoshida, T. Yatsu, T. Tamura, and T. Matsumoto, “Triterpene alcohols and sterols of Spanish olive oil,” Journal of the American Oil Chemists Society, vol. 58, no. 4, pp. 545–550, 1981. View at Publisher · View at Google Scholar · View at Scopus
  43. A. R. Aguilar-Gonzalez, G. J. Mena-Rejón, N. Padilla-Montaño, A. Toscano, and L. Quijano, “Triterpenoids from Hippocratea excelsa. The crystal structure of 29-hydroxytaraxerol,” Zeitschrift für Naturforschung B, vol. 60, no. 5, pp. 577–584, 2005. View at Google Scholar · View at Scopus
  44. P. Bouvier-Navé, T. Husselstein, T. Desprez, and P. Benveniste, “Identification of cDNAs encoding sterol methyl-transferases involved in the second methylation step of plant sterol biosynthesis,” European Journal of Biochemistry, vol. 246, no. 2, pp. 518–529, 1997. View at Google Scholar · View at Scopus
  45. S. Azadmard-Damirchi, G. P. Savage, and P. C. Dutta, “Sterol fractions in hazelnut and virgin olive oils and 4,4′- dimethylsterols as possible markers for detection of adulteration of virgin olive oil,” Journal of the American Oil Chemists' Society, vol. 82, no. 10, pp. 717–725, 2005. View at Publisher · View at Google Scholar · View at Scopus
  46. A. Sakurai and S. Fujioka, “Studies on biosynthesis of brassinosteroids,” Bioscience, Biotechnology and Biochemistry, vol. 61, no. 5, pp. 757–762, 1997. View at Google Scholar · View at Scopus
  47. L. Achnine, D. V. Huhman, M. A. Farag, L. W. Sumner, J. W. Blount, and R. A. Dixon, “Genomics-based selection and functional characterization of triterpene glycosyltransferases from the model legume Medicago truncatula,” Plant Journal, vol. 41, no. 6, pp. 875–887, 2005. View at Publisher · View at Google Scholar · View at Scopus
  48. D. Meesapyodsuk, J. Balsevich, D. W. Reed, and P. S. Covello, “Saponin biosynthesis in Saponaria vaccaria. cDNAs encoding β-amyrin synthase and a triterpene carboxylic acid glucosyltransferase,” Plant Physiology, vol. 143, no. 2, pp. 959–969, 2007. View at Publisher · View at Google Scholar · View at Scopus
  49. S. Lodeiro, Q. Xiong, W. K. Wilson, M. D. Kolesnikova, C. S. Onak, and S. P. T. Matsuda, “An oxidosqualene cyclase makes numerous products by diverse mechanisms: a challenge to prevailing concepts of triterpene biosynthesis,” Journal of the American Chemical Society, vol. 129, no. 36, pp. 11213–11222, 2007. View at Publisher · View at Google Scholar · View at Scopus
  50. T. Husselstein-Muller, H. Schaller, and P. Benveniste, “Molecular cloning and expression in yeast of 2,3-oxidosqualene-triterpenoid cyclases from Arabidopsis thaliana,” Plant Molecular Biology, vol. 45, no. 1, pp. 75–92, 2001. View at Publisher · View at Google Scholar · View at Scopus
  51. M. Suzuki, T. Xiang, K. Ohyama et al., “Lanosterol synthase in dicotyledonous plants,” Plant and Cell Physiology, vol. 47, no. 5, pp. 565–571, 2006. View at Publisher · View at Google Scholar · View at Scopus
  52. D. R. Phillips, J. M. Rasbery, B. Bartel, and S. P. Matsuda, “Biosynthetic diversity in plant triterpene cyclization,” Current Opinion in Plant Biology, vol. 9, no. 3, pp. 305–314, 2006. View at Publisher · View at Google Scholar · View at Scopus
  53. M. Shibuya, H. Zhang, A. Endo, K. Shishikura, T. Kushiro, and Y. Ebizuka, “Two branches of the lupeol synthase gene in the molecular evolution of plant oxidosqualene cyclases,” European Journal of Biochemistry, vol. 266, no. 1, pp. 302–307, 1999. View at Publisher · View at Google Scholar · View at Scopus
  54. M. D. Kolesnikova, Q. Xiong, S. Lodeiro, L. Hua, and S. P. Matsuda, “Lanosterol biosynthesis in plants,” Archives of Biochemistry and Biophysics, vol. 447, no. 1, pp. 87–95, 2006. View at Publisher · View at Google Scholar · View at Scopus
  55. R. Martín, J. Carvalho, E. Ibeas, M. Hernández, V. Ruiz-Gutierrez, and M. L. Nieto, “Acidic triterpenes compromise growth and survival of astrocytoma cell lines by regulating reactive oxygen species accumulation,” Cancer Research, vol. 67, no. 8, pp. 3741–3751, 2007. View at Publisher · View at Google Scholar · View at Scopus