Occurrence of iridoid glycosides in in vitro cultures and intact plants of Scrophularia nodosa L.
- First Online:
- Cite this article as:
- Sesterhenn, K., Distl, M. & Wink, M. Plant Cell Rep (2007) 26: 365. doi:10.1007/s00299-006-0233-3
- 251 Views
Shoot, root, and callus cultures of Scrophularia nodosa L. (Scrophulariaceae) were established and cultivated in vitro. Iridoid glycosides, such as harpagoside, aucubin, and catalpol were identified by LC-ESI-MS and their contents determined by HPLC. For comparison intact plants of S. nodosa were analysed. In shoot cultures slightly lower amounts of detectable iridoid glycosides (4.36% dry weight) were determined than in the field grown plants (4.88%). Concentration of harpagoside was highest in leaves of field plants (1.05%) and in flowers of in vitro plantlets (1.10%). For aucubin the highest amount was found in the leaves of in vitro plantlets (1.67%) whereas the levels of aucubin in the leaves of field plants were remarkably lower. Catalpol was produced as a trace compound in intact plants and shoot cultures. Callus and root cultures were apparently not able to synthesise iridoid glycosides.
KeywordsScrophularia nodosaIridoid glycosidesHarpagosideAucubinPlant in vitro culture
Woody plant medium (Lloyd and McCown 1980)
High performance liquid chromatography
Liquid chromatography – electrospray ionisation – mass spectrometry
The common figwort Scrophularia nodosa L. (Scrophulariaceae) is one of the over 300 known species of the genus Scrophularia, which has been used since ancient times in traditional medicine. S. nodosa is a 0.5–1.2 m high perennial herb showing a characteristic brown rhizome with nodal swelling and small inconspicuous flowers. It grows in deciduous and coniferous forests of Central Europe, Central Asia, and North America. The plant has been used in traditional medicine to treat scrofula, eczema, wounds, pain of the hips, eye complaints, goitre, ulcers, cancer, fistulae, and furthermore as diuretic and anthelmintic (Pauli et al. 1995; Weiss 1985; Pahlow 2001; Stevenson et al. 2002).
Characteristic for the genus Scrophularia is the accumulation of iridoid glycosides like aucubin, catalpol, and harpagoside (Pauli et al. 1995). Grabias et al. (1995) have reported the presence of the iridoids harpagide, harpagoside, harpagide acetate in the seeds of S. nodosa. From a pharmacological point of view the iridoids are among the most interesting compounds in these plants. They show a variety of properties such as choleretic, vasoconstrictory, hepatoprotective, antiinflammatory, antiviral, and antimicrobial effects. The antibacterial and further biological activities of aucubin are probably attributed to the aglycone aucubigenin, which is formed after cleavage by β-glucosidase (Rischer et al. 1998; Wichtl 1989). Antiinflammatory and also analgesic properties have been shown for Harpagophytum procumbens DC (Pedaliaceae) containing harpagoside as main iridoid glycoside. In vitro studies indicate that harpagoside, harpagide, procumbine, and harpagogenin may be responsible for the antiinflammatory and analgesic activity of the drug. Since herbal extracts have less adverse effects as compared to synthetic nonsteroidal anti-inflammatory drugs, there is a high demand for the corresponding plants (Levieille and Wilson 2002; Baghdikian et al. 1997; Chrubasik and Wink 1998). The increasing application of extracts from the secondary root tubers of H. procumbens has led to an excessive exploitation of the species in the Kalahari (Levieille and Wilson 2001). Therefore, an alternative source of harpagoside may be of interest. The combined presence of the three iridoid glycosides harpagoside, aucubin, and catalpol make S. nodosa an especially interesting candidate as a substitute.
The objective of the present investigation was to determine the amount of the iridoid glycosides harpagoside, aucubin, and catalpol produced in different plant tissues of S. nodosa. For this purpose, shoot, root, and callus cultures were established and cultivated in vitro. The iridoids were identified by liquid chromatography – electrospray ionisation – mass spectrometry (LC-ESI-MS) and quantified by high performance liquid chromatography (HPLC). In addition, plants collected in the wild were analysed for comparison.
Materials and methods
Intact plants and seed material
Mature seed capsules of S. nodosa were collected during August 2003 and intact plants were collected from sandy soil with low sun exposition in the forest of Dossenheim, Germany, in July 2004. A voucher specimen is deposited in the herbarium of the IPMB, Department Biology, University of Heidelberg, Germany.
Seeds were surface sterilised in 70% ethanol for 1 min followed by 1% NaOCl for 3 min. Finally, they were rinsed five times with autoclaved water.
In vitro cultures
Seeds were either placed on agar-solidified Woody Plant Medium (WP) (Lloyd and McCown 1980) without sucrose or on the same medium containing 250 mg/l (0.72 mM) gibberellic acid (GA3) (Carl Roth, Karlsruhe, Germany) for the enhancement of germination. The pH of the medium was adjusted to 5.7 before autoclaving at 1200 Hpa (121°C) for 20 min. GA3 stock solution was sterilised by filtration and added after autoclaving before the medium became solid. All cultures were maintained at 25°C±1°C under continuous cool-white fluorescent light (Phillips, 36 W) delivering 38 μmol m−2 s−1. After germination all seedlings were kept on hormone-free medium. Medium was changed every 4 weeks. Average germination rates and duration of germination were determined by repeating the experiment two times.
Root cultures were initiated from roots of the in vitro cultures. Intact root parts of at least 5 cm showing lateral roots were cut off and incubated in 50 ml liquid WP medium with 3% sucrose. Cultures were kept in the dark on a rotary shaker (110 rpm) at 25°C±1°C. They were transferred into fresh medium every 3 weeks.
A growth curve was calculated by determination of the net produced fresh weight (fresh weight at harvest − inoculum fresh weight at subculture) every second or third day over a cultivation period of 28 days. The experiment was repeated twice. The production of iridoid glycosides was monitored by HPLC analysis over the whole cultivation period.
Fragments of leaves (1–2 cm) and roots (2 cm) were placed on WP medium with 3% sucrose containing 0.1 mg/l (0.46 μM) kinetin and 0.5 mg/l (2.26 μM) 2,4-dichloroacetic acid (2,4-D). The callus appeared after 2 weeks and was then transferred every third week onto fresh agar medium with the same hormone concentrations as used for callus induction. Growth indices were calculated (fresh weight at harvest/inoculum fresh weight at subculture) over three passages. The concentration of iridoid glycosides was determined after the third passage by HPLC analysis.
Shoot tips from the in vitro cultured plants of S. nodosa with at least two young leaves were cut off. They were rooted on WP medium with 3% sucrose without growth regulators.
Preparation of extracts
Harvested intact plants and in vitro cultured plantlets were immediately divided into root, stem, leaves, and flowers. Samples were also taken from callus cultures and root cultures. The samples were stored at −20°C in aluminium foil. Before extraction, all samples were lyophilised (Christ, Osterode, Germany).
The lyophilised samples were powdered and weighed (10–100 mg). They were extracted in 20 ml methanol for 3 min under sonication and then heated at 50°C for 30 min. After shaking on a rotary shaker over night, the samples were centrifuged at 4000 rpm for 10 min. The supernatant was filtered and the residue was washed two times with 10 ml MeOH and also filtered. The combined supernatants were evaporated under reduced pressure and redissolved in 2 ml methanol. Recovery rate was determined to ensure that no harpagoside, aucubin, or catalpol was lost during the extraction process. The recoveries were 101.70%±3.13% for harpagoside, 99.46%±4.7% for aucubin, and 98.68%±6.44% for catalpol.
Harpagoside and aucubin were purchased from Carl Roth (Karlsruhe, Germany), catalpol from Wako (Neuss, Germany). Standards were dissolved in methanol.
Gradient 1 (harpagoside): 0–2 min, 10% B; 2–12 min, 10–40% B; 12–22 min, 40–100% B; 22–26 min, 100–10% B.
Gradient 2 (aucubin, catalpol): 0–1 min, 0% B; 1–16 min, 0–10% B; 16–17 min, 10–100% B; 17–18 min, 100–0% B.
The peak area was integrated by Gold Nouveau Chromatography Data System Software Version 1.72, 1996, Beckmann Instruments, with external standard. A standard calibration curve was plotted by using various concentration ranges of harpagoside (0.2–500 μg/ml), aucubin (7.8–1000 μg/ml), and catalpol (15.6–500 μg/ml), respectively.
Harpagoside, aucubin, and catalpol were additionally identified by LC-ESI-MS. The LC-ESI-MS system consisted of an HPLC from Latek Laborgeräte GmbH (Heidelberg, Germany) coupled with a Micromass VG Quattro II mass spectrometer (Waters, Manchester, United Kingdom). ESI-MS was operated under MassLynx software (version 4.0, Waters). Nitrogen was used as nebulising and drying gas, and was generated by a Parker nitrogen generator (Parker, Etten-Leur, The Netherlands).
An electrospray interface was utilised for ionisation. LC-separation was carried out with a LiChrospereTM 100 RP-18 column (250×4 mm, 5 μm) from Merck, Darmstadt, Germany. Separation of the iridoid glycosides at ambient temperature was achieved using a binary gradient system consisting of water (A) and acetonitrile (B), both containing 0.1% formic acid. The pump program reached from 0 to 10% B within 15 min, followed to 100% B in another 15 min. Post-flow splitting was set to 1:5. Sample volumes of 20 μl were injected by a Rheodyne injection valve.
Mass spectrometric detection of sodium adducts of iridoid glycosides (Harpagoside m/z 517 [M+Na]+, aucubin m/z 369 [M+Na]+, catalpol m/z 385 [M+Na]+) was performed in the positive ion mode over the range of m/z 50–1000 and the instrument was set to the following tune parameters: nebulising gas pressure of 13 l/h and drying gas pressure of 350 l/h, voltage of capillary electrophoresis was set to 3.5 kV, HV lens to 0.5 kV, cone voltage to 50 V, and the temperature of the heated transfer capillary was maintained at 120°C. Chromatograms were processed using MassLynx 4.0 software (Waters).
Results have been expressed as mean values in % dry weight±standard deviation (in vitro cultures, n=20; wild plant material, n=7).
In vitro cultures and intact plants
Seeds of S. nodosa germinated after 8 days on WP medium with an average germination rate of 4%. On WP medium supplemented with GA3 the germination rate could be enhanced to 61% after 7 days. The seedlings grew well on WP medium without sucrose. The in vitro growth conditions had a strong influence on the development of the in vitro cultivated plantlets of S. nodosa. Leaves and stems of these plantlets remained small compared to intact plants; roots were not capable of developing a rhizome. Flowering was induced after several weeks of cultivation. In vitro flowers showed similar size like those of intact plants.
In wild-grown plants, the iridoid glycosides could be detected in all plant parts. Catalpol was seen sporadically but only in traces. Different from the in vitro cultures, harpagoside was the most abundant of the three iridoid glycosides with 2.95%, followed by aucubin with 1.93%. In wild plants, the highest amount of aucubin was detected in stems and harpagoside concentration was highest in leaves and flowers. The detectable iridoid glycoside amounts were 4.88% for the wild plants and 4.36% for the in vitro cultured plantlets.
The attempt to establish stable and viable root cultures was only successful in some cases. In most instances the roots stopped growing or grew at a very slow rate. One well-growing root line initially showed increasing growth, which yielded a maximum net mass production of 6.47 g on day 23. In the roots analysed during the growth cycle none of the iridoid glycosides could be detected at any time. This corresponds to the low iridoid levels in the roots of the in vitro plantlets indicating that there may not be a site of iridoid biosynthesis.
Using the WP medium with 3% sucrose and supplementation of kinetin and 2,4-D we obtained three callus lines from leaf explants and six lines from roots. This medium proved to be appropriate for the induction of primary callus from leaves and roots of S. nodosa as well as for the cultivation of callus cultures.
All calli showed a green colour and a firm and friable consistency. The growth rates showed a high variation. For the leaf callus growth rate increased from 2.31±1.01 to 6.10±4.35 over three passages and for root callus from 1.59±0.62 to 3.11±1.29. After the third passage, the calli were analysed by HPLC; no iridoid glycosides could be detected.
In our experiments, the formation of roots was successful without growth regulators. In 70% of cases the cutoff shoots from in vitro plants had developed several roots from the basal end after 1–2 weeks. No callus formation or browning was observed during this process and shoot elongation was also induced. The regenerated plantlets developed as well as plants derived from seedlings. The easily regenerated plantlets of S. nodosa may be useful as a simple experimental system for studies on the iridoid glycoside biosynthesis.
Plant organs of in vitro grown plants showed a different development as those of intact field grown plants. However, the percentage of detectable iridoids in the in vitro grown plants (4.36%) was only slightly lower than that in the intact plants (4.88%). Generally, organs important for survival and reproduction have the highest and most potent secondary metabolites (Wink 2003). This may explain why a high iridoid content was found in the flowers of both in vitro grown and intact plants.
Our analysis revealed variations of the iridoid glycoside concentration between different plants of one population and between different organs of individual plants. This result can be expected taking into account that the production of secondary metabolites is a complex process between biosynthesis, transport, storage, and degradation in the intact plant (Wink 1987, 1989, 1990). The accumulation of a secondary metabolite depends on the equilibrium between these processes and can therefore change in a tissue-, organ-, and development-specific way (Wink 2003).
Concentration of the iridoid glycosides harpagoside, aucubin, and catalpol in different tissues of intact plants compared to the concentration of in vitro cultivated shoot cultures
Levieille and Wilson (2002) reported the presence of harpagoside and harpagide (0.1% dry weight) in the leaves of micropropagated plants of H. procumbens and questioned the site of iridoid biosynthesis in the plant. In the tubers of these plants they found about 1% iridoids, which is comparable to the values found in the tubers of the wild plants. They suggested that the leaves may provide an alternative source for the iridoids and assumed that they would show the same pharmacological activity as tuber extracts. In our investigations leaves of S. nodosa contained as much as or even more harpagoside (1.05%) than do the secondary root tubers of micropropagated plants of H. procumbens. Even the leaves of our in vitro cultivated plantlets of S. nodosa showed values of at least 0.41% harpagoside and 1.67% aucubin and represent therefore a rich and maybe alternative source of these two iridoid glycosides.
Our results indicate that some plants of the genus Scrophularia like S. nodosa may serve as an alternative source of harpagoside. Moreover, S. nodosa may be interesting with regard to the production of aucubin. Further compounds in the genera Scrophularia and Harpagophytum may enlarge the pharmacological profile of the extracts. For the comparison of the pharmacological properties of extracts of H. procumbens with those of S. nodosa further investigations are required.
We thank J. Freund, who participated as a research student in this project.