Triterpenoids and ellagic acid derivatives from in vitro cultures of Camptotheca acuminata Decaisne

Plant Physiology and Biochemistry 44 (2006) 220–225 www.elsevier.com/locate/plaphy

Research article

Triterpenoids and ellagic acid derivatives from in vitro cultures of Camptotheca acuminata Decaisne G. Pasqua a,*, A. Silvestrini b, B. Monacelli a, N. Mulinacci c, P. Menendez d, B. Botta e a

Dipartimento di Biologia Vegetale, Università degli Studi di Roma “La Sapienza”, Piazzale Aldo Moro 5, 00185 Rome, Italy b Istituto di Biochimica e Biochimica Clinica, Università Cattolica S. Cuore, Largo F. Vito 1, 00168 Rome, Italy c Dipartimento di Scienze Farmaceutiche, Via Ugo Shiff 6, 50019 Sesto Fiorentino, Florence, Italy d Catedra de Farmacognosia, Universidad de la Republica, 18 de Julio 1968, 11200 Montevideo, Uruguay e Dipartimento di Studi di Chimica e Tecnologia delle Sostanze Biologicamente Attive, Università degli Studi di Roma “La Sapienza”, Piazzale Aldo Moro 5, 00185 Rome, Italy Received 1 July 2005 Available online 25 April 2006

Abstract The metabolic profile of secondary products in calli and cell suspension cultures of Camptotheca acuminata Decaisne was investigated and compared to that of the leaves and roots taken from the plant. Neither in vitro system produced the anticancer quinoline alkaloid camptothecin (CPT); they accumulated discrete quantities of polyhydroxylated triterpenoids, different from those found in the plant organs, and ellagic acid derivatives. Nine ellagic acid derivatives (1a–1d and 2a–2e) and eight triterpenoid acids (3a–3e and 4a–4c) were isolated and characterised. All the identified triterpenes were related to ursane- or oleanane-type skeletons and their concentrations rose to 4.5% in suspended cells. © 2006 Elsevier SAS. All rights reserved. Keywords: Camptotheca acuminata; Cell cultures; Ellagic acid derivatives; Triterpenes; Camptothecin

1. Introduction Camptotheca acuminata Decaisne (Nyssaceae) is a well known natural source of the monoterpene-indole alkaloid camptothecin (CPT), one of the most promising anti-tumoural compounds, which was first isolated and structurally described in 1966 [1]. CPT blocks cell division by inhibiting topoisomerase I, an enzyme involved in DNA coiling. However, clinical trials conducted in the 1970s showed that CPT caused severe side effects, leading to the suspension of these trials [2]. In the 1990s, some semi-synthetic water-soluble CPT derivatives were successfully applied clinically. Up today these derivatives are synthesised from natural CPT extracted from C. acuminata and from other species belonging to unrelated orders and families of angiosperms [3]. CPT derivatives have also been studied as potent inhibitors of adenoviruses, papovaviruses, and

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0981-9428/$ - see front matter © 2006 Elsevier SAS. All rights reserved. doi:10.1016/j.plaphy.2006.04.001

herpesviruses [4], and they have shown promising activity against parasitic trypanosomes and Leishmania [5]. Although much is known about the pharmacological properties of CPT, little is known about its biosynthesis [3]. The first steps in the biosynthetic pathway, which are considered to be common to all monoterpene-indole alkaloids, lead to strictosidine, which derives from a condensation reaction between the indole tryptamine and secologanin [6]. Feeding experiments on C. acuminata plants using radio-labelled precursors have confirmed that tryptamine is incorporated in CPT [7]. Moreover, the genes encoding tryptophan decarboxylase (TDC) (the enzyme that converts tryptophan into tryptamine) have been cloned in C. acuminata, with the highest expression found in the organs with the highest CPT levels [8]. With regard to secologanin, which presumably derives from the monoterpene geraniol by secologanin synthase, already described in Catharanthus roseus [9], has never been isolated from C. acuminata. Furthermore, strictosidine and strictosidine synthase (SSS) have never been isolated from C. acuminata, although one study has reported an extremely low SSS activity in C. acuminata leaf [10], which is the organ with the greatest CPT accu-

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mulation [11]. Different strategies for sustainable CPT production from C. acuminata plants have been successfully used (i.e. biotic and environmental stress) [12,13]. In vitro cultures have also been tested to produce CPT, but the levels of production have not been comparable to those obtained in the plant [14– 16], and cell suspensions of C. acuminata have been reported to lose the biosynthetic ability to produce CPT after a few subcultures [17]. In a previous study, we reported that no CPT was produced by a CG1 cell line of C. acuminata also after precursor feeding [10]. Regarding other CPT-producing plants, calli obtained from Ophiorrhiza pumila did not produce CPT [18], and cell suspensions of Nathapodytes foetida only produced small amounts (100–1000-fold lower than the plant) [19]. In the present study, the metabolic profile of secondary products in calli and cell suspensions of C. acuminata were analysed and compared to that of the leaves and roots taken from the plant. The results from this research might contribute to explain the lack or scant CPT production in cell cultures. 2. Results and discussion

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Table 1 Yields (%) of the different extracts and metabolites content in in vitro systems and in plant organs (leaves and roots). Triterpenes and ellagic acid derivatives were gravimetrically determined while CPT was evaluated by HPLC according to Pasqua et al. 2004 [13] Sample Calli Suspended cells Culture medium Leaves Roots

Ethyl acetate extract 3.5 10.5 9.5 8 7.5

Triterpenes 1.5 4.6 3.8 (+) (+)

Ellagic acid derivatives 0.15 0.48 0.17 (+) (+)

CPT (–) (–) (–) 0.2 0.15

(–) not found; (+) traces amount. All data were expressed as percent with respect to the dried weight of the sample (DW).

in Table 1. Both of the in vitro systems failed to produce CPT. Histochemical analyses carried out on both calli and cell suspensions showed the presence of phenolic compounds in the vacuoles (Fig. 2A). Moreover, microscopic observations of the in vitro cells under UV light showed blue autofluorescence (Fig. 2B), which was due to the presence of ellagic acid derivatives, identified through chemical analysis. Differently, blue autofluorescence

2.1. Calli and cell suspension cultures After 14 days of culture, calli, which were cream in colour, were formed on the cut surface of the leaf explants. After several subcultures, some calli were chemically analysed and some were used to obtain cell suspension cultures. On day 30 (i.e. at the end of the exponential phase), the cell biomass was significantly higher when 3% inoculum was used, compared to inocula of 6, 10, and 20% (there were no significant differences among the latter three) (Fig. 1). 2.2. Secondary metabolites in the plant and in the in vitro systems The results of the semi-quantitative evaluation of the main metabolites found in the organic extracts from in vitro cultures and from the leaves and roots taken from the plant are shown

Fig. 1. Effect of inoculum concentration on the biomass increment in cell suspension cultures. Each point represents the mean of three increment values ± S.E. (Vf/Vi).

Fig. 2. Microscopic observation. A) Phenolic accumulation in the suspension cell (arrows) coloured blue–green by cresyl blue reagent (bar 10 μm). B) Autofluorescence blue in the suspension cells, showing ellagic-acid derivatives under UV light (bar 10 μm). C) Section of the leaf showing autofluorescent CPT crystals, observed under UV light, in the abaxial parenchyma of the midrib; (bar 1 mm) D). CPT crystals, observed under UV light, in a unicellular glandular trichome on the abaxial side of the leaf (bar 10 μm).

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observed, under UV light, in both leaf (Fig. 2C, D) and root tissues, was correlated to CPT accumulation in crystalline form, quantified by HPLC analysis (Table 1). In the ethyl acetate extracts of both calli and cell suspension cultures, the most abundant ellagic acid derivative was 3,3′,4tri-O-methylellagic acid (Fig. 3, 1b). Other components were 3,4-O-methylene-3′4′-di-O-methyl ellagic acid (Fig. 3, 2c) and 3,4-O-methylene-3′,4′,5′-tri-O-methylellagic acid (Fig. 3, 2d). 1 H NMR and mass spectral data of impure fractions of 1b, as compared with those of the literature [20–22], supported the finding of a minor component in the extracts: 3,4-O-methylene-3′-O-methyl ellagic acid (Fig. 3, 2b). Some ellagic acid derivatives identified in in vitro cultures were previously described in C. acuminata fruits and root bark [23,24]. Triterpenoids were present in the calli, the cell suspensions, and in the culture medium, in estimated quantities as shown in Table 1. Traces amount of these metabolites were detected in the extracts of both leaves and roots. All triterpenes identified were related to ursane- or oleanane-type skeletons: the two structures are regioisomeric and differ for the position on the

E ring of the β methyl group, which is located at C-19 in the first compound and at C-20 in the second compound. Both ursolic (3β-hydroxy-12-ursen-28-oic) acid (Fig. 3, 3a) and oleanolic (3β-hydroxy-12-oleanen-28-oic) acid (Fig. 3, 4a) were present in small quantities in the extracts. The parent compounds α-amyrin (3α-hydroxy-12-ursene) and β-amyrin (3β-hydroxy-12-oleanene) were also found in traces. The two pairs were isolated as a mixture but easily distinguishable by the chemical shifts (δ 122 and 143 for 3a vs. δ 125 and 138 for 4a) of the olefinic carbons 12 and 13. The 2α-hydroxy derivatives of both ursolic and oleanolic acids were isolated as methyl esters. The two stereoisomers are known as “corosolic acid” (Fig. 3, 3b) and “maslinic acid”. 2α,3β,23-Trihydroxy-12-ursen-28-oic (asiatic) acid (Fig. 3, 3c) and 2α,3β,23-tri-hydroxy-12-oleanen-28-oic (arjunolic) acid (Fig. 3, 4c) were the main components of the extracts from both the calli and cell suspensions. Finally, two minor triterpene metabolites were also isolated as methyl esters and were assigned the structures 2α,3α-dihydroxy-12-ursen-28-oic acid (Fig. 3, 3d) and 2α,3α-23-trihydroxy-12-ursen-28-oic acid

Fig. 3. Chemical structures of the main metabolites isolated from C. acuminata cell cultures: ellagic acid derivatives (1a–1d and 2a–2e) and triterpenoid acids (3a–3e and 4a–4c).

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(Fig. 3, 3e) (i.e. esculentic acid). The absence of the corresponding oleanoic acids may depend on steric hindrance of the D/E ring conformation towards the action of the 3α-hydroxylating enzyme. It is worth noting that the basic parent skeletons of ursolic and oleanolic acids show selective hydroxylations, probably performed by a cytochrome P-450. Most of the triterpenes isolated from C. acuminata cell cultures have also been found in the calli of Actinidia arguta [25], Hyssopus officinalis [26], Ternstroemia gymnanthera [27] and Eriobotrya japonica [28] and in the plant organs of different species, such as Crataegus pinnatifida [29], Cornus capitata [30], Lagerstroemia speciosa [31], Geum japonicum [32], and Cochlospermum tinctorium [33], Tabernaemontana divaricata [34]. Important biological activities of these compounds have been reported in the literature: inhibition of protein kinase C [29], inhibition of the activation of the Epstein–Barr virus early antigen [33], induction of apoptosis [35], inhibition of HIV-1 [32], neuroprotective action [36], anti-diabetic activity [31], and inhibition of insect growth [30]. 3. Conclusions The precursor of terpenoid biosynthesis is isopentenyl diphosphate (IPP); for a number of years, it was believed that IPP could only be formed through the mevalonate (MVA) pathway ([37] and references therein). In recent studies the existence of a separate pathway for the synthesis of IPP and DMAPP was detected [38–39]. Hampel et al. [40] showed that in Daucus carota roots and leaves, monoterpenes are synthesised exclusively via the 1-deoxy-D-xylulose/2-C-methyl-D-erythritol-4-phosphate (DOXP/MEP) pathway, whereas sesquiterpenes are generated by the mevalonic acid pathway (MVA) as well as the DOXP/MEP route. Both MVA and MEP pathways can operate simultaneously in higher plants. The enzymes of the MEP pathway are located in plastids, where they produce precursors for monoterpenes and some sesquiterpenes, diterpenes and carotenoids [39]. The enzymes of the MVA pathway are located in the cytoplasm, where they supply precursors of triterpenes, sesquiterpenes and sterols [37]. However, it has been recently shown that the compartmental separation of the two pathways in plant cells is not absolute, and cross-talk between MVA and MEP pathways can occur depending on the species and the physiological conditions [41– 43]. Our results support the hypothesis that in C. acuminata cell cultures MEP pathway might be inhibited or a possible shift of the isoprenoids from the MEP to the MVA might be occurred. Such hypothesis it has been previously proposed for other species producing monoterpene-indole alkaloids such as Tabernaemontana cell cultures [34]. Under elicitor treatment, these cultures showed a reduction of alkaloid biosynthesis and an increased production of pentacyclic triterpenes [34]. Further studies, are necessary to clarify the complex regulatory mechanisms involved in the C. acuminata terpenes meta-

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bolism, both for manipulating the biosynthetic capability of cell cultures and improving the in vitro production of CPT. 4. Methods 4.1. Plant material Two-year-old C. acuminata Decaisne (Nyssaceae) plants were used for the experiments. The plants were cultivated in the field, in the Botanical Garden of “La Sapienza” University (Rome, Italy). 4.2. Calli and cell suspension cultures Explants from young leaves were cultured in B5 Gamborg’s medium [44] supplemented with 1 mg l−1 2,4-dichlorophenoxyacetic acid (2,4-D), 0.5 mg l−1 kinetin (kin), 40 g l−1 sucrose, and 8 g l−1 agar, to induce callus formation. The pH was adjusted to 5.6 before autoclaving. The explants were cultured in the dark at 26 °C. Subcultures were carried out every 20 days and maintained in the same cultural conditions. Suspension cultures were initiated by transferring 2 g fresh weight (FW) of callus to 250 ml Erlemayer flasks containing 65 ml of the same medium without agar. The suspension cultures were incubated at 26 °C under a photoperiod of 16/8 h day/night on a rotary shaker (New Brunswick G10) at 100 rpm. The influence of different amounts of inoculum (3, 6, 10, and 20%) on the biomass growth was considered. Every 5 days, until day 35, the increment in cell growth (Vf/Vi) was measured by determining Packed Cell Volume (PCV). The PCV was measured by transferring 10 ml of cell suspension to a calibrated conical tube followed by centrifugation for 5 min at 600 × g. Each point on the growth curve represents the mean of three measurements (PCVf/PCVi ± standard error). The significance of the differences between the means were evaluated using the variance analysis at P < 0.05. 4.3. Microscopic analysis Calli, cell suspensions and leaf and root sections (~30 μm thickness obtained with a vibratome TPI series 1000) were examined with a Zeiss microscope (Axioscop 2 Plus) using a Zeiss UV-filter combination (BP 365 nm, LP 397 nm) for CPT detection, according to Pasqua et al. [13]. Some samples were observed in brightfield after having been stained using reagents specific for phenolic compounds: 1% toluidine blue or cresyl blue. 4.4. Extraction procedure We analysed calli (11.5 g DW), 20-day-old filtered cells (8 g FW), the culture media (500 ml – 4.2 g DW), and ground samples of plant leaf and root (15 g each – about 2 g DW). Suspended cells were separated from the medium by filtering under vacuum.

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All the samples were homogenised three times with ultra Turrax at 1000 rpm and extracted with MeOH (1 ml g−1 FW for three times). The filtered medium was concentrated to remove the MeOH and the aqueous residue was submitted to a three steps liquid/liquid extraction procedure using ethyl acetate. The ethyl acetate extracts were pooled, filtered on Whatman 1006 paper and dried under vacuum. The dried organic extract was dissolved in a 2:1 chloroform/methanol mixture and analysed by silica gel thin layer chromatography (TLC). At the same time, the dried water residue was dissolved in D2O and directly analysed by NMR to control its composition. The APT 13C NMR spectrum of the mixture revealed, by comparison with analogous spectra of common sugars, that sucrose was the main component and that glucose and fructose were minor components. Given that no other carbon signal with significant intensity was present in the spectra, the water extract was discarded. Firstly a TLC examination of the ethyl acetate extracts was carried out using CHCl3/EtOAc/MeOH mixtures of increasing polarity. The presence of the metabolites was revealed under UV light at 254 and 365 nm and/or by spraying with 10% H2SO4 in MeOH and heating at 100–110 °C. The organic extracts were finally purified on silica gel column or preparative TLC, eluting with solvent mixtures of the proper polarity, as suggested by TLC analysis. Complex fractions of triterpenes were methylated by CH2N2 and chromatographically resolved by isolation of methyl derivatives. 4.5. Identification of metabolites The structures of the secondary metabolites were determined by 1H, 13C NMR and mass spectra of the isolated products and/or of their derivatives, in comparison with the data in the literature. 1H and 13C NMR spectra (300 and 75 MHz, respectively) were run with a Varian Gemini, tetramethylsilane (TMS) as internal standard, and the appropriate solvent among CDCl3, CD3OD, and C5D5N. Electron impact (EI) mass spectra were performed with a VG 7070EQ spectrometer. 4.5.1. Ellagic acid derivatives 3,3′,4-Tri-O-methylellagic acid (1b) was identified by its UV, 1H, 13C NMR and mass spectral data, in comparison with those reported in the literature [20–22]. 3,4-O-Methylene-3′-O-methyl ellagic acid (2b): 1H NMR (CDCl3) δ: 7.77 (1H, s, H-5′), 7.62 (1H, s, H-5), 6.29 (2H, s, OCH2O), 4.18 (3′-OMe);13C NMR: δ 60.1 (3′-OMe); EIMS m/z (relative intensity): 328 [M]+ (100), 313 (49), 285 (20), 255 (32). 3,4-O-Methylene-3′,4′-di-O-methyl ellagic acid (2c): 1H NMR (CDCl3) δ: 7.72 (1H, s, H-5′), 7.64 (1H, s, H-5), 6.29 (2H, s, OCH2O), 4.23 (4′-OMe), 4.04 (3′-OMe); EIMS: 342 [M]+ (lit 342 [6]). 3,4-O-Methylene-3′,4′,5′-tri-O-methyl ellagic acid (2d): 1H NMR (CDCl3): δ 7.61 (1H, s, H-5), 6.29 (2H, s, OCH2O), 4.28 (4′-OMe), 4.04, 4.02 (3′-, 5′-OMe); 13C NMR: δ 62.4, 62.1, 62.0 (3′,4′,5′-OMe); EIMS m/z (relative intensity): 372

[M]+ (100), 357 (54), 343 (21), 329 (17), 314 (21), 271 (21), 215 (17). 4.5.2. Ursolic/oleanolic acids triperpene derivatives Ursolic acid (3a), oleanolic acid (4a), α- and β-amirin were identified by comparison (TLC, 1H and 13C NMR spectra) with authentic samples. The structure of the triterpenes 2α,3β,-dihydroxy-12-ursen-28-oic acid (3b), 2α,3β,dihydroxy-12-oleanen28-oic acid (4b), 2α,3β,23-trihydroxy-12-ursen-28-oic acid (3c) 2α,3β,23-trihydroxy-12-oleanen-28-oic acid (4c), 2α,3α,-dihydroxy-12-ursen-28-oic acid (3d) and 2α,3β- dihydroxy-12-oleanen-28-oic acid (3e) was assigned essentially based on a comparison of 1H and 13C NMR spectral data with those from the literature [45,46]. References [1] M.E. Wall, M.C. Wani, C.E. Cook, K.H. Palmer, A.T. McPhail, G.A. Sim, Plant antitumor agents. I. The isolation and structure of camptothecin, a novel alkaloid leukaemia and tumor inhibitor from Camptotheca acuminata, J. Am. Chem. Soc. 88 (1966) 3888–3890. [2] J.F. Pizzolato, L.B. Saltz, The camptothecins, Lancet 361 (2003) 2235– 2242. [3] A. Lorence, C.L. Nessler, Camptothecin, over four decades of surprising findings, Phytochemistry 65 (2004) 2735–2749. [4] P. Pantazis, Z. Han, D. Chatterjee, J. Wyche, Water-insoluble camptothecin analogues as potential antiviral drugs, J. Biomed. Sci. 6 (1999) 1–7. [5] A.L. Bodley, T.A. Shapiro, Molecular and cytotoxic effects of camptothecin, a topoisomerase I inhibitor, on trypanosomes and Leishmania, Proc. Natl. Acad. Sci. USA 92 (1995) 3726–3730. [6] T.M. Kutchan, Alkaloid biosynthesis: the basis for metabolic engineering of medicinal plants, Plant Cell 7 (1995) 1059–1070. [7] G.M. Sheriha, H. Rapoport, Biosynthesis of Camptotheca acuminata alkaloids, Phytochemistry 15 (1976) 505–508. [8] M. Lopez-Meyer, C.L. Nessler, Tryptophan decarboxylase is encoded by two autonomously regulated genes in Camptotheca acuminata which are differentially expressed during development and stress, Plant J. 11 (1997) 1167–1175. [9] S. Irmel, G. Schröder, B. St-Pierre, N.P. Crouch, M. Hotze, J. Schmidt, D. Strack, U. Matern, J. Schröder, Indole alkaloid biosynthesis in Catharanthus roseus: new enzyme activities and identification of cytochrome P450 CYP72A1 as secologanin synthase, Plant J. 24 (2000) 797–804. [10] A. Silvestrini, G. Pasqua, B. Botta, B. Monacelli, R. van der Heijden, R. Verpoorte, Effects of alkaloid precursor feeding on a Camptotheca acuminata cell line, Plant Physiol. Biochem. 40 (2002) 749–753. [11] M. Lopez-Meyer, C.L. Nessler, T.D. McKnight, Sites of accumulation of the antitumor alkaloid camptothecin in Camptotheca acuminata, Planta Med. 60 (1994) 558–560. [12] Z. Liu, J.C. Adams, H.P. Viator, R.J. Constantin, S.B. Carpenter, Influence of soil fertilization, plant spacing, and coppicing on growth, stomatal conductance, abscisic acid, and camptothecin levels in Camptotheca acuminata seedlings, Physiol. Plant. 105 (1999) 402–408. [13] G. Pasqua, B. Monacelli, A. Valletta, Cellular localization of the anticancer drug camptothecin in Camptotheca acuminata, Eur. J. Histochem. 48 (2004) 321–328. [14] K. Sakato, H. Tanaka, N. Mukai, M. Misawa, Isolation and identification of camptothecin from cells of Camptotheca acuminata suspension cultures, Agr. Biol. Chem. 38 (1974) 217–218. [15] A.J. Van-Hengel, M.P. Harkes, H.J. Wichers, P.G.M. Hesselink, R.M. Buitelaar, Characterization of callus formation and camptothecin production by cell lines of Camptotheca acuminata, Plant Cell Tiss. Org. Cult. 28 (1992) 11–18. [16] A.J. Van-Hengel, R.M. Buitelaar, H.J. Wichers, Camptotheca acuminata Decne: in vitro culture and the production of camptothecin, in: Y.P.S. Bajaj (Ed.), Biotechnology in Agriculture and Forestry, Vol. 28, Medic-

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