Glandular trichomes on aerial and underground organs in Chrysolaena species (Vernonieae – Asteraceae): Structure, ultrastructure and chemical composition

Flora 207 (2012) 878–887

Contents lists available at SciVerse ScienceDirect

Flora journal homepage: www.elsevier.com/locate/flora

Glandular trichomes on aerial and underground organs in Chrysolaena species (Vernonieae – Asteraceae): Structure, ultrastructure and chemical composition Beatriz Appezzato-da-Glória a,∗ , Fernando Batista Da Costa b , Vanessa Cristina da Silva b , Leonardo Gobbo-Neto c , Vera Lucia Garcia Rehder d , Adriana Hissae Hayashi e a

Departamento de Ciências Biológicas, Escola Superior de Agricultura ‘Luiz de Queiroz’, Universidade de São Paulo (USP), 13418-900 Piracicaba, São Paulo, Brazil Departamento de Ciências Farmacêuticas, Faculdade de Ciências Farmacêuticas de Ribeirão Preto, USP, 14040-903 Ribeirão Preto, São Paulo, Brazil c Departamento de Física e Química, Faculdade de Ciências Farmacêuticas de Ribeirão Preto, USP, 14040-903 Ribeirão Preto, São Paulo, Brazil d Centro Pluridisciplinar de Pesquisas Químicas, Biológicas e Agrícolas, Universidade Estadual de Campinas, C.P. 6171, 13083-970 Paulínia, São Paulo, Brazil e Núcleo de Pesquisa em Anatomia, Instituto de Botânica, 04301-902 São Paulo, São Paulo, Brazil b

a r t i c l e

i n f o

Article history: Received 13 June 2012 Accepted 4 October 2012

Keywords: Chrysolaena obovata Chrysolaena platensis Sesquiterpene lactones Ultrastructure HPLC–UV–MS HPLC–UV–DAD

a b s t r a c t Although the occurrence of glandular trichomes is frequently reported for aerial vegetative organs, many questions still remain opened about the presence of such trichomes in underground systems. Here, we present, for the first time, a comparative study concerning the structure, ultrastructure and chemical aspects of both, the aerial and underground glandular trichomes of two different Chrysolaena species, C. obovata and C. platensis. Glandular trichomes (GTs) were examined using LM, SEM, and TEM and also analyzed by GC–MS and HPLC coupled to UV/DAD and HR-ESI-MS (HPLC–UV–MS). In both aerial (leaf and bud) and underground (rhizophore) organs, the GTs are multicellular, biseriate and formed by five pairs of cells: a pair of support cells, a pair of basal cells, and three pairs of secreting cells. These secreting cells have, at the beginning of secretory process, abundance of smooth ER. The same classes of secondary metabolites are biosynthesized and stored in both aerial and underground GTs of C. platensis and C. obovata. These GTs from aerial and underground organs have similar cellular and sub-cellular anatomy, however the belowground trichomes show a higher diversity of compounds when compared to those from the leaves. We also demonstrate by means of HPLC–UV–DAD that the sesquiterpene lactones are located inside the trichomes and that hirsutinolides are not artifacts. © 2012 Elsevier GmbH. All rights reserved.

Introduction Chrysolaena obovata (Less.) Dematt. and C. platensis (Spreng.) H.Rob. (Vernonieae – Asteraceae) are perennial herbaceous species from the Brazilian savanna with thickened underground stem called rhizophores (Hayashi and Appezzato-da-Glória, 2005). The rhizophores possess glandular trichomes in their axillary and terminal buds and internodes similar to what occurs in the aerial stem and leaves. For the thickened underground systems in Asteraceae species usually only internal secretory structures, such as cavities, ducts and laticifers, are described (Appezzato-da-Glória et al., 2008; Cury and Appezzato-da-Glória, 2009; Fritz and Saukel, 2011; Lotocka and Geszprych, 2004). The ducts of Santolina leucantha roots are

∗ Corresponding author. E-mail addresses: [email protected] (B. Appezzato-da-Glória), [email protected] (F.B. Da Costa), [email protected] (L. Gobbo-Neto), [email protected] (V.L.G. Rehder), [email protected] (A.H. Hayashi). 0367-2530/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.flora.2012.10.003

storage reservoirs for terpenoids, alkaloids, tannins and flavonoids that can act as allelopathic compounds or as deterrents (Pagni and Masini, 1999). The compounds present in the polyacetylene ducts of the primary root of Ambrosia trifida (Lersten and Curtis, 1988) and of Tagetes patula (Poli et al., 1995) have cytotoxic and antimicrobial properties. The cavities from the roots of Eupatorium rugosum (Lersten and Curtis, 1986) and from the rhizome and roots of Solidago canadensis (Curtis and Lersten, 1990) secrete lipids. The secondary metabolites secreted by the glandular trichomes from aerial organs are related to defense of plants against the attack of herbivores and pathogens or act as attractants to pollinators or for fruit dispersal (Ascensão et al., 1998, 1999; Heinrich et al., 2002; Wagner, 1991; Werker et al., 1994). Earlier works on the secondary metabolite chemistry of glandular trichomes from the aerial organs of Brazilian members of the tribe Heliantheae using phytochemical tools revealed a complete dominance of sesquiterpene lactones – STLs (Schorr et al., 2007; Spring et al., 2003; Stefani et al., 2003), some of which showing ecological function like antiherbivorous (Ambrósio et al., 2008) or other biological effects (e.g. anti-inflammatory: Arakawa et al., 2008; Schorr et al., 2002, 2007; Stefani et al., 2006). STLs from some

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

species of Vernonieae displayed moderate molluscicidal, antimicrobial and analgesic activities (Borkosky et al., 2009; Burim et al., 1999). Thus, besides ecological functions, STLs also reveal a great pharmacologic potential. The report of so many STLs in the trichomes of Heliantheae can be easily correlated to the occurrence of lipids (or lipophilic compounds) which are usually detected by classical histochemical methods and reported in many species. This approach had not yet been applied widely to members of Vernonieae, although STLs were recently reported in trichomes of Vernonia galamensis ssp. galamensis var. ethiopica (Favi et al., 2008). Lipids have been reported in biseriate glandular trichomes on aerial organs of Artemisia dracunculus (Werker et al., 1994) and Stevia rebaudiana (Monteiro et al., 2001). According to Werker and Fahn (1981), the secretion of lipids and polysaccharides by glandular trichomes of Inula viscosa may be related to the reduction of leaf temperature by increasing the reflection of light and protection against herbivores. In glandular trichomes of involucral bracts of Sigesbeckia jorullensis, the secretion has adhesive properties which favor the fruit dispersal by animals (Heinrich et al., 2002). Although the occurrence of glandular trichomes is frequently reported for aerial vegetative organs in Asteraceae, there are a few records of glandular trichomes present in underground systems (Appezzato-da-Glória et al., 2008) and many questions about the presence of such trichomes in these systems still remain open. Therefore, the objective of this study was to compare the structure and ultrastructure of glandular trichomes from aerial and underground organs of two species of Chrysolaena to verify similarities and differences between the secretions of these trichomes, including their chemical composition, regarding first of all secondary metabolites. Materials and methods Plant material Adult individuals of Chrysolaena obovata and C. platensis (Vernonieae) were collected in natural habitats in Cerrado areas in Botucatu (22◦ 53 S, 48◦ 29 W) and Mogi Guac¸u (22◦ 15 S, 47◦ 09 W), State of São Paulo, Brazil. The vouchers specimens (94148 and 94146, respectively) are deposited in the ESA Herbarium, Brazil. For structural and ultrastructural analyses, samples of the aerial parts (apical buds and mature leaves) and of underground stems – rhizophores (internodes, axillary and apical buds) – were analyzed. For phytochemical analyses, glandular trichomes were manually collected from mature leaves, shoot apical buds, and rhizophore axillary and apical buds. STLs used as reference compounds were previously isolated from Lepidaploa rufogrisea (A.St.-Hil.) H.Rob. (Vernonieae) which was collected in natural habitats in areas of Cerrado in São João Batista do Glória (20◦ 38 S, 46◦ 19 W), State of Minas Gerais, Brazil. A voucher specimen (LG037) is deposited in the UEC Herbarium, Brazil. Authorization for access to genetic patrimony was issued by Conselho Nacional de Desenvolvimento Científico e Tecnológico – CNPq (proc. number 010091/2011-4). Structural and ultrastructural analyses For LM analyses, samples were fixed in Karnovsky solution (Karnovsky, 1965), dehydrated in a graded ethanol series, and embedded in Leica Historesin® . Serial sections (7 ␮m thick) were cut on a rotary microtome and stained with toluidine blue O (Sakai, 1973). Permanent slides were mounted in synthetic resin. Hand-made cuts from fixed and fresh material were used for the following histochemical tests: Periodic acid-Schiff’s (PAS)

879

reaction to detect 1,2-glycol groups present in total polysaccharides (McManus, 1948); Sudan IV for lipophilic compounds (Johansen, 1940); ruthenium red for pectins (Johansen, 1940); Nile blue sulfate for neutral and acidic lipids (Cain, 1947); Nadi reagent for terpenoids (David and Carde, 1964); aniline blue black for total proteins (Fisher, 1968); zinc chloride iodine for starch (Strasburger, 1913). Samples were also fixed in formalin–ferrous sulfate reagent (Johansen, 1940) for detection of phenolic compounds. Standard control procedures were performed simultaneously. Photomicrographs were taken with a Leica® DM LB photomicroscope equipped with a Leica® DC 300F camera. For analysis with Calcofluor white for cellulose (Hughes and McCully, 1975) the microscope was equipped for epi-illumination with an HBO 50 mercury lamp and a Leica® D filter, providing excitation (Bandpass filter 355–425 nm) and suppression (Long-pass filter 470 nm). For SEM analyses, samples were fixed in Karnovsky solution, dehydrated in a graded ethanol series and critical point-dried with CO2 (Horridge and Tamm, 1969). The samples, mounted on stubs and coated with gold (30–40 nm), were examined under a LEO VP 435 SEM (Zeiss, Oberkochen, Germany) at 20 kV. For TEM investigation, the samples were fixed in Karnovsky solution, post-fixed in 1% OsO4 , incubated in an aqueous solution of 0.5% uranyl-acetate, dehydrated using a graded acetone series, and embedded in Araldite resin. The ultrathin sections were contrasted with uranyl acetate (Watson, 1958) and lead citrate (Reynolds, 1963), and examined under a Philips® TEM 100 microscope at 60 kV. Cytochemical analyses were performed using the ZIO technique (zinc iodine and osmium tetroxide) to observe membrane systems involved in the secretion. Samples were fixed in Karnovsky solution and incubated in a solution containing Zn, I, Tris–aminomethane and 2% OsO4 (Reinecke and Walther, 1978). Dehydration, embedding, polymerization and sectioning followed the above-cited protocol. Phytochemical analyses Gas chromatography–mass spectrometry (GC–MS) analysis Glandular trichomes, 120 from leaves and 200 from rhizophores of Chrysolaena platensis, were manually and individually collected using a stainless steel needle and a stereomicroscope (Stemi 2000C, Carl Zeiss, Jena, Germany) and their content was extracted with ethyl acetate in an ultrasonic bath at room temperature (ca. 26 ◦ C) for 15 min. The extracts were concentrated and submitted to GC–MS analysis, which was carried out with a Hewlett-Packard 6890 gas chromatograph (Hewlett Packard, Palo Alto, CA, USA) using an HP-FFAP column (50 m × 0.20 mm × 0.33 ␮m), applying the following conditions: injection temperature 220 ◦ C; detector temperature 250 ◦ C; column temperature 150 ◦ C (2 min), 5 ◦ C (1 min), 240 ◦ C, 10 ◦ C (1 min), 300 ◦ C (34 min); carrier gas: He (1.0 mL min−1 ); and split injection. The MS was operated in the EI mode at 70 eV in the m/z range from 42 to 350. Compounds were identified by comparison of their mass spectra with a mass spectral library database (NIST-05). The relative proportions of the constituents were expressed as percentage obtained by peak area normalization; all relative response factors were taken as one. HPLC–UV–MS analyses HPLC grade methanol (MeOH), acetonitrile (MeCN) and acetic acid (AcOH) were obtained from J.T. Baker. De-ionized water 18 m (Milli-Q, Millipore) was used throughout the study. For the HPLC analyses, 500 trichomes were collected from the leaves of C. platensis as described above and diluted in 100 ␮L of MeCN. Further 200 trichomes were collected from the rhizophores of C. platensis and the same amount from C. obovata, which were diluted in 40 ␮L of MeCN. In each analysis the injected volume was

880

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

20 ␮L, thus comprising 100 foliar or underground trichomes per injection. Analyses were carried out on a LC-20A liquid chromatographic system (Shimadzu Corporation, Japan) equipped with a photodiode array detector – DAD (CBM20A, Shimadzu Corporation, Japan) controlled by the software CLASS-VP v. 6.14, coupled to an ESI-qQ-TOF high resolution mass spectrometer UltrOTOFq (Bruker Daltonics, Billerica, MA, USA) controlled by the software DataAnalysis v. 3.2. Samples were chromatographed in two coupled C18 monolithic columns (Phenomenex, Onyx, 4.6 mm × 100 mm) which in turn were coupled with a guard-column (4.6/10.0 mm) of equivalent material, using the following gradient (flow rate 3.0 mL min−1 ): solvent A = aqueous 1% AcOH; solvent B = MeCN and 1% AcOH; elution profile = 0.01 min: 3% B; 3.0 min: 3% B; 60.0 min: 45% B; 63.0 min: 100% B; 66.0 min: 100% B; 68.0 min: 3% B; 70.0 min: 3% B. The column eluent was split at a ratio of 5:1, the larger flow going to the UV/DAD detector and the lower to the mass spectrometer. The UV/DAD detector was set to record between 210 and 600 nm, and HPLC–UV chromatograms were recorded at 225, 245, 270 and 325 nm. HPLC–MS total ion current (TIC) chromatograms were recorded between m/z 50 and 1000 in both positive and negative ionization modes and the following parameters were maintained in all analyses: 1000 scans/s, spectrum interval 2 s, drying gas flow 6.0 L min−1 , drying gas temperature 180 ◦ C, and nebulizer gas pressure 4 bar. N2 was used as drying and nebulizer gas. Chromatographic peaks identification (dereplication) was carried out based on UV, and MS spectra obtained for each peak. UV spectra were used to infer the secondary metabolite class of the compounds (e.g. STLs or phenolics). Accurate masses obtained for precursor ions (protonated [M+H]+ and or deprotonated molecules [M−H]− ) were used to calculate possible molecular formulae while CID fragmentation leading to product ion spectra (MS/MS) of selected ions were performed whenever possible for peak’s identity determination/confirmation. This process was performed mainly based on previous phytochemical studies of Chrysolaena species and, whenever possible, retention time comparison with authentic standards was carried out.

Phytochemical standards (reference compounds) isolation and structural elucidation

Results Structural and ultrastructural studies The indumentum of the aerial and underground systems of Chrysolaena obovata and C. platensis is composed by several nonglandular and glandular trichomes (Fig. 1A–C). The glandular trichomes occur on the lower surface of leaf and, in general, are located in depressions (Fig. 1C). The glandular trichomes are multicellular and biseriate, usually formed by five pairs of cells: a pair of support cells embedded in the epidermis, a pair of basal cells, and three pairs of secreting cells (Fig. 1D). In front view, the head of the trichome is bilobed (Fig. 1B and C). There are trichomes at different stages of secretion positioned side by side (Fig. 1E). The secretion accumulates gradually in the subcuticular space formed in the distal pair of secreting cells (Fig. 1D, E and G). At the beginning, the cuticle is attached to the cellular surface and is then gradually separated from it (Fig. 1F), becoming fully distended at the end of secretory process (Fig. 1H). In both Chrysolaena species the secretion is released to the plant surface after the disruption of the cuticle (Fig. 1I and J) that occurs along a predetermined line between the two cells at the top of the head of the trichome (Fig. 1J). In both vegetative organs (above and underground), the secreting cells (Figs. 2A and 3A) of the trichomes have, at the beginning of secretory process, dense cytoplasm, plastids (Fig. 2B), active dictyosomes (Fig. 2C) and many mitochondria which vary in electron density (Figs. 2D and E and 3B and C). The abundance of smooth endoplasmic reticulum (SER) is noteworthy (Figs. 2D and 3B and C). Under ZIO technique, the SER membranes appear electron-dense in the two upper pair of secreting cells (Figs. 2D and 3B) and electrontransparent (Fig. 2D and E) or sometimes electron-dense (Fig. 3C) in the subjacent secreting cells. The basal cells have dense cytoplasm and the nucleus contains a substantial amount of heterochromatin (Fig. 2F). The support cells are longer and more vacuolated (Fig. 2G). At the beginning, the secretion has a granulated appearance, but gradually, there is accumulation of lipid droplets (Fig. 3D) which prevail at the final stage (Figs. 1H, 3E and 4J). In the post-secretory stage, the cuticle breaks up while the trichome cells exhibit degradation signs, such as strong electron-density of the cytoplasm (Fig. 3E) and phenolic compound accumulation (Fig. 4J). Histochemistry

Intact leaves of Lepidaploa rufogrisea (460 g) were rinsed with acetone for 15 s under ultra-sonic bath. Solvent was removed under reduced pressure to give a leaf rinse extract (1.9 g), which was submitted to vacuum liquid chromatography (silica gel 60H Merck; 50 mm × 45 mm; eluent: n-hexane with increasing amounts of 10% acetone for each fraction) leading to 11 fractions. Fractions 11 (100% acetone) and 8 (n-hexane–acetone 3:7) were identified as the flavonoids isoquercetrin (13 mg) and isorhamnetin (264 mg) respectively (Harborne, 1994). Fraction 6 (n-hexane–acetone 5:5; 415 mg) was identified as the STL glaucolide B (Padolina et al., 1974). Fraction 5 (n-hexane–acetone 6:4; 52 mg) was submitted to a semi-preparative scale reverse-phase HPLC (Shimadzu LC-6AD coupled to the UV-DAD detector SPD-M10Avp; column Shimpack ODS 5 ␮m – Shimadzu, 250 mm × 20 mm) using an MeCN–H2 O elution gradient (from 10% MeCN up to 50% MeCN in 40 min; flow rate 9.0 mL min−1 ) to afford the following hirsutinolide type STLs: 8␣-acetoxy-10␣-hydroxy-13-O-methylhirsutinolide – 12.1 min; 6 mg (Bardón et al., 1993) and 8␣,13-diacetoxy-10␣hydroxyhirsutinolide – 26.6 min; 5 mg (Catalán et al., 1986). Though this is the first report of these two hirsutinolides from Lepidaploa rufogrisea, the first has already been reported from Chrysolaena platensis (Pollora et al., 2000) and the latter from other Vernonia (sensu lato) species (Catalán et al., 1986; Valdés et al., 1998).

Trichomes in both the aboveground shoot and the underground system present a translucent secretion (Fig. 4A and B) which stains positively with Sudan IV (Fig. 4C and D), indicating predominantly lipophilic substances. The positive reaction to the Nadi reagent indicates the presence of terpenoids (Fig. 4E). Neutral lipids (Fig. 4F) and acidic lipids (Fig. 4G) were shown by Nile blue sulfate in the trichomes. The histochemical tests with aniline blue black (Fig. 4H), zinc chloride iodine, ruthenium red (Fig. 4I) and PAS reaction had negative results. Samples fixed in formalin–ferrous sulfate to identify phenolic compounds (Fig. 4J) showed negative result for secretion but positive one for content of the secreting cells. Phytochemical analysis Due to the high amount of non-glandular trichomes which are present on the leaf surface of Chrysolaena obovata their glandular trichomes could not be collected. Therefore the trichomes of this species could not be analyzed by GC–MS and HPLC–UV–MS. Through analysis by GC–MS of the ethyl acetate extracts from trichomes of leaves and rhizophores of C. platensis it was possible to identify the presence of several compounds with lipophilic characteristics (Table 1). The trichome extracts of the leaves are composed mainly of long-chain compounds (9,17-octadecadienal,

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

881

Fig. 1. SEM (A–C, F, and I) and LM (D, E, G, H, and J) showing glandular trichomes of Chrysolaena obovata (A, B, E–H, and J) and C. platensis (C, D, and I). (A–C) Glandular (arrowed) and nonglandular trichomes on the lower surface of the leaf (A and C) and rhizophore (B). (D–F) Biseriate glandular trichome and its different development stages, respectively. The secretion accumulates gradually in the subcuticular space formed in the distal pair of secreting cells. The arrow shows the cuticle distension. (G) Wall of the distal cell (arrow) stained with Calcofluor white. (H) Cuticle fully distended at the end of secretory process. (I and J) Cuticle rupture for releasing secretion. Scale bars: (A) 150 ␮m, (B) 50 ␮m, (C) 60 ␮m, (D, E, G, H, and J) 30 ␮m, (F) 15 ␮m, and (I) 20 ␮m.

882

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

Fig. 2. TEM showing the ultrastructure of glandular trichomes from the leaf of Chrysolaena obovata. (A) Trichome with five pairs of cells: secreting cells (numbers 1–3), basal cells (number 4) and support cells (number 5). (B–E) Secreting cells showing plastids (B), dictyosomes (C), and mitochondria (m) with different electron density and abundance of smooth endoplasmic reticulum (ser) (D and E). (F) Basal cells whose nucleus presents substantial amount of heterochromatin. (G) Pair of support cells. Scale bars: (A) 4.6 ␮m, (B and D) 0.3 ␮m, (C) 0.5 ␮m, (E and F) 0.9 ␮m, and (G) 2 ␮m.

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

883

Fig. 3. TEM showing the ultrastructure of glandular trichomes from the leaf of Chrysolaena platensis during the secretory stage (A–D) and from the rhizophore of Chrysolaena obovata at the post-secretory stage (E). (A–C) Mitochondria (m), dictyosomes (d) and smooth endoplasmic reticulum (ser) in the secreting cells (numbers 1–3). (D) Granulated secretion with gradual accumulation of lipid droplets (arrows) in the subcuticular space. (E) Cuticle fully distended and ruptured. The secreting cells exhibit a strong electron-density of the cytoplasm (arrow). Scale bars: (A) 1.7 ␮m, (B) 0.9 ␮m, (C) 2 ␮m, (D) 0.3 ␮m, and (E) 8 ␮m.

octadecanoic methyl ester, and octadecanoic acid) and cholestadiene, while in the rhizophore trichomes the major compounds are terpenoids, in particular the diterpene phytol and the triterpene cholestadiene. In rhizophore trichomes long-chain compounds occur in lower concentration than in the trichomes of the leaves. The total amounts of identified compounds from leaves and rhizophores trichomes were 70.37 and 41.00%, respectively. The HPLC–UV–MS analyses of the glandular trichomes from leaves (Chrysolaena platensis) and rhizophores (C. obovata and C. platensis) revealed that their content is qualitatively similar to each other and that some peaks are shared by the trichomes from both species (Fig. 5). The information from the chromatograms is summarized in Table 2, which shows the seven main compounds that occur in both species. Though HPLC–UV–MS analyses were

performed in both positive and negative ionization, in the latter no ESI-MS detection was observed in TIC-MS chromatograms, thus affording no profitable information regarding the chemical composition. Three chromatographic peaks (17.5 min, 18.2 min and 31.1 min) were identified by combined comparison of their retention times, MS and UV spectra with the authentic standards isolated from Lepidaploa rufogrisea. Spectrometric data obtained for the chromatographic peak at 13.0 min (Fig. 5) suggested that it represents a metabolite closely related to the peaks at 17.5 and 18.2 min. The first peak has an UV spectrum that is identical to that of the latter. The peak fragmentation pattern at 13.0 min in the positive MS spectrum suggested an initial loss of neutral water, followed by the acetate moiety leading to m/z 319 and m/z 277, respectively and, inversely, losses of acetate followed by water

884

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

Fig. 4. Glandular trichomes from leaves (A, E, F, H, and I) and rhizophores (B–D, G, and J) of Chrysolaena obovata (A–H) and C. platensis (I and J) submitted to different stains and reagents. (A and B) In vivo (translucent secretion). (C–G) Positive staining of the secretion (arrows) by Sudan IV for lipids (C and D), Nadi reagent for terpenoids (E), Nile blue sulfate for neutral and acidic lipids (F and G). (H and I) Negative staining of the secretion by aniline blue black for proteins (H) and Ruthenium red for pectin (I). (J) Ferrous sulfate in formalin for phenolic compounds. Negative reaction for secretion and positive reaction for content of the secreting cells (arrow). Scale bars: (A, B, E, F, H–J) 30 ␮m, (C) 200 ␮m, and (D, and G) 60 ␮m.

leading to m/z 295 and m/z 277, respectively. Further fragmentations of m/z 277 are results of consecutive losses of neutral water and CO molecules. This is essentially the same fragmentation pattern observed for the peaks at 17.5 and 18.2 min, therefore starting at m/z 337 (parent ion of peak at 13.0 min), which is present in both 17.5 and 18.2 min peak MS spectra, and denotes only one

acetate moiety for the peak at 13.0 min. The molecular formula obtained by HRMS for the peak at 13.0 min consists of the same molecular formula of 17.5 less CH4 O. Analyzing MS fragmentation data together with the molecular formula, we suggest that the compound at 13.0 min is another hirsutinolide-type STL, consisting of the same basic peak structure at 17.5 min less the methyl group

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

885

Table 1 Constituents identified in extracts from leaves and rhizophore trichomes of Chrysolaena platensis and their relative percentages. Compounds

Leaves relative (%)

Rhizophores relative (%)

Hexadecanoic acid methyl ester 9,17-Octadecadienal (Z) 9-Octadecenoic methyl ester Phytol Octadecanoic methyl ester Octadecanoic acid Cholestadiene

1.05 25.50 3.91 nd 21.32 7.15 11.44

3.68 t 4.62 20.24 1.94 t 10.52

Total

70.37

41.00

Nd: not detected; t: trace.

at C13 (leaving there an OH) and less a water molecule (loss of OH at C1, leading to an double bound between C1 and C2). The chemical structures of the two identified hirsutinolides (peaks at 18.2 and 17.5 min, respectively) as well as the proposal for the third one (peak at 13.0 min) are displayed in Fig. 6. Discussion All glandular trichomes in aerial and underground systems of Chrysolaena obovata and C. platensis are biseriate as described in Vernonia galamensis (Vernonieae) by Favi et al. (2008) and other Asteraceae genera and tribes, such as in Helichrysum/Inuleae (Ascensão et al., 2001), Inula/Inuleae (Werker and Fahn, 1981) and Sigesbeckia/Heliantheae (Heinrich et al., 2002). In both Chrysolaena species investigated here, the secretion is released to the plant surface after the disruption of the cuticle that occurs along a predetermined line at the top of the head of the trichome. This cuticle rupture may occur after mechanical contact by herbivores or pathogens due to the presence of STLs, terpenes and flavonoids. Secretion storage in the subcuticular space and rupture of the cuticle were also observed in other Asteraceae species (Ascensão et al., 2001; Heinrich et al., 2002; Werker and Fahn, 1981; Werker et al., 1994). At the secretory stage, the three pairs of secreting cells exhibit large amounts of smooth endoplasmic reticulum and the

Fig. 5. Chromatograms from the trichomes of Chrysolaena platensis leaves (top), C. platensis rhizophores (middle) and Chrysolaena obovata rhizophores (bottom).

cytochemical affinity to ZIO, with heavy deposits of metal on the membranes, varies among the cells. Machado and Gregório (2001) also described reaction variations among impregnated dictyosome cisternae in plant secretory systems, but the probable cause of the affinity to ZIO is not known. The abundance of endoplasmic reticulum in the apical secretory cells of trichomes may be related

peak at 27.5 min

peak at 18. 2 min

peak at 13.0 min Fig. 6. Chemical structures of the hirsutinolides from Chrysolaena platensis (leaves and rhizophores) and C. obovata (rhizophores).

886

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887

Table 2 Retention times (RT), mass spectrometric (MS) data obtained in positive ionization mode, UV maximum absorptions (UV Max) and peak’s identity (PI) or molecular formulae (MF) of the compounds detected by HPLC–UV–MS in the glandular trichomes of the two investigated species of Chrysolaena. RT (min)

C. platensis (leaves)

C. platensis (rhizophores)

C. obovata (rhizophores)

MS and MS/MS data (m/z)

UV max (nm)

PI and/or MF

10.1 13.0

+

++ nd

nd nd

272 286

– C17 H20 O7

18.2

++

+

+

285

C18 H24 O8 8␣-acetoxy-10␣hydroxy-13-Omethylhirsutinolide

27.5

+++

+++

+++

285

31.1 45.9 50.3

+ nd nd

+ ++ +

nd nd +

– [M+H]+ 337.1285 319; 295; 277 b.p.; 259; 241; 231; 213; 199; 173 [M+H]+ 369.1547 351; 337; 309; 291 b.p.; 277; 259; 241; 231; 213; 199; 173 [M+H]+ 397.1483 379; 337; 319 b.p.; 277; 259; 241; 231; 213; 199; 173 [M+H]+ 317.0669 – –

C19 H24 O9 8␣,13-diacetoxy10␣hydroxyhirsutinolide Isorhamnetin – –

to the synthesis and secretion of lipids, as observed by Machado and Gregório (2001) in the extrafloral nectary of Citharexylum mirianthum (Verbenaceae). At the post-secretory stage, all cells of a trichome present strong electron-density and full cuticle expansion. These characteristics are very similar to those described in Vernonia galamensis ssp. galamensis var. ethiopica, suggesting degradation of the cells (Favi et al., 2008). Kristen and Lockhausen (1985) found in Veronica beccabunga (Plantaginaceae), the release of phenolic substances of the vacuole to the cytoplasm during the degeneration of glandular cells. In fact, the cells of the studied trichomes, at this stage, present phenolic compounds after fixing samples in formalin–ferrous sulfate reagent. In Grindelia pulchella (Asteraceae) secreting cells of glandular trichomes had the outer tangential wall with an alveolar aspect in post-secretion stage (Bartoli et al., 2011) but this feature was not verified in Chrysolaena obovata and C. platensis. Our findings indicate that basically the same classes of compounds are biosynthesized and stored in the aerial and underground parts of C. platensis and C. obovata. Nevertheless, the trichomes from the underground parts of both species showed a higher diversity of compounds when compared to those from the leaves as observed in the GC–MS (Table 1) as well as HPLC–UV–MS analyses (Table 2). Moreover, the HPLC–UV–MS analysis showed that the peaks at 18.2 and at 27.5 min (Fig. 5), which refer to two different hirsutinolide-type STLs, occur in the trichomes of both species and therefore can be stated as their chemical markers. Moreover, a flavonoid was also identified. Flavonoids are widespread in Asteraceae and comprise the most common class of secondary metabolites; nevertheless, they are somewhat unusual in Vernonieae (Emerenciano et al., 2001) and there are only a few reports on their occurrence in Vernonia (sensu lato). Some authors argue that possibly the main function of flavonoids inside the trichomes is to prevent other compounds from oxidation. STLs occur in many taxa from Vernonieae, including the subtribe Vernoniinae and the genus Vernonia (sensu lato), where the glaucolide and hirsutinolide subtypes are the most widespread (Herz, 1996). In Vernonia (sensu lato) and related genera glaucolides and hirsutinolides are considered to be chemical markers. Some authors believe that hirsutinolides are artifacts, i.e., products that are formed from glaucolides during the extraction/isolation procedures when the plant extract is in contact with silica gel and protic acidic solvents (like methanol or ethanol) (Herz, 1996). In our study, both species were treated neither with such solvents nor silica gel because each trichome was individually collected with a needle and solubilized in MeCN, being

255; 354 230 230

further analyzed in a C18 monolithic column. Moreover, the same detected STLs are present in both species, either in the leaves or in the rhizophores. Therefore, we strongly believe that these STLs are not artifacts. Similarly, the same observation was found for flavonoids in glandular trichomes of Lychnophora ericoides (Vernonieae), where direct MS analysis of glandular trichomes proved that acetylated flavonoids were not artifact products (Gobbo-Neto et al., 2008). The methodology described herein consisting of individual collection of trichomes followed by solubilization in MeCN was successful in our HPLC–UV–MS analyses providing unequivocal compound identification. Recently Favi et al. (2008) described the identification of a STL from the glaucolide subtype in the glandular trichomes of Vernonia galamensis ssp. galamensis var. ethiopica. Nevertheless, in that work the compound was detected in a leaf rinse extract obtained with ethanol, which was later compared to the trichome content; moreover, compound identification was carried out based only on the molecular formula obtained from MS. In this work, we demonstrated that the STLs are located inside the trichomes by means of HPLC–UV–DAD as well as MS fragmentation pattern analyses of their peaks. The analysis of the glandular content by GC–MS and HPLC–UV–MS from both aerial and subterranean organs corroborates the results from the histochemical analysis, therefore confirming the lipophilic (long-chain compounds, STLs, diterpene, and triterpene) as well as the phenolic products (flavonoid). Acknowledgements We thank Dr João Semir for plant identification, Dr Norberto P. Lopes for MS measurements, Dr Silvia R. Machado for MET assistance and NAP-MEPA, ESALQ, USP for electron microscope facilities. This work was supported by São Paulo Research Foundation – FAPESP (Project, Process number 06/51370-9) and National Council for Scientific and Technological Development – CNPq (Edital Universal 470750/2006-5) and research grants awarded to B.A.G., F.B.C. and A.H.H. References Ambrósio, S.R., et al., 2008. Constituents of glandular trichomes of Tithonia diversifolia: relationships to herbivory and antifeedant activity. Phytochemistry 69, 2052–2060. Appezzato-da-Glória, B., Hayashi, A.H., Cury, G., Soares, M.K.M., Rocha, R., 2008. Occurrence of secretory structures in underground systems of seven Asteraceae species. Bot. J. Linn. Soc. 157, 789–796.

B. Appezzato-da-Glória et al. / Flora 207 (2012) 878–887 Arakawa, N.S., Schorr, K., Ambrósio, S.R., Merfort, I., Da Costa, F.B., 2008. Further sesquiterpene lactones from Viguiera robusta and the potential antiinflammatory activity of a heliangolide: inhibition of human neutrophil elastase release. Z. Naturforsch. C 63c, 533–538. Ascensão, L., Figueiredo, A.C., Barroso, J.G., Pedro, L.G., Schripsema, J., Deans, S.G., Scheffer, J.C., 1998. Plectranthus madagascariensis: morphology of the glandular trichomes, essential oil composition, and its biology activity. Int. J. Plant Sci. 159, 31–38. Ascensão, L., Mota, L., Castro, M.M., 1999. Glandular trichomes on the leaves and flowers of Plectranthus ornatus: morphology, distribution and histochemistry. Ann. Bot. 84, 437–447. Ascensão, L., Silva, J.A.T., Barroso, J.G., Figueiredo, A.C., Pedro, L.G., 2001. Glandular trichomes and essential oils of Helichrysum stoechas. Isr. J. Plant Sci. 49, 115–122. Bardón, A., Montanaro, S., Catalán, C.A.N., Diaz, J.G., Herz, W., 1993. Piptocarphols and other constituents of Chrysolaena verbascifolia and Lessingianthus rubricaulis. Phytochemistry 34, 253–259. Bartoli, A., Galati, B.G., Tortosa, R.D., 2011. Anatomical studies of the secretory structures: glandular trichomes and ducts, in Grindelia pulchella Dunal (Astereae, Asteraceae). Flora 206, 1063–1068. Borkosky, S., Ponce de León, S., Juárez, G., Sierra, M.G., Bardón, A., 2009. Molluscicidal sesquiterpene lactones from species of the tribe Vernonieae (Compositae). Chem. Biodivers. 6, 513–519. Burim, R.V., Canalle, R., Lopes, J.L.C., Takahashi, C.S., 1999. Genotoxic action of the sesquiterpene lactone glaucolide B on mammalian cells in vitro and in vivo. Genet. Mol. Biol. 22, 401–406. Cain, A.J., 1947. The use of Nile blue in the examination of lipids. Quart. J. Micros. Sci. 88, 383–392. Catalán, C.A.N., De Iglesias, D.I.A., Kavca, J., Sosa, V.E., Herz, W., 1986. Sesquiterpene lactones and other constituents of Vernonia mollissima and Vernonia squamulosa. J. Nat. Prod. 49, 351–353. Curtis, J.D., Lersten, N.R., 1990. Oil reservoirs in stem, rhizome, and root of Solidago canadensis (Asteraceae, tribe Astereae). Nord. J. Bot. 4, 443–449. Cury, G., Appezzato-da-Glória, B., 2009. Internal secretory spaces in thickened underground systems of Asteraceae species. Aust. J. Bot. 57, 229–239. David, R., Carde, J.P., 1964. Coloration différentielle des inclusions lipidique et terpeniques des pseudophylles du Pin maritime au moyen du réactif de Nadi. C. R. Acad. Sci. Paris, Ser. D 258, 1338–1340. Emerenciano, V.P., Militão, J.S.L.T., Campos, C.C., Romoff, P., Kaplan, M.A.C., Zambon, M., Brandt, A.J.C., 2001. Flavonoids as chemotaxonomic markers for Asteraceae. Biochem. Syst. Ecol. 29, 947–957. Favi, F., Cantrell, C.L., Mebrahtu, T., Kraemer, M.E., 2008. Leaf peltate glandular trichomes of Vernonia galamensis ssp. galamensis var. ethiopica Gilbert: development, ultrastructure, and chemical composition. Int. J. Plant Sci. 169, 605–614. Fisher, D.B., 1968. Protein staining of ribboned epon sections for light microscopy. Histochemie 16, 92–96. Fritz, E., Saukel, J., 2011. Secretory structures of subterranean organs of some species of the Cardueae, and their diagnostic value. Acta Biol. Cracov. Bot. 53, 62–72. Gobbo-Neto, L., Gates, P.J., Lopes, N.P., 2008. Negative ion ‘chip-based’ nanospray tandem mass spectrometry for the analysis of flavonoids in glandular trichomes of Lychnophora ericoides Mart. (Asteraceae). Rapid Commun. Mass Spectrom. 22, 3802–3808. Harborne, J.B., 1994. The Flavonoids: Advances in Research since 1986. Chapman and Hall, London. Hayashi, A.H., Appezzato-da-Glória, B., 2005. The origin and anatomy of rhizophores in Vernonia herbacea and V. platensis (Asteraceae) from the Brazilian Cerrado. Aust. J. Bot. 53, 273–279. Heinrich, G., Pfeifhofer, H.W., Stabentheiner, E., Sawidis, T., 2002. Glandular hairs of Sigesbeckia jorullensis Kunth (Asteraceae): morphology, histochemistry and composition of essential oil. Ann. Bot. 88, 459–469. Herz, W., 1996. A review on the terpenoid chemistry of the Vernonieae. In: Hind, D.J.N., Beentje, H.J. (Eds.), Compositae: Systematics. Proc. Int. Compositae Conf., vol. 1. Kew, 1994. Royal Botanic Garden, Kew, pp. 229–251. Horridge, G.A., Tamm, S.L., 1969. Critical point drying for scanning electron microscopy study of ciliary motion. Science 163, 817–818. Hughes, J., McCully, M.E., 1975. The use of an optical brightener in the study of plant structure. Stain Technol. 50, 319–329.

887

Johansen, D.A., 1940. Plant Microtechnique. McGraw-Hill, New York. Karnovsky, M.J., 1965. A formaldehyde–glutaraldehyde fixative of high osmolality for use in electron microscopy. J. Cell Biol. 27, 137–138. Kristen, U., Lockhausen, J., 1985. The leaf glands of Veronica beccabunga L.: ultrastructure and a possible pathway of secretion. Isr. J. Bot. 34, 147–156. Lersten, N.R., Curtis, J.D., 1986. Tubular cavities in white snakeroot, Eupatorium rugosum (Asteraceae). Am. J. Bot. 73, 1016–1021. Lersten, N.R., Curtis, J.D., 1988. Secretory reservoirs (ducts) of two kinds in giant ragweed (Ambrosia trifida; Asteraceae). Am. J. Bot. 75, 1313–1323. Lotocka, B., Geszprych, A., 2004. Anatomy of the vegetative organs and secretory structures of Rhaponticum carthamoides (Asteraceae). Bot. J. Linn. Soc. 144, 207–233. Machado, S.R., Gregório, E.A., 2001. Zinc iodide–osmium tetroxide (ZIO) reactive sites in the extrafloral nectary of Citharexylum mirianthum Cham. (Verbenaceae). Tissue Cell 33, 72–77. McManus, J.F.A., 1948. Histological and histochemical uses of periodic acid. Stain Technol. 23, 99–108. Monteiro, W.R., Castro, M.M., Mazzoni-Viveiros, S.C., Mahlberg, P.G., 2001. Development and some histochemical aspects of foliar glandular trichomes of Stevia rebaudiana (Bert.) Bert. – Asteraceae. Rev. Bras. Bot. 24, 349–357. Padolina, W.G., et al., 1974. Glaucolide-A and -B, new germacranolidetype sesquiterpene lactones from Vernonia (Compositae). Tetrahedron 30, 1161–1170. Pagni, A.M., Masini, A., 1999. Morphology, distribution, and histochemistry of secretory structures in vegetative organs of Santolina leucantha Bertol. (Asteraceae). Isr. J. Plant Sci. 47, 257–263. Poli, F., Sacchetti, G., Bruni, A., 1995. Distribution of internal secretory structures in Tagetes patula (Asteraceae). Nord. J. Bot. 5, 197–205. Pollora, G.C., Bardón, A., Catalán, C.A.N., Gedris, T.E., Herz, W., 2000. Sesquiterpene lactones from Chrysolaena platensis. Biochem. Syst. Ecol. 28, 707–711. Reinecke, M., Walther, C., 1978. Aspects of turnover and biogenesis of synaptic vesicles at locust neuromuscular junctions as revealed by iodide–osmium tetroxide (ZIO) reacting with intravesicular sh-groups. J. Cell Biol. 78, 839– 855. Reynolds, E.S., 1963. Use of lead citrate at high pH as an electron-opaque stain in electron microscopy. J. Cell Biol. 17, 208–212. Sakai, W.S., 1973. Simple method for differential staining of paraffin embedded plant material using toluidine blue O. Stain Technol. 48, 247–249. ˜ A.J., Siedle, B., Merfort, I., Da Costa, F.B., 2002. Guaianolides Schorr, K., García-Pineres, from Viguiera gardneri inhibit the transcription factor NF-␬B. Phytochemistry 60, 733–740. Schorr, K., Merfort, I., Da Costa, F.B., 2007. A novel dimeric melampolide and further terpenoids from Smallanthus sonchifolius (Asteraceae) and the inhibition of the transcription factor NF-␬B. Nat. Prod. Commun. 2, 367–374. Spring, O., Zipper, R., Conrad, J., Vogler, B., Klaiber, I., Da Costa, F.B., 2003. Sesquiterpene lactones from glandular trichomes of Viguiera radula (Heliantheae; Asteraceae). Phytochemistry 62, 1185–1189. Stefani, R., Eberlin, M.N., Tomazela, D.M., Da Costa, F.B., 2003. Eudesmanolides from Dimerostemma vestitum. J. Nat. Prod. 66, 401–403. Stefani, R., Schorr, K., Tureta, J.M., Vichnewski, W., Merfort, I., Da Costa, F.B., 2006. Sesquiterpene lactones from Dimerostemma species (Asteraceae) and in vitro potential anti-inflammatory activities. Z. Naturforsch. C 61c, 647– 652. Strasburger, E., 1913. Handbook of Practical Botany, 7th ed. George Allen, London. Valdés, D.A., Bardón, A., Catalán, C.A.N., Gedris, T.E., Herz, W., 1998. Glaucolides, piptocarphins and cadinanolides from Lepidaploa remotiflora. Biochem. Syst. Ecol. 26, 685–689. Wagner, G.J., 1991. Secreting glandular trichomes: more than just hairs. Plant Physiol. 96, 675–679. Watson, M.L., 1958. Staining of tissues sections for electron microscopy with heavy metals. J. Biophys. Biochem. Cytol. 4, 475–478. Werker, E., Fahn, A., 1981. Secretory hairs of Inula viscosa (L.) Ait. – development, ultrastructure, and secretion. Bot. Gaz. 142, 461–476. Werker, E., Putievsky, E., Ravid, U., Dudai, N., Katzir, I., 1994. Glandular hairs, secretory cavities, and the essential oil in leaves of tarragon (Artemisia dracunculus L.). J. Herbs Spices Med. Plants 2, 19–32.