JXB Advance Access originally published online on November 9, 2006
Journal of Experimental Botany 2006 57(15):4189-4200; doi:10.1093/jxb/erl195
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RESEARCH PAPER |
Pre-haustorial resistance to broomrape (Orobanche cumana) in sunflower (Helianthus annuus): cytochemical studies
1Agricultural and Plant Biochemistry Research Group, Department of Biochemistry and Molecular Biology, University of Córdoba, Campus de Rabanales, Edificio Severo Ochoa, E-14071 Córdoba, Spain
2IFAPA-CICE (Junta de Andalucía), CIFA ‘Alameda del Obispo’, Área de Mejora y Biotecnología, Apdo. 3092, E-14080 Córdoba, Spain
* To whom correspondence should be addressed. E-mail: bb2maala{at}uco.es
Received 16 May 2006; Accepted 11 September 2006
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Abstract |
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Sunflower broomrape (Orobanche cumana Wallr.) is a root holoparasitic angiosperm considered as one of the major constraints for sunflower production in Mediterranean areas. Breeding for resistance is regarded as the most effective, feasible, and environmentally friendly solution to control this parasite. However, the existing sources of genetic resistance are defeated by the continuous emergence of new more virulent races of the parasite. In this work, the interaction between sunflower and O. cumana has been analysed in order to gain insights into the mechanisms involved in resistance. Two sunflower genotypes were selected showing different behaviour against the new race F of O. cumana, HE-39998 (susceptible) and HE-39999 (resistant), and both compatible and incompatible interactions were compared. Pot and Petri dish bioassays revealed that only HE-39998 plants were severely affected, supporting a high number of successfully established broomrapes to mature flowering, whereas in HE-39999 root tubercles were never observed, resistance being associated with browning symptoms of both parasite and host tissues. Histological aspects of the resistance were further investigated. Suberization and protein cross-linking at the cell wall were seen in the resistant sunflower cells in contact with the parasite, preventing parasite penetration and connection to the host vascular system. In addition, fluorescence and confocal laser microscopy (CLM) observations revealed accumulation of phenolic compounds during the incompatible reaction, which is in agreement with these metabolites playing a defensive role during H. annuus–O. cumana interaction.
Key words: Confocal laser microscopy, defence responses, Helianthus annuus, histology, Orobanche cumana, parasitic plants, phenolics, protein cross-linking, resistance, suberization
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Introduction |
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Sunflower broomrape, Orobanche cumana Wallr., is considered as one of the major constraints for sunflower production in Mediterranean regions, including Eastern Europe and Spain (Alonso et al., 1996; Shindrova et al., 1998). This parasitic angiosperm depends entirely on the host for its supply of water and nutrients. Upon infection of sunflower (Helianthus annuus L.) roots, O. cumana successfully competes for sunflower nutritional resources, thereby damaging crop development and reducing yields drastically, by up to 50% (Parker and Riches, 1993; Domínguez, 1996). Several methods have been proposed to control sunflower broomrape, including mechanical, biological, and chemical practices, but nowadays genetic resistance is regarded as the most effective, feasible, economically and environmentally friendly solution. However, the use of resistant cultivars, usually of monogenic nature, is followed by the appearance of new more virulent races of the parasite that overcome the existing sources of resistance. This is the case of the new race F of O. cumana which has recently appeared in the south of Spain (Domínguez, 1999). Resistance to this new race has been found (Domínguez, 1999; Jan et al., 2002), and quantitative trait loci (QTLs) associated with this trait have been identified (Pérez-Vich et al., 2004). Sunflower resistance to parasitic plants seems to be a complex multicomponent process and, consequently, in order to achieve an optimal durable and effective resistance, the combination of several resistance elements in one cultivar would be desirable. Hence, a detailed knowledge of sunflower–O. cumana interaction and the mechanisms underlying resistance is mandatory to reach this goal.
The Orobanche spp. biological cycle comprises well-defined steps, separated both spatially and temporally, that are potential targets for host defence strategies. Upon germination, stimulated by host root-exuded chemical signals, broomrape seed develops a small seedling that attaches to the host root and differentiates in the attachment organ (appressorium). After host tissue penetration and connection to the vascular system through the haustorium, the parasite becomes a major sink for plant photosynthates, gradually forming a tubercle from which a shoot arises to emerge from the soil to flower and produce seeds (Parker and Riches, 1993; Westwood, 2000). Although much is known about the parasite's biological cycle on susceptible hosts, the molecular bases of host resistance to this parasite remain to be elucidated. Most of the studies have focused on identifying the host signals that induce germination and appressorium formation (Yoder, 1999; Galindo et al., 2002; Bouwmeester et al., 2003). Several reports reveal that typical plant defence responses against pathogenic micro-organisms are also induced in response to parasitic plant infection (Joel and Portnoy, 1998; Westwood et al., 1998; Goldwasser et al., 1999) such as phytoalexin induction (Serghini et al., 2001), high levels of peroxidase activities (Goldwasser et al., 1999; Pérez-de-Luque et al., 2005a), lignification of host endodermis (Perez-de-Luque et al., 2005c), induction of pathogenesis-related (PR) proteins (Joel and Portnoy, 1998; Pérez-de-Luque et al., 2006a), and sealing of host xylem vessels by deposition of mucilage (Pérez-de-Luque et al., 2005c, 2006b). Callose deposition and reactive oxygen species (ROS) production and accumulation are also typical responses to biotic and abiotic stresses and are involved in the interaction between plant-parasitic plants (Pérez-de-Luque et al., 2006a). Several mechanisms involved in resistance of sunflower to O. cumana have been identified, each interrupting broomrape development at different stages. Those include excretion of phytoalexins (Serghini et al., 2001), cell wall deposition, vessel occlusion, development of an encapsulation layer in the cortical parenchyma, and lignification of host xylem vessels (Dörr et al., 1994; Labrousse et al., 2001). A recent work, using a proteomics approach to investigate the pea–O. crenata interaction, allowed the identification of several proteins involved in resistance such as peroxidases and typical PR proteins, and revealed that carbon and nitrogen metabolism are affected in relation to pea resistance/susceptibility to broomrape (Castillejo et al., 2004).
The aim of the present work is to identify defence responses involved in conferring resistance to broomrape, an essential step for the development of effective sunflower breeding programmes. For this purpose, advantage was taken of the genetic variability available for sunflower, and two genotypes were used, HE-39998 (susceptible) and HE-39999 (resistant), which are very close genetically but display different behaviour against the new race F of O. cumana (Advanta Ibérica, S.A., personal communication). Both compatible and incompatible reactions were compared. Data are discussed and compared with previous data obtained in legumes (Perez de Luque et al., 2005a, b, c, 2006a) and sunflower (Serghini et al., 2001).
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Materials and methods |
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Plant material, growth conditions, and inoculation
The sunflower genotypes HE-39999 and HE-39998 and O. cumana seeds (race F) used in this study were kindly provided by Advanta Ibérica S.A. (Sevilla, Spain). Sunflower lines were selected on the basis of differences in resistance to O. cumana race F as previously tested by field experiments: resistant HE-39999 and susceptible HE-39998 (Advanta, personal communication).
Orobanche cumana seeds, race F ‘El Rubio’, belong to a homogeneus population and were collected from sunflower-infected fields in southern Spain in 2000. Once cleaned, seeds were kept in darkness at room temperature. The viability of Orobanche seeds was determined by the 2,3,5-triphenyl tetrazolium chloride (TTC) test (López-Granados and García-Torres, 1996). Germination tests with the synthetic germination stimulant GR-24 were performed as previously reported (Pérez de Luque et al., 2000), the in vitro percentage germination being 
82%.
Sunflower seeds, previously disinfected with sodium hypochlorite (20%, for 20 min), were germinated in Petri dishes (15 cm diameter) on wet paper and kept in the dark at 22 °C for 5–6 d (until the radicle reached 3 cm in length).
The characterization of the resistant/susceptible phenotype of sunflower genotypes was conducted by pot and Petri dish infection experiments. During all the experiments, plants were kept in a controlled environment chamber with a day/night temperature of 25/18 °C, 12 h photoperiod, and an irradiance of 300 µmol m–2 s–1, and watered regularly with Hoagland nutrient solution (Hoagland and Arnon, 1950).
For pot experiments, seedlings were transferred to trays filled with perlite and grown until the root was 5–7 cm long. Inoculation was carried out in eight plants by sprinkling O. cumana seeds (30 mg) on the surface of previously washed sunflower roots. Inoculated plants were then placed into pots with sand and vermiculite (3:1 v:v) as substrate and the infection process was evaluated. Established broomrapes were quantified and classified according to their different development stages from day 41 to day 80 after inoculation, when the experiment was concluded, and expressed as absolute value per plant. Non-infected plants were grown and evaluated in parallel.
For dish experiments, the Petri dish system described by Pérez-de-Luque et al. (2005a) was used. Sunflower seedlings (3 cm radicle in length) were transferred to Petri dishes (15 cm diameter) filled with perlite and covered with glass fibre papers (Whatmann GF/A). After 7–9 d, inoculation was performed by changing the glass fibre paper for one on which O. cumana seeds (10 mg) were homogeneously spread, and conditioned with MES 0.3 mM in darkness at 20 °C for 10 d. Dishes were sealed with parafilm, covered with aluminium foil to exclude the light, and placed vertically, the germinating host plant upwards, in trays with Hoagland nutrient solution (Hoagland and Arnon, 1950).
Infection was monitored daily under a binocular microscope (Leica L2; Leica Microsystems Wetzlar GmbH, Wetzlar, Germany). At least 200 Orobanche seeds close (<3 mm) to the sunflower roots were visualized in each Petri dish. Quantification of O. cumana seed germination, attachments, and tubercles was performed at 7, 10, 14, 17, 21, and 24 days post-inoculation (DPI) and expressed as the percentage of the total number of visualized seeds, the percentage of the total number of germinated seeds, and the absolute number per Petri dish, respectively. Seeds having an emerged seedling were scored as germinated and seedlings contacting a host root were recorded as attached. Images were taken with a digital camera (Leica DC 500, Leica Microsystems). Ten plants per genotype (HE-39999 and HE-39998) and treatment (infected and control) were used. The whole experiment was repeated three times.
Collection and fixation of samples for conventional microscopy
The Petri dish system was used for plant material collection. Observations were taken using a binocular microscope (Leica L2). Two independent experiments were set, each of them consisting of 12 plants per genotype (HE-39999 and HE-39998). At 11, 14, and 17 DPI, seedlings of O. cumana were sampled with the corresponding attached parts of host roots. In parallel, sunflower roots from control (non-infected) plants were sampled.
For cytochemical analysis of bright field and epi-fluorescence observations by light microscopy, the sampled material (10 samples for each staining procedure) was fixed in FAA (50% ethanol+5% formaldehyde+10% glacial acetic acid, in water) for 48 h. Fixed samples were then dehydrated in an ethanol series (50, 80, 95, 100, 100%: 12 h each), transferred to an embedding solvent (Xylene; Panreac Quimica S.A., Montcada i Reixac, Spain) through a xylene–ethanol series (30, 50, 80, 100, 100%: 12 h each), and finally saturated with paraffin (Paraplast Xtra; Sigma, St Louis, MO USA). Sections of 7 µm thickness were cut with a rotary microtome (Nahita 534; Auxilab S.A., Beriain, Spain) and attached to adhesive-treated microscope slides (polysine slides; Menzel GmbH & Co KG, Braunschweig, Germany).
Ten samples were kept after fixation in FAA and used for protein cross-linking determination, and some others were used fresh for determination of phenolic compounds, H2O2, and peroxidase activity.
Cytochemical methods for light microscopy
After removal of paraffin, the sections were stained with different dyes: (i) Alcian green–safranin (AGS) (Joel, 1983). The slides were dried and mounted with DePeX (BDH). With this staining method, carbohydrates (including cell walls and mucilage) appeared green, yellow, or blue, while lignified, cutinized, and suberized walls, as well as tannin and lipid material inside the cells, appeared red (Joel, 1983). (ii) Phloroglucinol (2% in ethanol)–HCl (35%) (Ruzin, 1999) stains the aldehyde groups of lignin and suberin, but quenches lignin autofluorescence and retains suberin fluorescence (Baayen et al., 1996; Rioux et al., 1998). (iii) Aniline blue fluorochrome was used for callose detection under UV fluorescence (340–380 nm). The samples were stained for 15–30 min in a solution of 0.1% aniline blue fluorochrome in water (Bordallo et al., 2002). (iv) Protein cross-linking in cell walls was determined following the procedure described by Mellersh et al. (2002). Fixed samples were cut by hand and submerged in 1% SDS for 24 h at 80 °C. They were stained for 3–5 min in 0.1% Coomassie blue in 40% ethanol/10% acetic acid, rinsed in a solution of 40% ethanol/10% acetic acid, and mounted in distilled water. Cell walls with protein cross-linking show a deep blue colour. (v) For determination of H2O2 and peroxidase activity, fresh samples were stained with 3,3-diaminobenzidine (DAB) using a modification of the procedure described by Thordal-Christensen et al. (1997). Fresh samples were submerged in DAB solution (1 mg ml–1, pH 3.8) prepared in distilled water for 2–3 h. After that, the samples were washed with lactic acid, glycerol, and water (1:1:1) for 1 h, hand cut with a razor blade, and mounted on slides with lactic acid, glycerol, and water (1:1:1 by vol.). (vi) Accumulation of phenolic compounds was determined on fresh hand-cut samples observed under epi-fluorescence (340–380 nm).
The sections were observed using a light microscope (Leica DM-LB; Leica Microsystems) and photographed at a magnification of x100 to x400 using a digital camera (Nikon DXM1200F; Nikon Europe B.V., Badhoevedorp, The Netherlands). The samples were also observed by epi-fluorescence under excitation at 340–380 and 450–490 nm.
Confocal microscopy
Ten fresh samples of O. cumana seedlings with the corresponding attached parts of the host roots were immersed in a solution of 0.1% (w/v) diphenyl boric acid 2-aminoethylester (Naturstoffreagenz A; NA) in buffer [100 mM KPi, pH 6.8, 1% NaCl (w/v)] according to Hutzler et al. (1998). This treatment induces secondary fluorescence of flavonoids. Confocal optical section stacks were collected using a Leica TCS-SP2-AOBS-UV confocal laser scanning microscope (Leica Microsystems) with an excitation wavelength of 488 nm and emission spectra monitored from 515 to 565 nm. Analysis of confocal images was performed with the LEICA software LCS, version 2.5 Build 1227.
Statistical analysis
Each Petri dish assay was performed with 10 replicates per genotype and treatment. The pot assay was carried out with eight replicates per genotype. The Student's t-test was performed for both assays and the P-value calculated. Percentages and whole numbers were transformed according to the formulae Y=arcsin(
(X(%)/100) and Y=(X+1/2), respectively.
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Differences in resistance to broomrape between sunflower genotypes HE-39999 and HE-39998
Differences in resistance to broomrape between the sunflower genotypes studied were evaluated by pot and Petri dish experiments (Table 1; Fig. 1). Quantification and classification of O. cumana tubercles, shoots, and flowering stalks were performed during the pot bioassay. At 41 DPI, one inoculated plant of each genotype was uprooted to quantify the underground established broomrapes. Thirty-eight tubercles and 21 stalks were found to be attached to the root of the HE-39998 plant, while no established broomrapes were observed on the root of the HE-39999 sunflower plant. The first emerged broomrape shoot was detected on HE-39998 plants at 47 DPI (Fig. 1A) and the first flowered broomrapes appeared at 60 DPI, the average number of emerged shoots per plant at this time being between eight and 10 (Fig. 1B). At 80 DPI, the experiment was concluded, plants were uprooted, and all established broomrapes quantified and classified according to their developmental stage. Each HE-39998 plant supported an average of 49 successfully established broomrapes, most of them being emerged shoots (19) or at the flowering stage (22) (Table 1). On the other hand, a detailed examination of roots of the HE-39999-infected plants revealed no established broomrapes.
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The two genotypes were further evaluated for differences in resistance against O. cumana at earlier times after infection using the Petri dish inoculation bioassay. The interaction was monitored by observations under a binocular microscope, annotating broomrape seed germination, attachment, and tubercle formation (Fig. 1G–L; Table 1). Germination of O. cumana seeds started to be visible at 4 DPI, once a conditioning period of
10 d required for germination was completed. The average percentage of O. cumana seeds germinated near the host roots of HE-39999 and HE-39998 plants was 93% and 90%, respectively (Table 1), a figure slightly higher than that obtained for the in vitro germination assays conducted with the synthetic germination stimulant GR-24. Microscopic observations revealed that O. cumana development is arrested at the first stages of the parasite life cycle in the incompatible interaction, and this is highly associated with darkening of the seedling of the parasite (Fig. 1G, H). These symptoms were already evident at 7 DPI, when most seedlings in contact with the root of HE-39998 displayed a translucent appearance, while the seedlings in contact with roots of resistant HE-39999 plants appeared darker. At 14 DPI, the percentage of germinated O. cumana seeds attached to the roots is similar for the resistant sunflower (33%) and the susceptible genotype (29%) (Table 1). Nevertheless, attachments in the incompatible interaction had a brownish aspect that extended towards the host tissues (Fig. 1G–I), while a root swelling caused by the development of the parasite can be detected in the compatible interaction (Fig. 1J, K). The first tubercles appeared at 14 DPI only in the susceptible genotype, the average number per plant, quantified at 24 DPI, being 89 (Table 1; Fig. 1L), which continued their development to emerge as a shoot (Fig. 1E).
Cytochemical analyses
Cytochemical analyses were performed on sections of infected roots of both genotypes containing an attached seedling. First, these sections were stained with AGS revealing that, during the incompatible interaction of O. cumana with the resistant sunflower plants, the intrusive cells of the parasite are stopped in the host cortex, before reaching the endodermis (Fig. 2A). In some cases, the parasite is scarcely able to pierce the epidermis (Fig. 2B). In contrast, on the roots of the susceptible sunflower accession, the infection developed normally, with the intrusive cells reaching the central cylinder and the host vascular tissues (Fig. 2D). A more detailed observation of the incompatible interactions showed that the host cells in contact with the parasite intrusive cells presented a thickening of their walls that stained intensely red with safranin. This reddish coloration was also observed at the intercellular spaces, the interface between both organisms, and inside the parasite cells (Fig. 2C). The thickened host cell walls did not show refringence when observed under polarized light, in contrast to lignified host xylem walls (Fig. 2E). However, an intense yellow fluorescence was observed at the infection point under excitation of 450–490 nm that co-localized with the safranin red staining (Fig. 2G). None of these was observed in sections of the compatible interaction (Fig. 2F, H).
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In order to uncover the nature of the host cell wall thickening, phloroglucinol-stained sections were observed under light and fluorescence microscopes. Xylem walls acquired a characteristic pink staining for lignins, and the host cell walls in contact with the parasite and parasite intrusive cells also showed this pink coloration inside, indicating the presence of compounds of a phenolic nature (Fig. 3A). Observation of these sections under a fluorescence microscope (340–380 nm) showed an intense blue fluorescence at the infection point that was absent from the xylem cell walls (quenched by the staining) (Fig. 3B), strongly suggesting that suberins are responsible for the cell wall thickening observed.
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Aniline blue fluorochrome staining was used to detect callose accumulation at the infection site. Some scarce callose depositions were detected in root sections of resistant plants close to but outside of the infection point (Fig. 3D).
However, protein cross-linking was observed in host cell walls at the infection point, in contact with O. cumana tissues during incompatible interactions (Fig. 3E, G). Soluble proteins were previously extracted with the SDS treatment, and only those proteins strongly linked to cell walls remained and were stained by the dye. The blue staining was present in the walls of host cells next to parasite tissues, being absent in the case of compatible interactions (Fig. 3F, H).
Samples corresponding to compatible and incompatible interactions were stained with DAB for the detection of H2O2 and peroxidase activity. Data were not conclusive; a strong staining was detected at the interface between the host and parasite in both compatible and incompatible interactions (data not shown).
Accumulation of phenolic compounds was analysed by fluorescence and confocal laser microscopy (CLM). Observation of fresh hand-cut samples under the appropriate light (excitation wavelength of 340–380 nm) showed an intense blue fluorescence inside parasite intrusive cells and neighbouring host tissues during incompatible interactions (Fig. 4C), which was not observed in the case of compatible interactions (Fig. 4D). CLM analysis revealed the accumulation of intense fluorescence due to phenolic compounds in the parasite and host tissues, as well as at the interface between both organisms during incompatible interactions (Fig. 5A, C, E). At the early stages of the infection process, phenolics were detected in parasite tissues (attachment organ) in contact with the resistant host root (Fig. 5A). Once the parasite penetrates the host, accumulation of phenolic compounds was observed in host cortical cell walls in contact with parasite intrusive cells, in the apoplast, at the host–parasite interface, and again inside parasite tissues (Fig. 5E, C). By contrast, this fluorescence was absent in every one of the compatible interactions analysed (Fig. 5D, F).
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Discussion |
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A range of host resistance strategies to prevent parasite infection have been documented, each interrupting specific stages of the parasite biological cycle (Jorrín et al., 1999; Labrousse et al., 2001). The observed results suggest that resistance in line HE-39999 must operate during the early stages of the parasite life cycle. The differences observed between both genotypes with respect to the capability of inducing O. cumana seed germination is unlikely to account for the differences in resistance. Thus, it could be possible that resistance in the HE-39999 genotype, associated with darkening of host and parasite tissues at the infection site, operates during host root tissue penetration, preventing connection to the vascular system, and hence broomrape development (Fig. 1; Table 1).
Previous works investigating the mechanisms of resistance to Orobanche spp. with other plant species have shown that the parasite intrusion can be stopped in the cortex (Pérez-de-Luque et al., 2006a), at the endodermis (Pérez-de-Luque et al., 2005c), or after reaching the central cylinder (Pérez-de-Luque et al., 2005b). In the present study with sunflower, the resistant accession HE-39999 does not allow parasite penetration further than the cortex (Fig. 2). However, the mechanisms described here are different from those reported previously for legume–O. crenata interaction (Pérez-de-Luque et al., 2005c, 2006a). HE-39999 develops a thickening by suberization of the cell walls in contact with O. cumana intrusive cells (Fig. 2). At the same time, reinforcement of nearby host cell walls by protein cross-linking takes place. As a result of these actions, the parasite is unable to continue further penetration and its establishment and development are prevented. Suberization of host cell walls as a mechanisms preventing root parasitic plant penetration has not been described previously, and only Zaitoun et al. (1991) reported the development of a ‘corky tissue’ in faba bean (Vicia faba) resistant to O. crenata. Dörr et al. (1994) reported the thickening of sunflower cell walls as a resistant response to O. cumana infection and the formation of an ‘isolation layer’ around the parasite tissues. More recently, Pérez-de-Luque et al. (2006a) showed protein cross-linking as a defensive mechanism against O. crenata in pea (Pisum sativum). Therefore, to date, only lignification and protein cross-linking have been reported as mechanisms responsible for cell wall reinforcement against root parasitic plant intrusion (Pérez-de-Luque et al., 2005c, 2006a). On the other hand, suberization of tissues has been reported as a response to wounds (Lulai and Corsini, 1998; Polito et al., 2002), vessel invasion by vascular pathogens (Rioux et al., 1995; Beckman, 2000; Lulai, 2005), and compartmentalization against intruding fungi (Baayen et al., 1996; Rioux and Baayen, 1997; Reissinger et al., 2003), this last case being the one which shares more similarities with the phenomenon observed here.
Peroxidases belong to a large family of enzymes to which a variety of functional roles have been ascribed, including lignification and cell wall phenol deposition, suberization, developmental-related processes, defence against pathogens, and response to other stresses. The formation of protein cross-links and of papillae, two types of cell wall fortification which enhance cell wall resistance within just a few minutes after pathogen attack (Bradley et al., 1992), are processes accomplished by peroxidases in the presence of H2O2 (Hammond-Kosack and Jones, 1996; Brown et al., 1998). The key role of peroxidases in the process of cell wall reinforcement in relation to resistance to parasitic plants has been demonstrated previously in several pathosystems (Bradley et al., 1992; Hammond-Kosack and Jones, 1996; Brown et al., 1998). An increase in lignification and peroxidase activity has been observed in vetch plants infected with O. aegyptiaca as part of defence mechanisms conferring mechanical and chemical barriers confronting the invading parasite (Goldwasser et al., 1999). Recent works using suppression-subtractive hybridization approaches (Vieira Dos Santos et al., 2003) and proteomic techniques (Castillejo et al., 2004) have led to the identification of several genes encoding peroxidases expressed during the process of resistance to Orobanche spp. In addition, in situ hybridization studies have shown the expression of a peroxidase gene in cells of pea plants resistant to O. crenata. Its expression was restricted to those cells near to the penetration point, which is in agreement with peroxidase activity data and the H2O2 accumulation pattern (Pérez-de-Luque et al., 2006a). In spite of the above strong evidence supporting the implication of peroxidases in plant resistance to parasitic plants, histochemical differences in peroxidase activity between genotypes could not be detected in the present study because an intense staining was observed in both cases (resistant and susceptible). This is probably due to peroxidase activity from the parasite itself (Antonova and ter Borg, 1996) which masks the host peroxidase activity.
The dispersed callose depositions observed, located near the infection point but in cells that are not in direct contact with the parasite, do not seem to be implicated in cell wall reinforcement, contrary to what has been found with O. crenata (Pérez-de-Luque et al., 2006a) and other pathogens (Brown et al., 1998). However, another possible role for callose depositions as a reservoir of ß-glucan elicitors as suggested by Esquerré-Turgayé et al. (2000) cannot be ruled out.
In addition to suberization of cell walls, sunflower produces and excretes phytoalexins (phenolic compounds) as a defensive response against O. cumana attack. Previous work showed induced synthesis and excretion of coumarins in response to O. cumana infection in sunflower (Serghini et al., 2001), and accumulation of phenolic compounds in response to O. aegyptiaca attack in Vicia athropurpurea (Goldwasser et al., 1999) and O. crenata attack in pea (Pisum spp.) (Pérez-de-Luque et al., 2005a). The results obtained here by fluorescence and confocal microscopy studies strongly indicate that resistant sunflower is poisoning the parasite by locally releasing toxic compounds very probably of a phenolic nature. The excretion of phytoalexins precedes O. cumana penetration into the host, taking place during the attachment of the parasite to the sunflower roots (Fig. 5A), and this excretion lasts until the parasite intrusive cells are stopped and the seedling dies. During the penetration step, phenolic compounds are excreted into the apoplast by resistant host cells neighbouring parasite intrusive cells. As a consequence, at the same time that the parasite finds its penetration attempt halted, a toxic environment is created around the infection point.
To conclude, according to the present results, it can be asserted that resistance to O. cumana race F in HE-39999 plants is a consequence of the co-ordinated induction of several defence mechanisms. These defence responses have a highly localized nature, being restricted to the cells in contact with the pathogen and to the interface between both organisms. These mechanisms constitute on one hand a physical barrier to prevent parasite intrusion, i.e. protein cross-linking and suberization of cell walls, and on the other hand a chemical response, by producing and secreting toxic compounds of a phenolic nature that will finish poisoning the parasite.
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Acknowledgements |
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The authors thank Advanta Ibérica S.A. (Sevilla, Spain) for kindly providing sunflower genotypes and Orobanche cumana seeds. We thank M Dolores Lozano and Rafael Susín for their help in the realization of this work, and the microscopy service of the University of Córdoba-SCAI where observations were made. AP-d-L is a researcher at the IFAPA funded by the programme ‘Juan de la Cierva’ of the Spanish Ministry of Education and Science, and AMM is the recipient of a postdoctoral contract from ‘Junta de Andalucía’. This research was partly supported by a fellowship of Advanta Ibérica S.A. in collaboration with FUNDECOR and the University of Córdoba through the PRAEM programme.
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Abbreviations |
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AGS, alcian green–safranin; CLM, confocal laser microscopy; DAB, 3,3-diaminobenzidine; DPI, days post-inoculation; QTL, quantitative trait locus; PR, pathogenesis related; ROS, reactive oxygen species.
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References |
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