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1. Plants possess numerous mechanisms to control infections by deleterious organisms. Unspecific resistance mechanisms may, however, also exert ecological costs when they have a negative effect on beneficial plant–microbe interactions. Such negative effects may even cross the border between a plant’s aerial parts and its roots and then affect very central functions such as nutrient uptake and root resistance to micro-organisms. Whereas an impaired nodulation indeed appears common after resistance expression in the leaves, contradictory results have been published for the case of arbuscular mycorrhizal (AM) fungi.
2. We analysed the effect of induction of resistance mechanisms in foliar tissues on AM colonization in soybean plants, using acibenzolar-S-methyl (ASM) as chemical elicitor. By determining different physiological and biochemical parameters, we assessed whether the effects are related to the activation of the plant defence mechanisms or rather to the re-allocation of primary metabolites.
3. Colonization with AM fungi transiently decreased after pathogen resistance mechanisms were elicited in the aerial parts of the plant. The induction with ASM led to a significant, yet moderate, defence response in the roots, which was modulated in mycorrhizal plants. No allocation or fitness costs associated with the induction of resistance were detected in this study.
4.Synthesis. Our study confirms a transient negative impact of the elicitation of foliar defences on root–AM interactions. The results show that induced resistance to foliar pathogens can (i) move from the above-ground to the below-ground compartment and (ii) affect mutualistic micro-organisms as well as plant pathogens. We also conclude that (iii) the negative effect is likely linked to changes in the defence status of the plant rather than to changes in resource allocation patterns and (iv) the AM association can modulate the activation of the plant defence mechanisms and overcome such effects.
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Higher plants largely rely on mutualisms with a diverse set of organisms such as pollinators, natural enemies of herbivores, and micro-organisms such as leaf endopyhtic fungi and soil-borne rhizobia and fungi. Fungi forming arbuscular mycorrhizas (AM) are believed to associate with the roots of over 80% of all higher plants in nature and have significant influences on the nutrition of the plant (Parniske 2008; Smith & Read 2008) as well as on its defensive status (Pozo & Azcón-Aguilar 2007; Kempel et al. 2010). As a consequence, AM have a significant impact on plant interactions with other organisms and, ultimately, on ecosystem functioning (van der Heijden, Bardgett & van Straalen 2008).
Plant must, however, also cope with pathogenic micro-organisms, and they do so by activating various defence mechanisms (Pieterse et al. 2009). Because plants interact simultaneously with multiple other organisms, their responses to specific events may have a significant impact on other interactions, and these effects can even cross the border between the aerial compartments and the roots (van Dam 2009). In fact, the relevance of induced plant defence responses for above-ground and below-ground communities and their interactions is now being recognized (Bezemer & van Dam 2005; Erb et al. 2009; van Dam and Heil 2011). Because of the intimate association of plants with mutualistic micro-organisms, several cross-reactions may occur between plant resistance to pathogens and beneficial associations with, for example, nodulating rhizobia, free-living bacteria in the rhizosphere and mycorrhizal fungi (Pozo & Azcón-Aguilar 2007; Soto et al. 2009; Yang et al. 2011). In this study, we investigate whether the induction of resistance mechanisms to pathogens in foliar tissues can cross the above-ground–below-ground border and have an impact on the symbiosis of roots with AM fungi.
Systemic acquired resistance (SAR) is an induced resistance in plants that is expressed upon local pathogen infection and confers resistance to a broad spectrum of attackers (Ryals et al. 1996; Sticher, Mauch-Mani & Métraux 1997). In most plant species, this response requires the synthesis of the endogenous signalling molecule salicylic acid (SA) in the infected tissue and its accumulation in the systemic tissue (Heil & Ton 2008; Xu et al. 2009) and it is characterized by the synthesis of pathogenesis-related (PR) proteins (Durrant & Dong 2004). Fourteen families of PR proteins have been described so far, and chitinases and β-1,3-glucanases are among them (van Loon & van Strien 1999). Chitinases and β-1,3-glucanases catalyse the hydrolysis of chitin and β-d-glucans, respectively, so both possess direct antimicrobial activity by degrading common microbial cell wall components (Dumas-Gaudot et al. 1996; van Loon 1997). Thus, PR proteins, together with other elements of SAR, contribute to a broad-spectrum resistance against diverse pathogenic bacteria, fungi and viruses. Because of this lack of specificity, they may also have a negative impact on microbial mutualists of the plant, then leading to significant ‘ecological costs’ of SAR (Heil 2002).
How does a resistance against leaf pathogens affect below-ground symbioses with mutualistic micro-organisms? Unfortunately, the physiological mechanisms of the above-ground–below-ground interactions are unclear in most cases, although a re-allocation of primary metabolites or a systemic resistance expression appears to be the most common reason (van Dam & Heil 2011). Besides the induction of active resistance, elicitation of SAR can cause a reduction in plant productivity and fitness, which results from a shift in the allocation pathways from growth to defence (Heil 2001a; Walters & Heil 2007). A common phenomenon associated to this ‘shift from housekeeping to resistance expression’ is a reduction in photosynthesis (Logemann et al. 1995; Scheideler et al. 2002; Swarbrick, Schulze-Lefert & Scholes 2006; Schwachtje & Baldwin 2008; Heil & Walters 2009). Thus, negative effects of foliar resistance expression on below-ground mutualisms could result from re-allocation of assimilates, as described from those cases in which plant tolerance responses increased root susceptibility to root herbivores (Kaplan et al. 2008). The goal of our present study was, therefore, to elucidate the physiological mechanisms that underlie any putative phenotypic effects of elicitation of resistance in above-ground tissues on the interaction of roots with AM fungi.
Considering the contradictory studies on the effect of SAR on mycorrhizas cited above, and the lack of knowledge on the underlying mechanisms, detailed research is required to understand the impact of plant resistance regulation on AM. Using a well-characterized chemical elicitor and a mycotrophic model plant such as soybean allows a detailed investigation of such potential effects and the underlying biochemical and molecular mechanisms. We tested the hypothesis that SAR-expressing soybean plants are colonized by the AM fungus Glomus mosseae to a lower extent than untreated plants. In order to assess if such effects are related directly to the defence mechanisms or to the allocation costs that are associated with the expression of the defence, we analysed defence responses and growth-associated parameters. Protein content, the activity of chitinases and β-1,3-glucanases and the expression of the PR-1a gene were chosen as disease resistance markers, whereas plant weight, sugar content, photosynthetic activity and seed production were chosen as markers for allocation and fitness costs.
Materials and methods
Plant and fungal material, growth conditions and chemical induction of resistance
Soybean seeds (Glycine max (L.) Merr. cv. Williams 82) were surface sterilized with a commercial bleach solution (10%, v/v) and germinated under sterile conditions on wet filter paper at 28 °C for 3 days. Plants were grown in 500 mL pots containing a sterile mixture of quartz sand and soil (1:1, v/v).
Arbuscular mycorrhizal fungal inoculum consisted of propagules from G. mosseae (Nicol. and Gerd.) Gerd. and Trappe (BEG 12) including spores and chopped Allium porrum L. roots colonized by the fungus in a sand:sepiolite (1:1, v/v) substrate. Mycorrhizal inoculation was carried out by mixing with the growing substrate 7% of the inoculum. In control plants 7% sand:sepiolite (1:1, v/v) was mixed with the growing substrate. At potting, all plants received a filtrate (< 20 μm) of the AM fungal inoculum in order to provide the microbial populations accompanying the mycorrhizal inocula but free from AM propagules.
Soybean plants were cultivated in a controlled environment room (25/18 °C day/night temperature, 60% relative humidity, 16 h photoperiod, with a photosynthetic photon flux of 400 μmol photons m−2 s−1). Plants were watered three times a week, the first 3 weeks only with water, and then twice a week with water and once a week with 50% Long Ashton nutrient solution (Hewitt 1966) at one-quarter phosphorus strength.
Six weeks after planting half of the plants were sprayed with an aqueous 400 mg L−1 Bion® solution at 10 mL per plant in order to induce resistance. Bion® is the name under which ASM is marketed by Syngenta (Basel, Switzerland). The rest of the plants were sprayed with the same amount of distilled water. Plant roots had no direct contact with ASM as the application was done by spraying only the shoots. The experimental design consisted, thus, of four different treatments: non-induced non-mycorrhizal plants (Nm−), induced non-mycorrhizal plants (Nm+), non-induced plants inoculated with G. mosseae (Gm−), and induced plants inoculated with G. mosseae (Gm+).
Plant harvest and assessment of mycorrhizal colonization
Plants were harvested 1 and 2 weeks after the application of ASM (7 and 8 weeks after planting). The root system was carefully washed in running tap water and then the fresh weight of shoots and roots was annotated. About one-third of the root system from each plant was kept for determination of mycorrhizal colonization, and the rest was immediately frozen in liquid nitrogen and stored at −80 °C until protein and RNA extraction. Leaf samples were also frozen and stored at −80 °C.
For the estimation of mycorrhizal colonization, samples from the root system were cleared and stained (Phillips & Hayman 1970), and the percentage of total root length colonized by the AM fungi was determined by using the gridline intersect method (Giovannetti & Mosse 1980).
The remaining plants were harvested at the end of their growth period (17 weeks after planting) to record the seed dry weight per plant and the number of seeds per plant.
Sugar extraction and quantification
Sugars were extracted following a modified procedure of the method described by Bligh & Dyer (1959). One gram of fresh leaf material from the plants harvested 1 and 2 weeks after the application of ASM was homogenized in a mortar filled with 7.5 mL methanol for 2 min. Then 7.5 mL chloroform was added and the mixture was homogenized for 1 min. Finally, 3.75 mL of water with 0.88% NaCl were added, the solution was centrifuged at 4500 g for 10 min at 0 °C, and the upper layer containing the sugars was kept for further analysis.
Quantification of sugars was carried out following the anthrone method (Morse 1947). One hundred microlitre sugar solution, 900 μL water and 3 mL anthrone solution (200 mg anthrone + 100 mL 72% H2SO4) were mixed and boiled for 10 min at 100 °C. Samples were then put on ice to stop the staining reaction. The absorbance was measured at 620 nm in a spectrophotometer. The amount of sugars released was determined by comparison with a glucose standard curve with concentrations ranging from 0 to 400 μg mL−1. One millilitre of each glucose concentration was mixed with 3 mL anthrone solution, boiled and the absorbance measured as described above.
Photosynthetic assimilation was measured with a portable photosynthesis system (Li-Cor 6400, Lincoln, NE, USA) 1 week after ASM treatment. The CO2 concentration was set at 400 p.p.m. and the photosynthetically active radiation at 1000 μmol m−2 s−1. All measurements were performed in the morning, between 9:00 and 12:00 am.
Protein extraction and quantification
Frozen plant material (root and leaf samples) from the plants harvested 1 week after the application of ASM was ground at 4 °C in an ice-chilled mortar with liquid nitrogen and the resulting powder was suspended in 100 mm MacIlvaine extracting buffer (citric acid/Na2HPO4), pH 6.8 (1:1, w/v). Crude homogenates were centrifuged at 15 000 g for 30 min at 4 °C and the supernatant fractions were kept frozen at −20 °C. Protein contents were determined by the method of Bradford (1976) using BSA as standard.
Chitinase and β-1,3-glucanase activity assays
Chitinase activity was determined with a fluorimetric assay (Ren, Wee & Chang 2000). Five microlitre methylumbelliferyl β-d-N, N’,N’-triacetylchitotrioside hydrate (0.5 mg mL−1; Sigma, Alcobendas, Madrid, Spain) were added to 95 μL plant extract (root and leaf samples) in a black 96-well microplate. Samples were incubated at 40 °C for 30 min in a shaker. The fluorescence was measured using a Perkin Elmer luminescence spectrometer LS50 (excitation 365 nm, emission 450 nm). All values were reported as fluorescence units.
The activity of β-1,3-glucanase was performed in 96-well microplates using an adaptation of the method described by Somogyi (1952). Laminaria digitata laminarin (Sigma) was used as substrate. The total volume of 180 μL reaction preparation contained 50 μL plant extract (root and leaf samples), 10 μL laminarin (20 mg mL−1 in 50 mm Na-acetate buffer, pH 5.0), 60 μL copper reactive and 60 μL arsenic reactive. The absorbance was measured at 650 nm in a spectrophotometer. The amount of sugars released was determined by comparison with a glucose standard curve with concentrations ranging from 0 to 200 μg mL−1.
RNA Extraction and gene expression analysis by quantitative real-time PCR
Total RNA was isolated from root and leaf samples using Tri Reagent® (Ambion, Foster City, CA, USA) and treated with DNase RQ1 (Promega, Madison, WI, USA). cDNA synthesis was performed using iScript™ cDNA Synthesis Kit (Bio-Rad Laboratories, Hercules, CA, USA) starting with 1 μg of total DNAse-treated RNA.
Gene expression levels were determined by quantitative real-time PCR, using the primers detailed in Table 1. The sets of primers for Gm18S and GintEF have been previously used as constitutive expression controls for soybean and Glomus intraradices, respectively (Porcel et al. 2006; Benabdellah et al. 2009). The soybean PR-1a gene was selected as marker for SA-regulated defence responses, and the corresponding set of primers was designed using the primer 3 software (Rozen & Skaletsky 2000). Prior to real-time PCR analysis, the specificity of the selected primers was checked by conventional RT-PCR. The efficiency of the primer sets was evaluated by performing real-time PCR on several dilutions of genomic DNA. All quantitative real-time PCR reactions were performed using iQ SYBR Green Supermix (Bio-Rad Laboratories) on an iQ5 Multicolor Real-Time PCR Detection System (Bio-Rad Laboratories). The amplification protocol included an initial denaturation at 95 °C for 3 min followed by 35 cycles of 95 °C for 30 s, 58 °C for 30 s, and 72 °C for 30 s. The specificity of the PCR amplification procedure was checked with a heat dissociation protocol (from 70 to 100 °C) after the final cycle of the PCR. The results obtained for the different treatments were standardized to the soybean 18S rRNA levels, which were amplified with the primers Gm18S shown in Table 1.
Table 1. Primers used for the quantitative real-time PCR
Real-time PCR experiments were carried out at least three times, with the threshold cycle (CT) determined in triplicate. The relative levels of transcription were calculated by using the 2-DDCT method (Livak & Schmittgen 2001). Negative controls without cDNA were included in all PCR reactions.
Replication and statistical analysis
Eighteen replicated plants were used per treatment, four of them were harvested at each time point (7 and 14 days after treatment) and used for the physiological and biochemical analyses and the rest (10 replicated plants) were used for the quantification of seed production. statistica 6.1 software (StatSoft Inc., Tulsa, OK, USA) was used for the statistical analysis. A Student’s t-test was carried out to detect significant effects of induction of resistance on AM colonization. Mann–Whitney U tests were conducted to detect significant effects of induction of resistance on fresh weight, sugar content, photosynthetic activity, protein content, chitinase and β-1,3-glucanase activity, and seed production parameters. Mean ± SE were calculated. Significance levels were set at 5%.
Soybean plants inoculated with G. mosseae harvested 1 week after the application of ASM (Gm+) showed significantly lower AM colonization than Gm plants sprayed only with water (Gm−) (Fig. 1a; t =2.63; d.f. = 6; P =0.038; t-test). A second harvest of Gm+ plants 2 weeks after ASM treatment revealed that the differences in AM colonization between Gm+ and Gm− plants had levelled off (t =0.35; d.f. = 6; P =0.735; t-test). Thus, the negative effect of the induction of systemic resistance on the AM colonization of soybean plants appeared transitory. The absence of AM colonization was confirmed in non-inoculated plants (Nm) (data not shown).
Quantifying the expression level of a fungal ‘housekeeping gene’ is known to be an accurate and sensitive method to estimate the amount of active, living AM fungus within root tissues (Isayenkov, Fester & Hause 2004). The quantitative real-time PCR of transcript levels of GintEF, constitutively expressed in AM fungi (Benabdellah et al. 2009), confirmed a significant reduction of the AM fungus in Gm+ roots 1 week after treatment (0.4-fold compared to untreated plants; Fig. 1b). The fungal presence increased with time in both treated and untreated plants, and 2 weeks after the chemical treatment the inhibition was completely overridden. The results confirm that the negative effect of ASM application on AM colonization was transitory.
Resistance induction has been associated to a shift in resources allocation from growth to defence, and this shift may potentially limit the resources available for the fungal symbiont, thus limiting AM colonization. To assess if this is the case, we compared plant biomass, sugar content and photosynthetic activity in induced and non-induced plants. Neither the fresh weight, nor the sugar content, nor the photosynthetic activity changed significantly in either Nm or Gm plants after the induction of resistance (P > 0.05; Mann–Whitney U test; Table 2).
Table 2. Photosynthetic activity, sugar content and fresh weight of soybean plants 1 and 2 weeks after the application of acibenzolar-S-methyl (ASM)
One week after induction
Two weeks after induction
Photosynthesis (μmol CO2 m−2 s−1)
Sugar content (mg g−1 fresh wt. leaves)
Fresh wt. shoots (g)
Fresh wt. roots (g)
Sugar content (mg g−1 fresh wt.leaves)
Fresh wt. shoots (g)
Fresh wt. roots (g)
Fresh wt., fresh weight; Nm−, non-induced non-mycorrhizal plants; Nm+, induced non-mycorrhizal plants; Gm−, non-induced plants inoculated with Glomus mosseae; Gm+, induced plants inoculated with G. mosseae. Mean values of Nm− and Nm+ and Gm− and Gm+ in each column were compared using paired Mann–Whitney U tests. n = 4; ns P >0.05.
Nm+ plants produced fewer seeds than Nm− ones (Fig. 2a,b), although the difference was insignificant both for seed dry weight per plant (P =0.155; Mann–Whitney U test) and numbers of seeds per plant (P =0.073; Mann–Whitney U test). No hints towards changes in either the seed dry weight per plant (P =0.934; Mann–Whitney U test) or the number of seeds per plant (P =0.866; Mann–Whitney U test) were observed between Gm+ and Gm− plants.
We also analysed the impact of ASM treatment on the protein content and defence-related enzyme activities. The chemical induction of resistance led to a significant increase in protein content both in leaves and roots (P <0.05; Mann–Whitney U tests) of Gm+ plants harvested 1 week after the application of ASM as compared to Gm− plants (Fig. 3a,b). No significant differences in protein content were observed between Nm+ and Nm− plants (P > 0.05; Mann–Whitney U test).
Chitinase activity (Fig. 4a,b) in leaves of Gm+ plants (P <0.01; Mann–Whitney U test) 1 week after the induction of resistance was significantly higher than in Gm− plants. No significant differences were observed in root chitinase activity between Gm+ and Gm− plants or in either roots or leaves of Nm+ and Nm− plants (P > 0.05; Mann–Whitney U test).
β-1,3-glucanase activity (Fig. 5a,b) was significantly higher in leaves of induced Nm+ and Gm+ plants compared to non-induced plants, as well as in roots of Nm+ plants when compared with Nm− plants (P <0.05; Mann–Whitney U tests). No significant changes in β-1,3-glucanase activity were observed in roots of Gm plants after the induction of resistance (P > 0.05; Mann–Whitney U test).
Acibenzolar-S-methyl is a functional analogue of SA, and its application induces resistance in plants through the activation of the SA signalling pathway. In order to monitor this pathway we analysed the expression levels of the soybean PR-1a gene, coding for an acidic isoform of PR-1, a common marker for SA-regulated responses. ASM application resulted in an increase in PR-1a transcript levels in shoots in Nm and Gm plants, confirming the activation of the SA signalling pathway (Fig. 6a). In agreement with the reported systemic nature of the elicitor, the expression levels of the gene were also significantly higher in the roots of Nm+ plants. Although the basal level of PR1a expression was higher in mycorrhizal plants than in non-mycorrhizal controls, no increase in PR-1a expression was observed in roots of Gm plants upon induction by ASM treatment (Fig. 6b). Elevated levels of PR1a expression in roots were still found 2 weeks after induction in Nm+ plants, but again no difference was detected between Gm+ and Gm− plants (Fig. 6c).
What is the biochemical or genetic mechanism that underlies this phenomenon? Negative effects of SAR induction on the interaction of plants with beneficial microbes can principally result from two different phenomena: (i) the ‘metabolic shift’ from primary to secondary metabolism that usually occurs during resistance elicitation might lower photosynthesis and, thus, the allocation of assimilates to the root system (as suggested by Russin et al. 1990), or (ii) defensive traits such as PR proteins might exert direct negative effects on the beneficial microbes (as suggested by Heil 2001b). In fact, the AM association, although a mutualistic one, is still a form of fungal invasion, and the plant activates its defence mechanisms (Riedel, Groten & Baldwin 2008). Remarkably, local defence responses such as increases of chitinase and β-1,3-glucanase activities are activated during early steps of compatible AM interactions (Ruiz-Lozano et al. 1999; Pozo & Azcón-Aguilar 2007), but these enzymatic activities are repressed at a later stage of mycorrhiza formation (Dumas-Gaudot et al. 1996; Gianinazzi-Pearson et al. 1996; Kapulnik et al. 1996). However, induction of specific hydrolytic enzymes (chitinase, chitosanase and β-1,3-glucanase isoforms) as a response to the AM symbiosis that differ from those induced as the result of general defence mechanisms have been reported in different plants (Dumas-Gaudot et al. 1996; Pozo et al. 1996, 1998, 1999). The induction of these hydrolytic enzymes and other defence-related compounds in AM symbiosis seems to be involved in the plant control over the fungal symbiont, but it may affect other organisms (Pozo et al. 2002; López-Ráez et al. 2010). Indeed, the AM symbiosis is usually associated with an enhanced disease resistance or mycorrhiza-induced resistance (Pozo & Azcón-Aguilar 2007). Thus, besides a shift in assimilate allocation patterns, a shift in the delicate balance between host resistance and AM infection (Bennett, Bever & Bowers 2009) provides an alternative explanation of the pattern reported here.
In order to distinguish among these two alternatives we quantified photosynthetic rates and resistance expression in our plants. No changes in photosynthetic activity, sugar content or plant growth were recorded in either mycorrhizal or non-mycorrhizal plants after the induction of resistance. We conclude, therefore, that the negative effects of resistance induction on AM colonization were linked to the biochemical changes associated with the defence response rather than a shift in the allocation pathways from growth to defence. The application of ASM elicited a significant, yet moderate, increase in protein content and PR protein activity. To further analyse the activation of defence responses by ASM, we monitored expression levels of the PR-1a gene, commonly used as marker for SA-dependent defence responses (Uknes et al. 1993). Gene expression levels confirmed the activation of this signalling pathway not only in shoots, but also in roots of ASM treated non-mycorrhizal plants. SA-dependent responses are effective against biotrophic pathogens (Glazebrook 2005), but they may also impact mycorrhizal fungi because of their biotroph condition (Pozo & Azcón-Aguilar 2007). It is, therefore, likely that the activation of the SA signalling pathway upon ASM treatment results in an inhibition of AM root colonization.
Interestingly, ASM treatment led to a significant change in leaf chitinase activity in Gm plants but not in Nm plants. This pattern was caused by both, a lowered chitinase activity in Gm controls and a higher chitinase activity in Gm-induced plants, compared to the respective groups of Nm plants (Fig. 4a). This pattern is consistent with a successful suppression of chitinase activity by G. mosseae, which is transiently lost after ASM treatment. Our results are, therefore, consistent with the hypothesis that a shift in the balance between host resistance and control of AM infection explains why resistance induction transiently inhibits mycorrhization.
It is assumed that the induction of disease resistance incurs allocation and/or fitness costs, but only few studies have demonstrated or quantified them (Heil et al. 2000; Cipollini 2002; Heil 2002). In other cases, the results depended on growing conditions (Dietrich, Ploss & Heil 2005). In our study, plant growth was not affected by the induction of resistance. Similar results were reported by Iriti & Faoro (2003), and Zhu et al. (2003) and Faessel et al. (2008) recorded only transitory negative effects on plant growth. As for the seed production, the induction of resistance was associated with a lower number of seeds per plant and a lower seed dry weight per plant in non-mycorrhizal plants, although the differences were not significant. This trend was not observed in plants inoculated with G. mosseae.
In conclusion, our results support the hypothesis that the elicitation of foliar plant defence responses may negatively affect beneficial plant–microbe interactions below-ground, as a decrease in AM colonization of soybean roots was detected after the application of the chemical elicitor ASM to the leaves. The negative effects of the induction of resistance on AM seemed to be directly linked to the biochemical changes associated with the defence response and not to a possible shift in the allocation pathways from growth to defence. Because AM colonization plays an important role in plant nutrition and resistance, these effects likely will affect the future capacity of the plant to survive in the presence of competitors and enemies. Thus, it appears adaptive from the perspective of the plant to minimize these effects as far as possible and indeed, the decrease in AM colonization was only transitory. Because the ability of AM fungi to modulate the plant defence responses likely explains the transitory nature of the effect, we speculate that traits of both the host and its symbiont may be involved in overcoming the negative effects of above-ground resistance expression on the below-ground mutualism. Future studies aiming at an identification of the mechanisms of this co-operative effort would involve studying the interaction under different nutrient conditions and densities of AM colonization (Vannette & Hunter 2011) and monitoring physiological and transcriptomic changes in both plant roots and AM fungi during the various phases of the interaction.
We thank Juan M. García for his excellent technical assistance. M. de R. was supported by a postdoctoral fellowship from the Alexander von Humboldt Foundation, and I.F. by a PhD grant from the Spanish Ministry of Science and Innovation MINCIN. This work was funded by grant AGL2006-08029 from the MINCIN.