The immediate activation of defense responses in Arabidopsis roots is not sufficient to prevent Phytophthora parasitica infection


  • Agnès Attard,

    1. Unité Mixte de Recherches Interactions Biotiques et Santé Végétale, INRA1301-CNRS6243-UNS, 400 route des Chappes, F-06903 Sophia Antipolis, France
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  • Mathieu Gourgues,

    1. Unité Mixte de Recherches Interactions Biotiques et Santé Végétale, INRA1301-CNRS6243-UNS, 400 route des Chappes, F-06903 Sophia Antipolis, France
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  • Nicolas Callemeyn-Torre,

    1. Unité Mixte de Recherches Interactions Biotiques et Santé Végétale, INRA1301-CNRS6243-UNS, 400 route des Chappes, F-06903 Sophia Antipolis, France
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  • Harald Keller

    1. Unité Mixte de Recherches Interactions Biotiques et Santé Végétale, INRA1301-CNRS6243-UNS, 400 route des Chappes, F-06903 Sophia Antipolis, France
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Author for correspondence:
Agnès Attard
Tel: +33 492386594


  • The outcome of plant–microbe interactions is determined by a fine-tuned molecular interplay between the two partners. Little is currently known about the molecular dialogue between plant roots and filamentous pathogens. We describe here a new pathosystem for the analysis of molecular mechanisms occurring during the establishment of a compatible interaction between Arabidopsis thaliana roots and a root-infecting oomycete.
  • We performed cytological and genetic analyses of root infection during the compatible interaction between A. thaliana and Phytophthora parasitica.
  • Phytophthora parasitica uses appressoria to penetrate A. thaliana roots. Initial biotrophic growth is accompanied by the formation of haustoria, and is followed by a necrotrophic lifestyle. Arabidopsis thaliana mutants with impaired salicylic acid (SA), jasmonic acid (JA) or ethylene (ET) signaling pathways are more susceptible than the wild-type to infection. The salicylate- and jasmonate-dependent signaling pathways are concertedly activated when P. parasitica penetrates the roots, but are downregulated during invasive growth, when ethylene-mediated signaling predominates.
  • Thus, defense responses in A. thaliana roots are triggered immediately on contact with P. parasitica. Our findings suggest that the pattern of early defense mechanism activation differs between roots and leaves.


When a pathogen comes into contact with a plant, this contact may result in the onset of resistance or the establishment of disease. The first level of plant resistance is activated through the recognition of pathogen-associated molecular patterns (PAMPs), small molecular motifs conserved within a class of microbes and exposed on infection. The resulting PAMP-triggered immunity (PTI) involves the production of reactive oxygen species (ROS), pathogenesis-related (PR) protein synthesis and reinforcement of the cell wall with callose, lignin and other phenolic polymers at infection sites. PTI protects plants effectively against most pathogen invasions (Jones & Takemoto, 2004; Nürnberger et al., 2004). A second, more specialized level of plant resistance is triggered by the direct or indirect recognition of effectors from the pathogen by specific plant resistance gene products. This effector-triggered immunity (ETI) frequently leads to rapid, localized programmed host cell death at sites of infection, inhibiting pathogen development (Wojtaszek, 1997; Heath, 2000; Jones & Dangl, 2006). This onset of resistance results in so-called incompatible interactions.

The activation of plant defense responses against pathogens is controlled by signaling pathways involving hormones, such as salicylic acid (SA), ethylene (ET) and jasmonic acid (JA) (Dong, 1998; Thomma et al., 2001). SA is involved in both PTI and ETI, and in a systemic immune response called systemic acquired resistance. ET and JA frequently act in concert to regulate pathogen resistance and wound responses (Ryals et al., 1996; Turner et al., 2002; Wang et al., 2002; Robert-Seilaniantz et al., 2007). However, the SA- and JA/ET-mediated signaling cascades have been shown to have antagonistic effects, and the balance between these pathways depends on the lifestyle of the invading pathogen. A plant generally activates the SA pathway and represses JA/ET signaling when dealing with biotrophic pathogens, whereas JA/ET signaling is activated and SA signaling is repressed when repelling necrotrophs (Glazebrook, 2005).

Unrecognized pathogens multiply and spread within the host plant tissues, thus causing disease. In such conditions, the plant is considered to be susceptible and the interaction is described as compatible. Defense pathways are also activated during compatible interactions. Genome-wide analyses of the Arabidopsis thaliana transcriptome after infection with Pseudomonas syringae and Hyaloperonospora arabidopsidis have shown that the defense responses activated are similar in compatible and incompatible interactions, but that these responses are weaker and slower in compatible than in incompatible interactions (Tao et al., 2003; Desender et al., 2007; Rinaldi et al., 2007; Silvar et al., 2008; Huibers et al., 2009). More than 97% of the genes displaying a change in expression during incompatible interactions are also activated or repressed when the interaction is compatible (Huibers et al., 2009).

Most research projects aiming to decipher the molecular plant–pathogen dialogue have focused on interactions involving the aerial parts of plants. The molecular mechanisms underlying aerial plant defenses are thus well documented, whereas little is known about the mechanisms underlying root responses to soil-borne pathogens. However, there is growing evidence to suggest that defense mechanisms are different in roots and leaves (Hermanns et al., 2003; Schlink, 2009). For example, Schlink (2009) found that beech root responses to Phytophthora citrocola differed from leaf responses, and showed that most of the genes activated in roots had no known function or no matches with database sequences for genes activated in aerial parts of plants. Moreover, and unlike in leaves, expression of the A. thaliana resistance gene RPP1 in roots was found to be insufficient to trigger ETI to the H. arabidopsidis isolate Noco2 (Hermanns et al., 2003). An understanding of the molecular events governing root–pathogen interactions is thus of considerable interest to plant pathologists.

The available data concerning interactions between filamentous pathogens and plant roots are limited, because only a few model systems are available. Assays developed for the analysis of plant responses to soil-borne pathogens have often been performed on leaves, with extrapolation of the results to root interactions. For example, gene expression analyses to determine the molecular events underlying the susceptibility of A. thaliana to the soil-borne pathogens Verticillium dahlia and Fusarium oxysporum were performed on leaf tissues (Veronese et al., 2003; Johansson et al., 2006; Berrocal-Lobo & Molina, 2008).

Most root diseases in crops are caused by soil-borne oomycetes, such as biotrophic downy mildews, hundreds of necrotrophic Pythium species and > 85 Phytophthora species. Oomycetes infect a wide spectrum of plants (Erwin & Ribeiro, 1996). Phytophthora species have a major ecological and economic impact, with annual losses caused by pathogens from this genus estimated at 5 billion dollars (Stokstad, 2006). All Phytophthora species are considered to be hemibiotrophic pathogens, but they differ in their host range. Some species, such as Phytophthora infestans and Phytophthora sojae, are specialized pathogens infecting only a few hosts, whereas others, such as Phytophthora parasitica, are able to infect > 60 plant families (Erwin & Ribeiro, 1996). The infection cycle of Phytophthora spp. is initiated by the attraction of swimming zoospores to plant roots. In most cases, penetration of the root epidermis is mediated by appressorium-like structures (Tyler, 2007; Attard et al., 2008), but the direct penetration of hyphae between root cells has been reported for P. sojae (Enkerli et al., 1997). Following penetration, bulbous hyphae invade the roots intercellularly (Benhamou & Côté, 1992; Widmer et al., 1998; Le Berre et al., 2008). During the interaction between soybean and P. sojae, this stage of infection involves a short, difficult-to-observe biotrophic phase (Hanchey & Wheeler, 1971) that seems to be associated with the differentiation of specialized feeding structures, called haustoria (Enkerli et al., 1997; Perfect & Green, 2001; Tyler, 2007).

Arabidopsis thaliana is an attractive model for dissection of the cellular and molecular events underlying root responses to pathogens, but only a few pathosystems involving oomycetes are currently available for study. Phytophthora cinnamomi can invade A. thaliana roots, but causes no symptoms, A. thaliana thus being considered to be tolerant to this oomycete (Rookes et al., 2008). Phytophthora brassicae is thought to colonize A. thaliana roots, but data are available only for leaf interactions (Roetschi et al., 2001). A compatible interaction between A. thaliana roots and Phytophthora palmivora has been described (Daniel & Guest, 2006). However, under natural conditions, P. palmivora causes fruit and stem rot disease (Erwin & Ribeiro, 1996; Daniel & Guest, 2006). There is thus still a need for a model system based on an interaction between A. thaliana roots and a root-invading oomycete species, mimicking the natural events occurring in host plants and suitable for the study of oomycete–root interactions.

In this study, we aimed to fill this gap by developing a new pathosystem involving A. thaliana and the broad-host-range oomycete pathogens, P. parasitica and Phytophthora capsici, which are both susceptible to genetic transformations. We analyzed plant responses to P. parasitica and identified striking differences between this interaction and those described for aerial parts of the plant. This work provides the basis for the analysis of the molecular mechanisms underlying compatible plant–oomycete interactions in the roots.

Materials and Methods

Plant material and growth conditions

The Arabidopsis ecotypes used in this study were Columbia-0 (Col-0), C24 (C24), Landsberg erecta (Ler-0), Wassilewskija (Ws), Blanes (Bla-1), Edinburgh (Edi-0), Geneva (Ge-0), JEA, Kaunas (Kn-0), Pitztal (Pi-0), Ran, Spandau (Sp-0), Stockholm (St-0), Stobowal (Stw-0), Tenela (Te-0) and Tsushima (Tsu-0) (McKhann et al., 2004). All natural accessions and the mutant Col-0 lines, enhanced disease susceptibility 5 (eds5-1), phytoalexin deficient 4 (pad4-1) (Glazebrook et al., 1996), ethylene insensitive 2 (ein2-1) (Guzman & Ecker, 1990), ethylene receptor 1 (etr1-3) (Bleecker et al., 1988), jasmonate resistant 1 (jar1-1) (Staswick et al., 1992) and salicylic acid induction deficient 2 (sid2-1) (Nawrath & Metraux, 1999), were obtained from The European Arabidopsis Stock Centre, Nottingham, UK. The enhanced disease susceptibility 1 (eds1-1) mutant line in the Ws genetic background (Parker et al., 1996) was obtained from Dr Jane Parker through a material transfer agreement with Plant Bioscience Limited, Norwich Research Park, Norwich, UK. Seeds of Col-0 plants expressing the naphthalene hydroxylase G (nahG) gene (Lawton et al., 1996) were obtained from Dr Leslie Friedrich, Research Triangle Park, NC, USA. All Arabidopsis seeds were surface sterilized for 5 min with sodium hypochlorite (20% commercial bleach, 2.5% active ingredient) and rinsed twice in 95% aqueous ethanol. Seeds were cold-stratified for 2 d and sown on 1 × Murashige and Skoog (MS) medium (Sigma Chemical Company) supplemented with 10 g l−1 sucrose and 20 g l−1 agar. Two-wk-old plants were transferred to square Petri dishes containing a 2-cm-wide strip of solid MS agar separating the root compartment (growing in 10 ml of 0.1 × MS medium) from a compartment without medium for the aerial parts of the plants. These dishes, each containing six plants, were then placed upright for 1 month at 25°C under an 8-h photoperiod. For inoculations, zoospore suspension was added directly to the root compartment containing the medium.

Growth conditions for P. parasitica and inoculation of A. thaliana plantlets

Phytophthora parasitica Dastur isolates 310 and 149 were initially isolated from tobacco in Australia, and maintained in the Phytophthora collection at INRA, Sophia Antipolis, France. Phytophthora parasitica strain 149, which constitutively expresses cytoplasmic green fluorescent protein (GFP), has been described elsewhere (Le Berre et al., 2008). Phytophthora growth conditions and the production of zoospores have been described elsewhere (Galiana et al., 2005). For the investigation of early infection steps, we added 106 zoospores to the MS medium of Petri dishes containing 30-d-old plantlets grown as described already. For studies of late infection processes, we added 103 zoospores to the roots of these plants. After inoculation, plantlets were incubated at 25°C, as described already. Statistical analyses of disease severity were based on Scheirer–Ray–Hare nonparametric two-way analysis of variance (ANOVA) for ranked data (H-test; Sokal & Rohlf, 1995). The statistical analysis was carried out on inoculated plants (= 11–15) and all experiments were performed at least twice.

Tissue-staining protocols

To visualize oomycete colonization of host tissues, samples were first fixed with glutaraldehyde–formaldehyde (1% and 4%, respectively, in 50 mM sodium phosphate buffer at pH 7.2). The oomycete was then visualized by staining for 3 min with propidium iodide (PI, 1 μg ml−1 in water, Invitrogen). Alternatively, fixed samples were cleared with ethanol, and stained with Chlorazol black E (Sigma-Aldrich; 0.03%, in water–glycerol–lactic acid, 1 : 1 : 1, v/v/v) for 24 h, according to Adler et al. (1948). Calcofluor white was used for the localization of cell wall appositions within fixed, cleared tissues, as described previously (Ruzin, 1999; Ton et al., 2005). The viability of infected root cells was assessed by staining fresh root segments for 3 min with PI and then for 3 min with fluorescein diacetate (FDA; 2 μg ml−1 in water; Toda et al., 1999).


Infected roots were observed with a Zeiss Axioplan microscope. Samples were viewed with light transmission, differential interference contrast (DIC) and epifluorescence optics. For epifluorescence microscopy, we used the following filters with the HBO-100 mercury vapor lamp light source: a GFP/FITC filter for PI/FDA and GFP observation, and a rhodamine filter for PI observation. Images were acquired with a Zeiss AxioCam camera and analyzed with Zeiss Axiovision digital image-processing software, version 4.4. Confocal laser scanning microscopy images were obtained with a Zeiss LSM 510 META confocal microscope (Carl Zeiss GmbH, Jena, Germany). For GFP visualization, an argon laser was used for excitation at 488 nm, whereas an HeNe laser was used for excitation at 543 nm, with a 530-nm long-pass filter and DIC-transmitted light, for the visualization of PI.

Quantitative reverse transcription-polymerase chain reaction (qRT-PCR) analyses

Total RNA was extracted from freeze-dried infected roots, as described previously (Laroche-Raynal et al., 1984). RNA was treated with DNAse I (Ambion, Austin, TX, USA), and 1 μg was reverse transcribed to generate first-strand cDNA, using the I Script cDNA synthesis kit according to the manufacturer’s instructions (Bio-Rad, Hercules, CA, USA). qRT-PCR experiments were performed with 5 μl of a 1 : 50 dilution of first-strand cDNA and SYBR Green, according to the manufacturer’s instructions (Eurogentec SA, Seraing, Belgium). Gene-specific oligonucleotides (PR1-At2g14610: cggagctacgcagaacaact-ctcgctaaccatgttca, 92% efficiency; PR2-At3g57260: tggtgtcagattccggtaca-tcatccctgaaccttccttg, 98% efficiency; PR3-At3g12500: caatgcaactgtcgtggaac-tgagcagtcatccagaacca, 100% efficiency; PR4-At3g04720: tagtggaccaatgcagcaac-gatcaatggccgaaacaag, 93% efficiency, PR5-At1g75040: cgtacaggctgcaactttgc-tgaattcagccagagtgacg, 95% efficiency; PDF1.2-At2g26020: accaacaatggtggaagcac-cacttgtgagctgggaagac, 100% efficiency; SID2-At1g74710: ccaattgaccagcaaatcg-caaggtcacggaagaaaactg, 97% efficiency; ICS2-At1g18870: ccaattgaccagcaaatcg-ataccagcaacgctaaccaga, 100% efficiency; FAD8-At5g05580: ctttgtcatgggtccaatcc-tgagccctcctctcaggtaa, 95% efficiency; ACO4-At1g05010: gctgatcggaaaagaagcag-cattgttggccacagttgtc, 98% efficiency; ACS2-At1g01480: tgtgtctcctggctcttcct-cctcgtaaagtcgtcggaaa, 95% efficiency) were designed with Primer3 software and their specificity was validated by analyzing dissociation curves after each run. Genes encoding a mitochondrial NADH-ubiquinone oxidoreductase subunit (At5g11770) and a mitochondrial inner membrane translocase (At5g62050) were selected as constitutive internal controls (Quentin et al., 2009). Two biological replicates of the entire experiment were performed, each as a technical triplicate. For each time point, six results were analyzed. Gene expression was quantified and normalized with respect to constitutively expressed internal controls.


Phytophthora parasitica and P. capsici undergo compatible interactions with A. thaliana

The roots of A. thaliana plantlets grown in vitro were inoculated with motile P. parasitica or P. capsici zoospores, and disease development was followed over a period of 4 wk (Supporting Information Fig. S1). A wilting disease index was established on the basis of the observed symptoms. Disease severity was ranked from 1 (healthy plants) to 7 (dead plants). In the first few days after inoculation, mycelium was observed around the roots, and the plants continued to appear healthy (Fig. S1). The first symptoms appeared at the hypocotyl (Fig. S1), and older leaves then began to be invaded (disease index 2 and 3; Fig. 1a,b). This was rapidly followed by leaf necrosis and severe damage (disease index 4; Fig. 1c). The symptoms progressed through the center of the rosette (disease index 5 and 6, Fig. 1d,e), and the whole plant died within 20 d of inoculation with P. parasitica (disease index 7; Fig. 1f,g).

Figure 1.

 Development of disease symptoms on Arabidopsis thaliana roots challenged with Phytophthora parasitica and Phytophthora capsici. Plants of Arabidopsis ecotype Col-0 inoculated with 1000 zoospores of P. parasitica strain 310 or P. capsici strain 450. (a–f) Phenotypes illustrating the disease index, which is based on disease progression over time and describes the disease severity in terms of the number and age of invaded leaves. No symptoms are visible for disease index 1. (a) Disease index 2. One or two old leaves are invaded. (b) Disease index 3. Three to four old leaves are invaded. (c) Disease index 4. Only one or two old leaves remain healthy. (d) Disease index 5. The first stage of the rosette is completely invaded and only young leaves remain healthy. (e) Disease index 6. Two to three young leaves remain healthy. (f) Disease index 7. The plant is dead. (g, h) Disease progression on three A. thaliana ecotypes inoculated with P. parasitica (g) and P. capsici (h), according to the disease index. The illustrations show the results of a representative experiment. Differences between ecotypes on inoculation with P. parasitica were statistically significant, as determined by Scheirer–Ray–Hare nonparametric two-way analysis of variance (ANOVA) for ranked data (H < 0.05). Squares, Ws-0; circles, Col-0; triangles, Ler.

The same macroscopic symptoms were observed when A. thaliana was challenged with either P. parasitica or P. capsici (Fig. S1). However, the symptoms developed more slowly in plants inoculated with P. parasitica than in those inoculated with P. capsici (Fig. 1g,h). The first macroscopic symptoms appeared 7–8 d after inoculation with P. parasitica, whereas similar symptoms occurred 4 d after inoculation with P. capsici (Fig. 1g,h). Similarly, the whole plant died within 20 d after inoculation with P. parasitica and 8 d after inoculation with P. capsici (Fig. 1g,h). Analysis of the interaction with P. parasitica using ANOVA also revealed significant differences in disease severity between the ecotypes Col-0, Ler and Ws-0 (Fig. 1g; H = 10.8; df = 2; P = 9 × 10−3). Although all ecotypes were susceptible to P. parasitica and displayed disease index 7 (dead plant) by 20 d, they differed significantly in the timing of symptom expression. By contrast, no such differences in symptom severity were observed when Ler, Col-0 and Ws-0 were inoculated with P. capsici (Fig. 1h; H = 0.13; df = 2; P = 0.93). As A. thaliana responded more slowly to P. parasitica than to P. capsici, it was easier to observe all the events of infection, including the early biotrophic stage, for this pathogen. We therefore further characterized the A. thalianaPhytophthora interaction with P. parasitica-inoculated plants.

We investigated possible differential responses to P. parasitica by evaluating the interaction of 13 additional A. thaliana ecotypes with strain 310. All ecotypes were susceptible, and the interaction eventually led to plant death. However, the disease severity differed between ecotypes, resulting in the definition of three susceptibility groups (Table 1). In the least susceptible ecotype, Ge-0, leaf invasion began 20 d after zoospore application (mean disease index of 2.5), whereas leaf invasion was observed 13 d post-inoculation (dpi) for Col-0. Infected Ge-0 plants died 30–40 dpi. By contrast, infection of the most susceptible ecotype, Kn-0, led to the death of infected plants within 15 dpi.

Table 1.   Susceptibility groups defined according to the disease severity of 16 Arabidopsis thaliana ecotypes challenged with Phytophthora parasitica (H = 169, df = 14, P = 8 × 10−29)
Susceptibility groupEcotype
Least susceptiblePi-0, Te-0, Ge-0
Moderately susceptibleCol-0, Bla-1, Edi-0, Stw-0, Tsu-0
Most susceptibleLer, Ran, JEA, Sp-0, St-0, Kn-0, Ws-0, C24

We checked that the interaction between A. thaliana and P. parasitica could also be established in semi-in vitro conditions. Plants grown in sand or hydroponic culture conditions and infected with P. parasitica developed macroscopic symptoms 15 d after the addition of zoospore suspensions to the roots. The inoculated plants died within 25 d of inoculation (data not shown).

Appressorium-mediated root penetration

Following inoculation, zoospores were attracted to the roots, accumulating massively in the elongation and differentiation zones, where they formed cysts (Fig. 2a). Cysts were observed along the entire root surface, including root hairs, with the exception of the root tips. However, the density of cysts was consistently highest in the root elongation area (Fig. 2a). Germination began 10–20 min after inoculation, and 91% of the cysts germinated within 2 h (Fig. 2a–d,h). Most of the germ tubes subsequently stopped growing and their tips swelled to form appressoria (Fig. 2e). Appressoria were formed by 65% of germinating cysts at 2 h post-inoculation (hpi), and by 95% of germinating cysts at 4 hpi (Fig. 2h). These structures were separated from the germ tube by a septum (data not shown). Successful penetration between the cells of the rhizodermis was observed for 45% of the appressorium-forming germlings at 2.5 hpi, and for 80% at 4 hpi (Fig. 2f–h). Direct penetration occurred from appressoria that differentiated on cell junctions. Attempts by the oomycete to initiate appressoria elsewhere were rapidly arrested, and hyphae continued to grow until reaching an appropriate site for penetration. However, penetration was always preceded by appressorium formation, and was accomplished by intercellular hyphae growing between cell junctions. By 6 hpi, 100% of the germinating cysts achieved penetration of the epidermis (Fig. 2h).

Figure 2.

 Penetration of Phytophthora parasitica into Arabidopsis thaliana roots. (a) Phytophthora parasitica zoospores accumulate principally at the elongation zone of roots, 1 h post-inoculation (hpi). Zoospores germinate, and germ tubes detach the spore from the surface. (b) Encysted zoospore at the root surface, 3 min post-inoculation (mpi). (c) Germinating zoospore, 20 mpi. (d) Growing germ tube, 22 mpi. (e) Appressorium differentiation, 32 mpi. (f, g) Appressorium-mediated penetration between two rhizodermis cells by P. parasitica expressing green fluorescent protein (GFP), as observed under light transmission (f) and fluorescence excitation (g) conditions. (a-g) Differential interference contrast (DIC) images. Zoospores (zsp), germinating hyphae (gHy), appressoria (app) and penetration pegs (peg) are indicated. (h) Quantitative analysis of the penetration process. At 30 mpi, 50% of the encysted zoospores had germinated. At 2 hpi, 50% of the encysted zoospores had differentiated an appressorium. At 2.5 hpi, 50% of the encysted zoospores had penetrated the root surface. By 6 hpi, all the encysted zoospores had successfully penetrated the roots. The illustration presents results from a representative experiment, in which > 100 zoospores were analyzed at each time point. Squares, triangles and circles indicate the percentage of cysts germinating, forming appressoria and leading to successful penetration of roots, respectively. Bars: (a), 50 m; (b-g), 10 m.

Invasive growth towards the leaves

Invasive growth occurred once the epidermal cell layer had been breached. During the first 8 h after inoculation, hyphae grew intercellularly, crossing the cortex towards the stele. By c. 5 hpi, 81% of the germlings had crossed three cell layers. When the hyphae reached the stele, they changed direction, growing towards the shoots along the central cylinder, without entering it (Fig. 3a,b). The root life cycle was completed by sporulation at the root surface at 5 dpi (Fig. 3c). Between days 7 and 9 after inoculation, the hyphae reached the aerial parts of the plant through the hypocotyl and petioles (Fig. 3d). The hyphae grew intercellularly in the leaf lamina (Fig. 3e), and sporulated on leaf surfaces through stomata, at 10 dpi (Fig. 3f).

Figure 3.

 Cytological analysis of Phytophthora parasitica growing invasively in Arabidopsis thaliana roots and leaves. (a, b) Invasive growth within the cortex, along the stele, 2 d post-inoculation (dpi). (c) Sporulation at the root surface, 5 dpi. (d) Leaf colonization from the petiole towards the limb, 7 dpi. (e) Massive intercellular invasion of the leaf by oomycete hyphae, 7 dpi. (f) Sporulation at the leaf surface through stomata, 7 dpi. (g, h) Haustorium-like structures, linked to the hyphae by a narrow neck, differentiate in root cells. These structures and intercellular hyphae were visualized by propidium iodide (PI) staining under fluorescence excitation alone (g) or with a merged image of PI staining under fluorescence excitation and differential interference contrast (DIC) (h). (i, j) Double vital staining with fluorescein diacetate (FDA) and PI, under light transmission (i) and fluorescence excitation (j) conditions, with live cells shown in green and dead cells shown in red (j), 3 dpi. (k–m) Various views of haustorium-like structures differentiating in leaf cells. (k, l) Two focal planes of leaf sections, with one plane (k) located 2 μm further into the tissue than the other (l). Haustorium-like structures in leaves and roots, measuring 4 μm in diameter (m). Intercellular leaf invasion with haustoria. (n, o) Calcofluor white staining of infected leaves showing cell wall accumulation at the base of the haustoria, as visualized by DIC (n) and fluorescence excitation (o). Micrographs were taken under light transmission conditions (d, i), with DIC images of unstained (b, c, f, k–n), PI-stained (a, g, h, j) and Calcofluor white-stained (o) tissues. Chlorazol black-stained leaf tissues (e) were analyzed by dark-field microscopy. Hyphae (Hy), haustoria (Ha) with haustorial necks (N), papillae (Pap), stomata (st) and xylem (Xy) are indicated. Bars: (b, c, f–h, k–o), 10 μm; (a, e), 50 μm; (i, j), 500 μm; (d), 50 mm.

The interaction is hemibiotrophic

Haustorium-like structures differentiated along cortex-invading hyphae, reaching their maximum abundance at 10 hpi (Fig. 3g,h). Haustoria are involved in the biotrophic development of pathogens, mediating nutrient uptake during biotrophic growth. Their formation during the early stages of infection thus indicates an initial biotrophic phase before the switch to necrotrophy. Vital staining showed that the invaded parts of the roots were still alive at 2 dpi, demonstrating the existence of a biotrophic phase during the first 2 d of infection, when hyphae expanded within the living cortex (data not shown). At 3 dpi, the infected parts of the root were dead (Fig. 3i,j). Chlorazol black staining showed that leaf-invading hyphae differentiated haustoria-like structures, before the appearance of the first macroscopic symptoms (Fig. 3k–m). Calcofluor white staining and DIC-transmitted light microscopy revealed glucan apposition and cell wall thickening at the haustorial neck in leaf cells (Fig. 3n,o). No such apposition was observed for haustoria-like structures in roots (data not shown).

Defense signaling in the roots is activated during the early stages of infection

Based on the detailed cytological analysis already described, we selected key infection stages for the analysis of the activation of plant defense pathways during the early events of root infection. We sampled A. thaliana roots at 2.5, 6, 10.5 and 30 hpi. The 2.5-hpi time point corresponded to the response of roots to initial contact with the oomycete, whereas the 6-hpi sample corresponded to the establishment of a compatible interaction (two to three layers of cortex cells colonized). As haustorium-like structures were most abundant at 10.5 hpi, this sample corresponded to the biotrophic phase of the interaction. Cell death in infected roots was initiated between 2 and 3 dpi, and the 30-hpi sample most probably corresponds to early events in the switch from biotrophy to necrotrophy. For all time points, RNA was recovered from two biological replicates, each corresponding to a pool of root sections from at least 30 infected plants. qRT-PCR experiments were then performed to assess the expression profile of plant genes encoding proteins involved in SA-, JA- or ET-mediated defense signaling.

The marker genes studied for the SA-mediated signaling pathway were ISOCHORISMATE SYNTHASE 2 (ICS2) and SALICYLIC ACID INDUCTION DEFICIENT 2 (SID2), encoding proteins involved in SA biosynthesis (Wildermuth et al., 2001; Shah, 2003; Loake & Grant, 2007). The genes encoding the PR proteins PR1, PR2 and PR5 are downstream markers of SA signaling (Wildermuth et al., 2001; Tjamos et al., 2005). In mock-inoculated plant roots, PR1 transcripts were not detectable, and SID2, ICS2, PR2 and PR5 transcripts accumulated in only small amounts (Fig. 4a,b). Transcripts of SID2, ICS2, PR1 and PR5 accumulated at 2.5 hpi, and were 18, 6, 60 and 7 times more abundant, respectively, in P. parasitica-inoculated than in mock-inoculated plants. The relative abundance of SID2 and ICS2 mRNAs varied little over these time points (Fig. 4a), whereas PR1, PR2 and PR5 transcript abundance decreased from 2.5 to 30 hpi (Fig. 4b).

Figure 4.

 Real-time reverse transcription-polymerase chain reaction (RT-PCR) profile of defense gene expression during establishment of the compatible interaction between Arabidopsis thaliana and Phytophthora parasitica. RNA was isolated 2.5 h post-inoculation (hpi), 6 hpi, 10.5 hpi and 30 hpi. Data are presented as the relative transcript abundance for genes involved in salicylic acid (SA), jasmonic acid (JA) and ethylene (ET) biosynthesis (a, c, e, respectively), and for marker genes of the SA-, JA- and ET-mediated signaling pathways (b, d, f, respectively). Transcript levels are normalized with respect to At5g11770 and At5g62050 expression, determined for the same samples. The means with error bars (± 2 SE) of two independent replicates are indicated.

We followed JA- and ET-mediated signaling events by studying 1-AMINOCYCLOPROPANE-1-CARBOXYLIC ACID SYNTHASE 2 (ACS2) and 1-AMINOCYCLOPROPANE-1-CARBOXYLIC ACID OXIDASE 4 (ACO4), which encode proteins involved in ET biosynthesis (Gomez-Lim et al., 1993; Van der Straeten et al., 1993), and FATTY ACID DESATURASE 8 (FAD8), which encodes a protein involved in JA production (Gibson et al., 1994). PR3, PR4 and PLANT DEFENSIN FAMILY 1.2 (PDF1.2) are downstream markers of ET- and JA-mediated signaling pathways (Potter et al., 1993; Chen & Bleecker, 1995; Penninckx et al., 1998). The transcription of FAD8, PDF1.2 and ACS2 was only weakly detectable in mock-inoculated roots (Fig. 4c–e). By contrast, ACO4, PR3 and PR4 displayed weak but constitutive expression in roots (Fig. 4e,f). FAD8 and PDF1.2 presented similar expression profiles, with transcripts 5 and 16 times more abundant, respectively, at 2.5 hpi in infected than in mock-inoculated plants (Fig. 4c,d). The abundance of transcripts from these genes then decreased between 2.5 and 30 hpi (Fig. 4c,d). By contrast, the expression of ACO4, ACS2, PR3 and PR4 changed little until 6 hpi, increasing gradually thereafter until 30 hpi, corresponding to induction by factors of 3, 300, 10 and 11, respectively (Fig. 4e,f).

Three different plant defence signaling pathways mediate basal resistance to P. parasitica

We assessed the contribution of the SA-, ET- and JA-mediated pathways to the interaction by analyzing the susceptibility to P. parasitica of A. thaliana mutants in which these pathways were impaired (Fig. 5). eds1-1, eds5-1 and pad4-1 mutants, and transgenic nahG plants, which have impaired SA signaling and accumulation, appeared to be significantly more susceptible than wild-type plants (Fig. 5a). By contrast, the sid2-1 mutant, in which SA synthesis is impaired, displayed a response to infection similar to that of wild-type plants (Fig. 5a). The ET-defective mutants etr1-3 and ein2-1, and the JA response mutant jar1-1, were also more susceptible than wild-type Arabidopsis to P. parasitica (Fig. 5b).

Figure 5.

 Outcome of the interaction in Arabidopsis thaliana defense-related signaling pathway mutants. Mutant and wild-type plants were inoculated with 1000 zoospores of Phytophthora parasitica strain 310. Disease severity was recorded over time, with a disease index ranging from 1 to 7. Disease progression is illustrated for mutants with impaired salicylic acid (SA)-mediated (a) and jasmonic acid/ethylene (JA/ET)-mediated (b) signaling. Results from a representative experiment are shown. Differences in disease progression between wild-type ecotypes (Col-0 or Ws) and the mutants eds5-1, pad4-1, eds1-1, ein2-1, etr1-3, jar1-1 and NahG transgenic line were statistically significant, as shown by Scheirer–Ray–Hare nonparametric two-way analysis of variance (ANOVA) for ranked data (H < 0.05). The differences between the Col-0 ecotype and the sid2-1 mutant were not statistically significant. Standard deviations (SD) for each time point are presented in Supporting Information Table S1.


The aim of this study was to provide a basis for the analysis of a compatible interaction between a plant and a root-infecting oomycete. We decided to use the model plant A. thaliana, for which many genetic and molecular tools are available. We demonstrated that P. parasitica and P. capsici successfully colonize A. thaliana roots, and are able to complete their disease cycle in this host. The two oomycetes caused similar symptoms on the plants, but symptoms developed later following the infection of A. thaliana with P. parasitica than after infection with P. capsici. As the early stages of infection were prolonged following infection with P. parasitica, we used this oomycete for detailed characterization of the compatible interaction.

The analysis of P. parasitica disease on A. thaliana and on the natural host, tomato (Le Berre et al., 2008), showed that the infection process was similar on the two plants, but that disease development was slower on A. thaliana. The first wilting symptoms appeared at 4 dpi on tomato hypocotyls, but at 7 dpi on A. thaliana plants. Moreover, P. parasitica killed the plant within 7 d of infection on tomato, whereas infected A. thaliana plants died between 15 and 20 dpi. A screen of 16 A. thaliana ecotypes with P. parasitica revealed differences in the speed of invasion, but no incompatible interaction. This finding is consistent with the ability of the P. parasitica strain used in this study to infect all tomato, tobacco and potato cultivars tested to date (F. Panabières, pers. comm.). Root resistance to pathogens currently seems to be the exception rather than the rule, because it is not observed in most pathosystems (Geraats et al., 2002; Berrocal-Lobo & Molina, 2004; Thaler et al., 2004; Okubara & Paulitz, 2005; Johansson et al., 2006; Rookes et al., 2008). It is found only in highly specialized interactions, such as those between soybean and P. sojae, between tomato and F. oxysporum f. sp. lycopersici, and between rice and Magnaporthe oryzae (Hoffman et al., 1999; Mes et al., 2000; Sesma & Osbourn, 2004; Gao et al., 2005; Okubara & Paulitz, 2005).

As shown previously for P. parasitica infections of host plant roots (Swiecki & MacDonald, 1986; Dale & Irwin, 1991; Widmer et al., 1998; Le Berre et al., 2008; Kebdani et al., 2010), A. thaliana roots were penetrated by appressoria. Within the root, hyphae developed through an initial biotrophic phase lasting 2 d after inoculation. Staining showed that, during the first 24 hpi, haustorium-like structures developed within living host root cells. These structures met the criteria for true haustoria (Bushnell, 1972), providing the first experimental evidence for the occurrence of a biotrophic phase during the root life cycle of P. parasitica. Phytophthora species are generally described as hemibiotrophic pathogens, but, for root infections, true haustoria have been observed previously only during a short intracellular biotrophic phase preceding necrotrophy, in the interaction between soybean and P. sojae (Enkerli et al., 1997).

By contrast with the abundance of knowledge about defense mechanisms in leaves, little is known about the genetic basis of root responses to soil-borne pathogens. Using A. thaliana mutants impaired in the SA, JA and ET defense signaling pathways, we showed that all but one (sid2-1) of the mutants was more susceptible than the wild-type to P. parasitica. The absence of a sid2-1 mutant phenotype may be accounted for by potential functional redundancy between ICS2 and SID2, both of which encode an isochorismate synthase. Small amounts of SA are produced in sid2-1 mutants and this may slow P. parasitica infection (van Wees & Glazebrook, 2003). Some of the previously described phenotypes of NahG plants have been attributed to the accumulation of catechol (van Wees & Glazebrook, 2003). However, the enhanced oomycete susceptibility observed on these transgenics concurred perfectly with a similar phenotype of eds mutants, thus correlating with the deficiency in SA signaling rather than with an accumulation of catechol. Previous studies analyzing root defense responses to oomycetes and fungi have generated conflicting results. Mutations affecting ET-mediated or JA-mediated signaling in A. thaliana have no effect on the response to the oomycete P. cinnamomi, but such mutants of tobacco and Arabidopsis are more susceptible to Pythium species and to fungi such as Verticillium longisporum, F. oxysporum and Rhizoctonia solani (Staswick et al., 1998; Vijayan et al., 1998; Hoffman et al., 1999; Geraats et al., 2002; Rookes et al., 2008). ET- and JA-dependent defenses seem to be frequently triggered in root responses to filamentous pathogens, but SA involvement does not seem to be a general feature of these responses. SA mutants of A. thaliana display wild-type phenotypes during the interaction with the oomycete P. cinnamomi and the fungus V. longisporum (Johansson et al., 2006; Rookes et al., 2008). In Arabidopsis root–microbe interactions, the activation of SA signaling has been reported only for the interaction between A. thaliana roots and the ascomycete F. oxysporum (Liljeroth et al., 2001; Berrocal-Lobo & Molina, 2004). Our results for P. parasitica are thus consistent with published data indicating that ET/JA signaling pathways are activated during the infection of roots with filamentous pathogens. They also provide the first evidence that SA-mediated pathways are activated in roots during the interaction with an oomycete. We found that SA-, ET- and JA-mediated pathways limited, but could not prevent, infection.

We used the A. thalianaP. parasitica root interaction system to analyze the activation profile of marker genes for the SA, ET and JA signaling pathways during infection, because we were able to cover biotrophy and the switch to necrotrophy. Transcript accumulation was evaluated during: (1) penetration of the first cell layer by P. parasitica; (2) the establishment of biotrophic invasive growth; (3) established invasive growth along the stele; and (4) early in the switch to necrotrophy. Relative transcript levels for genes encoding proteins involved in SA or JA biosynthesis, and for downstream marker genes of the SA and JA pathways, increased transiently when the pathogen penetrated the rhizodermis. By contrast, sustained upregulation of the genes encoding proteins involved in ET biosynthesis and signaling was observed throughout the invasive growth of P. parasitica within roots. Genetic and molecular data are currently available for the invasive growth phase of plant–pathogen interactions, but not for penetration (Hu et al., 2008; Huibers et al., 2009; Schlink, 2009). The earliest stage assessed to date is the onset of infection during the interaction between A. thaliana and Alternaria brassicola and between Fagus sylvatica and Phytophthora citricola in leaves and/or roots (Schenk et al., 2003; Zimmerli et al., 2004; Schlink, 2009). Defense gene transcripts were found to have accumulated 6 h after infection, a time point coinciding with the establishment of invasive growth.

During leaf infections, switches from biotrophy to necrotrophy are frequently accompanied by a shift of plant defenses from SA- to JA-mediated responses (Glazebrook, 2005). During the infection of roots with P. parasitica, a transient accumulation of transcripts for marker genes of both the SA- and JA-dependent pathways occurred during penetration. These findings suggest that the JA and SA signaling pathways cooperate in the defenses induced in A. thaliana roots challenged with P. parasitica. Similar observations have been reported following the inoculation of A. thaliana roots with the fungus F. oxysporum (Berrocal-Lobo & Molina, 2008). These findings contrast with the reported antagonistic action of the signaling pathways involving SA and JA/ET in leaves (Glazebrook, 2005), demonstrating that the extrapolation of findings from one plant organ to another might lead to misleading generalizations. For a full understanding of the molecular mechanisms governing root interactions with soil-borne pathogens, experiments must therefore be performed on plant roots. The pathosystem described, based on interactions between A. thaliana roots and P. parasitica, thus provides a unique tool for the molecular analysis of the mechanisms underlying the interaction between roots and soil-borne oomycetes. Such analysis may make use of the genomic tools available for the model plant and of the forthcoming genomic sequence of P. parasitica (F. Panabières, pers. comm.). As all steps in the process of infection in A. thaliana are similar to those observed in the host plant, tomato, the knowledge obtained in this model pathosystem may provide clues to the possible mechanisms underlying infectious oomycete root diseases in agriculturally important solanaceous plants. In leaves, early defense responses are of prime importance, determining the outcome of an interaction, because the principal differences between compatibility and incompatibility are observed once the pathogen has crossed two to three cell layers (Roetschi et al., 2001). We found that P. parasitica infection triggered defense responses in A. thaliana roots from the moment at which the pathogen first penetrated the roots. The transient activation of SA- and JA-mediated responses may reflect an ability of P. parasitica to overcome the defense signaling pathways coordinated by these hormones. The decrease in the expression of hormone-related genes may reflect the manipulation of host responses by pathogen effectors (Schornack et al., 2009).


We would like to thank Nicolas Ris (INRA, Sophia Antipolis, France) for statistical analysis and Gilbert Engler for his help with microscopic analysis. We are grateful to Dr Jane Parker (Max-Planck-Institut für Züchtungsfors-chung, Cologne, Germany), Dr Leslie Friedrich (Research Triangle Park, NC, USA), Dr Yves Marco and Dr Claudine Balagué (INRA-CNRS, Toulouse, France), and the Nottingham Arabidopsis Stock Centre, for providing seeds of Arabidopsis mutants and transgenics.