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PLANTS INTERACTING WITH OTHER ORGANISMS

Genetic Elucidation of Nitric Oxide Signaling in Incompatible Plant-Pathogen Interactions^[w]

John Innes Centre, Norwich Research Park, Colney, Norwich NR4 7UH, United Kingdom (J.Z., E.S., C.L.); Dipartimento Scientifico e Tecnologico, Università degli Studi di Verona, 37134 Verona, Italy (M.D., E.S.); and Julius-von-Sachs-Institut für Biowissenschaften, Lehrstuhl für Botanik II, Universität Würzburg, D–97082 Wurzburg, Germany (J.Z., T.M., M.S.)

	ABSTRACT

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 
Recent experiments indicate that nitric oxide (NO) plays a pivotal role in disease resistance and several other physiological processes in plants. However, most of the current information about the function of NO in plants is based on pharmacological studies, and additional approaches are therefore required to ascertain the role of NO as an important signaling molecule in plants. We have expressed a bacterial nitric oxide dioxygenase (NOD) in Arabidopsis plants and/or avirulent Pseudomonas syringae pv tomato to study incompatible plant-pathogen interactions impaired in NO signaling. NOD expression in transgenic Arabidopsis resulted in decreased NO levels in planta and attenuated a pathogen-induced NO burst. Moreover, NOD expression in plant cells had very similar effects on plant defenses compared to NOD expression in avirulent Pseudomonas. The defense responses most affected by NO reduction during the incompatible interaction were decreased H₂O₂ levels during the oxidative burst and a blockage of Phe ammonia lyase expression, the key enzyme in the general phenylpropanoid pathway. Expression of the NOD furthermore blocked UV light-induced Phe ammonia lyase and chalcone synthase gene expression, indicating a general signaling function of NO in the activation of the phenylpropanoid pathway. NO possibly functions in incompatible plant-pathogen interactions by inhibiting the plant antioxidative machinery, and thereby ensuring locally prolonged H₂O₂ levels. Additionally, albeit to a lesser extent, we observed decreases in salicylic acid production, a diminished development of hypersensitive cell death, and a delay in pathogenesis-related protein 1 expression during these NO-deficient plant-pathogen interactions. Therefore, this genetic approach confirms that NO is an important regulatory component in the signaling network of plant defense responses.

One of the earliest events following pathogen recognition is a burst of oxidative metabolism leading to the generation of superoxide () and subsequent accumulation of H₂O₂ (Lamb and Dixon, 1997). These ROI are directly protective and drive the oxidative cross-linking of cell wall structural proteins (Brisson et al., 1994). The H₂O₂ originating from the oxidative burst induces some plant genes involved in cellular protection and defense such as gluthathione S-transferase (GST) and is necessary for the initiation of host cell death following the HR (Levine et al., 1994).

Recent pharmacological experiments indicate that nitric oxide (NO), which acts as a signal in the immune, nervous, and vascular system in vertebrates (Schmidt and Walter, 1994), also plays an important role in plant disease resistance. Generation of NO by chemical NO donors augments the induction of hypersensitive cell death by H₂O₂ in soybean (Glycine max) suspension cultures (Delledonne et al., 1998, 2001). Likewise, inhibitors of NO synthesis as well as NO scavengers are able to block the HR induced by avirulent Pseudomonas syringae in soybean cell cultures and in Arabidopsis plants. Compared to ROI, NO induces a complementary set of plant defense genes, including two key enzymes of the phenylpropanoid pathway, namely Phe ammonia lyase (PAL) and chalcone synthase (CHS). Furthermore, NO-treated tobacco (Nicotiana tabacum) cells were shown to induce the pathogenesis-related protein 1 (PR-1) together with an accumulation of SA (Durner et al., 1998), a key molecule for the expression of systemic acquired resistance (Gaffney et al., 1993). Moreover, the molecular components of NO signaling in plants appear to be similar to those in animals, regarding the involvement of NO producing NO synthases (NOS; Chandok et al., 2003; Guo et al., 2003) and cGMP as a second messenger (Clark et al., 2000).

As an increasing number of recent reports suggests, a regulatory function of NO in plants seems to be essential in other physiological processes, including guard cell abscisic acid signaling (Desikan et al., 2002; Garcia-Mata and Lamattina, 2003), regulation of iron homeostasis (Graziano et al., 2002; Murgia et al., 2002), execution of programmed cell death in barley (Hordeum vulgare) aleurone layers (Beligni et al., 2002), root organogenesis (Pagnussat et al., 2002), and wound signaling (Orozco-Cardenas and Ryan, 2002). However, despite the recent identification of a pathogen-inducible NOS (Chandok et al., 2003), assumptions of NO function in plants emerging from all these studies are almost exclusively based on pharmacological studies, i.e. either exogenous application of NO donors, NO scavengers, and inhibitors of mammalian NOS or detection of NO by essentially indirect methods using fluorescent dyes or photometric indicator molecules (Delledonne et al., 1998; Foissner et al., 2000). If indeed the application of pharmacological compounds reflects a physiological NO situation without exerting nonspecific side effects is far from clear, and additional experimental approaches are therefore desirable.

We report here a novel genetic approach to manipulate NO levels in planta, which has been used to gain a better understanding of the function of NO in the signaling network underlying incompatible plant-pathogen interactions. We first generated transgenic Arabidopsis plants overexpressing the Escherichia coli hmp gene encoding NO dioxygenase (NOD), a flavohemoglobin capable of converting NO to nitrate by use of NAD(P)H and O₂ (Vasudevan et al., 1991; Gardner et al., 1998; Poole and Hughes, 2000). In this way, we attempted to directly reduce the levels of NO in plant cells. We then compared the defense responses of NO-deficient plants and wild-type Arabidopsis following challenge with avirulent P. syringae pv tomato (Pst) bacteria. Additionally, we employed avirulent Pst expressing the hmpX gene from Erwinia chrysanthemi [Pst(avr-hmpX)] (Favey et al., 1995) that encodes a highly similar NOD to lower NO levels specifically at the side of pathogen infection. We then challenged wild-type Arabidopsis and hmp-expressing Arabidopsis plants with Pst(avr-hmpX) to study the effect of NO removal at both the plant and the pathogen side.

	RESULTS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 

Arabidopsis Plants Expressing a Bacterial NOD

For the production of Arabidopsis ecotype Col-0 plants expressing a functional NOD, the hmp coding sequence from E. coli (Vasudevan et al., 1991) was cloned into the dexamethasone (DEX)-based pTA7001-inducible vector system (Aoyama and Chua, 1997). After Agrobacterium-mediated transformation of wild-type Col-0 plants with the vector construct, homozygous T3 plants segregating for a single T-DNA insertion were used for further experiments. Upon treatment with 3 µM DEX, the various lines expressed the hmp transgene at different levels (data not shown). We selected one of the lines that expressed hmp to a higher degree, designated hmp8, for more detailed analysis. In hmp8 plants, hmp transcripts accumulated at 5 h after spraying rosette leaves with DEX (Fig. 1A). Transcriptional levels increased 1 d after DEX treatment and remained nearly constant for at least 1 week. Western-blot analysis with antibodies raised against E. coli Hmp revealed the production of a full-size Hmp (43 kD) protein in planta (Fig. 1B). The kinetics of Hmp protein expression was similar to transcriptional hmp induction.

View larger version (28K): [in this window] [in a new window]   Figure 1. Expression of E. coli hmp in transgenic Arabidopsis as a functional NOD. A, Northern-blot analysis illustrating time-dependent accumulation of hmp transcripts after treatment of hmp8 plants with 3 µM DEX (c, no DEX). B, Western blot demonstrating the appearance of correctly sized Hmp protein in leaf extracts; time course as in A. C and D, Electrochemically measured degradation kinetics of 10 µM NO in leaf extracts of wild-type and hmp8 transgenic Arabidopsis. Error bars represent the SDs of five independent measurements. C, Hmp8 plants without DEX-induced transgene expression. D, Wild-type and hmp8 plants 1 d after DEX treatment. NO concentrations of less than 1 µM were reached for wild-type, noninduced hmp8, and DEX-induced hmp8 plants at 197 s, 202 s, and 131 s, respectively. E, NO emission from intact wild-type and hmp8 plants 1 d after DEX treatment. Plants were first incubated in the dark for 10 min and then illuminated for 60 min to cause nitrate reductase-dependent NO emission. The light was switched off again, which gave rise to a characteristic light-off peak (see "Results"). Experiments were repeated three times with similar results.

In leaves, NO can be produced from nitrite by nitrate reductase, and this NO production is measurable as emission by chemiluminescence (Rockel et al., 2002). A temporary rise of NO emission resulting from increased nitrate reductase activity was detected when dark-adapted plants were transferred to light. When the light source was switched off again, a light-off peak caused by transient nitrite accumulation resulted (Kaiser et al., 2002). Both the light-induced NO emission and the NO light-off peak were significantly lower in DEX-treated hmp8 plants compared to the wild type (Fig. 1E).

P. syringae Expressing a Bacterial NOD

To expand our genetic approach to study NO deficiency in incompatible plant-pathogen interactions, we transformed Pst (avrB) with the hmpX gene from E. chrysanthemi. Like Hmp from E. coli, E. chrysanthemi HmpX represents a NOD (M. Delledonne and R. Poole, unpublished data). The expression of HmpX in the bacterium should decrease the concentration of the diffusible molecule NO specifically at the site of pathogen infection. In this way, we were able to study plant-pathogen interactions in which NO is simultaneously removed at the infection site from both the plant and the pathogen side.

Plant Defense Responses under NO-Deficient Conditions

Pst carrying the avrB avirulence gene is recognized by Arabidopsis ecotype Col-0 carrying the Rpm1 resistance gene (Bisgrove et al., 1994). The recognition results in the oxidative burst, a production of ROI like H₂O₂ and at the site of infection, induction of defense and cellular protectant genes, development of the HR, and a limitation of pathogen growth in comparison to isogenic virulent Pst.

NO Production

To investigate whether NO is produced during the infection of Col-0 plants with avirulent bacteria, we infiltrated DEX-treated wild-type leaves with the NO-sensitive, cell-permeable fluorescent dye 4,5-diaminofluorescein diacetate (DAF2-DA; Foissner et al., 2000) 3 h after challenge with 2 x 10⁶ colony-forming units (cfu) mL^–1 Pst (avrB). Inside the area of pathogen infiltration, various brightly green fluorescing cell groups were discernible 1 h after DAF2-DA treatment (Fig. 2C, white stars), whereas MgCl₂-infiltrated control leaves only showed a weak fluorescence when treated with the fluorophore for the same period of time (Fig. 2B). Pathogen infiltration without fluorophore treatment did not cause any fluorescence (Fig. 2A). As compared to wild-type leaves, DEX-induced hmp8 plants challenged with Pst (avrB) and subsequently treated with DAF2-DA showed a markedly reduced fluorescence at the site of pathogen infiltration (Fig. 2D). A similar reduction of DAF2-DA fluorescence was observed when wild-type leaves were challenged in the presence of 100 µM CPTIO, a NO scavenging compound (data not shown). These results suggest that NO is produced during the earlier stages of the Arabidopsis-Pst (avrB) interaction and that this NO burst is attenuated in NOD-expressing hmp8 plants.

View larger version (181K): [in this window] [in a new window]   Figure 2. Pathogen-induced DAF2-DA fluorescence as a measure for NO production in DEX-treated wild-type and hmp8 plants. Leaves were pretreated with Pst (avrB) or MgCl2 for 3 h and subsequently infiltrated with 10 µM DAF2-DA or control buffer (10 mM Tris/KCl, pH 7.2). Infiltrated leaf areas were analyzed 1 h later by confocal laser scanning microscopy. DAF2-DA fluorescence (green) was recorded using a channel with a 505- to 530-nm band-pass filter, and autofluorescence of chloroplast (red) was captured with a channel equipped with a 560-nm long-pass filter. A, Treatment of a wild-type Arabidopsis leaf with Pst (avrB) and control buffer. B, Wild-type Arabidopsis-MgCl2 and DAF2-DA. C, Wild-type Arabidopsis-Pst (avrB) and DAF2-DA. D, Hmp8-Pst (avrB) and DAF2-DA. Seven independent samples were recorded for each condition, and representative leaf areas are shown.

To assess the accumulation of ROI in response to infection with an avirulent pathogen, DEX-treated leaves from wild-type and hmp8 plants were infiltrated with 2 x 10⁶ cfu mL^–1 Pst (avrB) and stained with diaminobenzidine (DAB), a histochemical reagent that forms a reddish-brown precipitate upon contact with H₂O₂ (Thordal-Christensen et al., 1997). Wild-type leaves showed a strong production of H₂O₂ at the infection site 4 h after pathogen challenge (Fig. 3). By contrast, leaves from hmp8 plants showed significantly lower levels of H₂O₂ compared to the wild type when challenged with Pst (avrB). Moreover, when DEX-treated leaves from hmp plants where challenged with Pst (avrB/hmpX) bacteria, the suppression of H₂O₂ levels during the oxidative burst was even more pronounced.

View larger version (55K): [in this window] [in a new window]   Figure 3. DAB staining of Arabidopsis leaves to assess H2O2 accumulation during the oxidative burst in DEX-treated wild-type and hmp8 transgenic plants. Solutions of Pst (avrB) were infiltrated into Arabidopsis leaves, and DAB staining was initiated 4 h after infection. A, Staining patterns of representative, MgCl2-infiltrated wild-type or hmp8 leaves (control), Pst (avrB)-infected wild-type Arabidopsis leaves (Wt-avr), Pst (avrB)-infected hmp8 plants (hmp8-avr), and Pst (avrB/hmpX)-infected hmp8 plants (hmp8-avr/hmpX) 4 h after the respective treatment (100-fold magnification). B, Quantification of DAB staining in Pst (avrB)-infected wild-type and hmp8 leaves. The percentage of stained pixels inside the infiltration area was assessed as described in "Materials and Methods." Values are shown as the mean ± SD of at least five leaves from different plants. Experiments were repeated three times with similar results.

View larger version (73K): [in this window] [in a new window]   Figure 4. DAB staining of Arabidopsis leaves to assess their capability to degrade H2O2 in DEX-treated wild-type and hmp transgenic plants. Solutions of 2.5 mM Glc/2.5 units mL–1 Glc oxidase were infiltrated into leaves and DAB staining was performed 1 h after infiltration. A, Staining patterns of representative leaves inside the infiltrated area (100-fold magnification). B, Percentage of stained pixels inside the infiltration zone. Values are shown as the mean ± SD of at least five leaves from different plants. Experiments were repeated three times with similar results.

We next examined whether the expression of three typical defense-related genes, GST, PAL, and PR-1, was affected in the hmp8 line (Fig. 5). GST functions in cellular protection, and gst transcripts are induced during the oxidative burst (Levine et al., 1994; Delledonne et al., 2001). Gst transcripts accumulated 4 to 10 h after DEX-treated wild-type plants were challenged with Pst (avrB). Similar induction kinetics were observed when DEX-induced hmp8 plants were challenged with Pst (avrB) or Pst (avrB/hmpX). Only in the latter case was the amount of gene induction slightly diminished, presumably reflecting the extremely low H₂O₂ levels during the oxidative burst.

View larger version (42K): [in this window] [in a new window]   Figure 5. Expression of defense and cellular protectant genes in wild-type and hmp8 transgenic plants after Pst (avrB) challenge. Three parallel leaf samples were collected at the indicated times after infection for RNA extraction and northern-blot analysis. MgCl2 (m)-infiltrated leaves were collected 4 h after infection. Experiments were repeated three times with similar results.

Transcriptional induction of the antimicrobial PR-1 protein occurred 10 h after challenge of both wild-type and hmp8 plants with Pst (avrB) (Fig. 5). PR-1 induction was delayed, albeit not fully suppressed, when hmp8 plants were challenged with Pst (avrB/hmpX).

The finding that pathogen-induced pal expression is blocked under NO-deficient conditions prompted us to test whether NO could act as a general signal for the activation of the phenylpropanoid pathway. Another stimulus activating this pathway is UV light (Chappell and Hahlbrock, 1984), and we examined the UV-induced expression of PAL and CHS, the key enzyme for flavonoid biosynthesis, in DEX-treated wild-type and hmp8 plants (Fig. 6). Whereas wild-type plants strongly expressed chs and pal after 24 h of UV treatment, this UV-induced gene expression was markedly attenuated in hmp8 plants, indicating that NO is required for phenylpropanoid pathway activation by different environmental stimuli.

View larger version (57K): [in this window] [in a new window]   Figure 6. Expression of PAL and CHS in DEX pretreated wild-type and hmp8 plants after UV exposure. Three parallel leaf samples were collected at the indicated times after the beginning of UV treatment. Experiments were repeated three times with similar results.

When leaves from wild-type plants were challenged with Pst (avrB) at concentrations of 5 x 10⁶ cfu mL^–1, a dry, colorless lesion limited to the site of pathogen infiltration developed within 2 d in at least 5 out of 7 leaves (Fig. 7A). These macroscopic symptoms characteristic of hypersensitive cell death developed to the same extent when noninduced hmp8 plants were used (data not shown). However, when infected with Pst (avrB), DEX-treated hmp8 plants showed a more chlorotic lesion that was reminiscent of the symptoms caused by isogenic virulent Pst in about 50% of leaves (Fig. 7B). Infection of induced hmp8 plants with Pst (avrB/hmpX) further increased this percentage. In this case, virtually every challenged leaf developed chlorotic symptoms (Fig. 7C).

View larger version (82K): [in this window] [in a new window]   Figure 7. Macroscopic HR symptoms 2 d after infiltration of DEX-pretreated wild-type and hmp8 plants with avirulent Pseudomonas. Bacteria were infiltrated into the left side of leaves. Seven parallels are shown for each condition. A, Wild-type plants-Pst (avrB). B, hmp8 plants-Pst (avrB). C, hmp8 plants-Pst (avrB/hmpX).

View larger version (88K): [in this window] [in a new window]   Figure 8. Microscopic cell death after pathogen challenge of DEX-pretreated wild-type and hmp8 plants. Trypan blue staining was performed 24 h after infection. A, Staining patterns of representative leaves inside the infiltrated area (100-fold magnification). B, Percentage of stained pixels inside the infiltration area. Values are shown as the mean ± SD of at least six leaves from different plants. Experiments were repeated three times with similar results.

SA plays a central role in the activation of plant defense responses, and a relationship between NO and SA signaling pathways has been discussed (Klessig et al., 2000). To determine the effects of NO on SA levels in the Pst (avrB)-Col-0 interaction, we determined the levels of SA and SA glucoside (SAG) in our system 8 h after pathogen infection (Fig. 9). DEX-treated wild-type and hmp8 plants infiltrated with 10 mM MgCl₂ had SA levels of about 120 ng g^–1 leaf fresh weight and SAG contents of about 600 ng g^–1. Upon Pst (avrB) challenge, SA levels increased in wild-type plants about 4-fold to 500 ng g^–1, and SAG levels reached values of 1,700 ng g^–1, representing a 3-fold increase. In DEX-treated hmp8 plants, the SA content in Pst (avrB)-challenged leaves showed a small, albeit statistically not significant, decrease to about 80% of the wild-type value (Fig. 9A). A similar trend was observed for the glucoside (Fig. 9B). Again, the most striking effect was detected when induced hmp8 plants were infiltrated with Pst (avrB/hmpX). Here, SA and SAG were reduced to about 50% of the wild type-Pst (avrB) values, reflecting a pathogen-induced increase of only about 2-fold (SA) and 1.5-fold (SAG), respectively.

View larger version (15K): [in this window] [in a new window]   Figure 9. SA contents of wild-type and hmp8 transgenic plants after challenge with avirulent Pst (± hmpX). Leaf samples were collected 8 h postinfection. Leaves were pretreated with DEX for 16 h. Bars indicated mean values of three independent measurements. Control, MgCl2-infiltrated plants. A, Free SA. B, SAG.

	DISCUSSION

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

Transgenic Arabidopsis plants were produced that express the Hmp protein from E. coli, and whole-plant NO emission was evaluated. The emission was shown to be markedly reduced in Hmp-expressing plants, and leaf extracts from transgenic plants degraded NO significantly faster than extracts from control plants. These findings demonstrate that Hmp is a functional NOD in planta (Fig. 1). Using the NO-sensitive fluorescence indicator DAF2-DA, we furthermore showed that NO is produced during the incompatible interaction of Arabidopsis and Pst (avrB) and that this NO burst is attenuated in NOD-expressing plants (Fig. 2). We also produced avirulent Pseudomonas expressing HmpX, a similar NOD from E. chrysanthemi. Biochemical experiments suggest that HmpX is located in the periplasm and represents a functional NOD in transformed Pseudomonas (R. Poole and M. Delledonne, unpublished data).

With these genetic tools, we examined the hypersensitive disease resistance response when NO accumulation was attenuated by the action of NODs in two different surroundings. We could generally state that the removal of NO from the plant resulted in strikingly similar tendencies compared to NO removal from the pathogen side. When comparing the interaction of hmp8 plants with Pst (avrB) on the one hand and the interaction of wild-type plants with Pst (avrB/hmpX) on the other hand, we found very similar tendencies (data not shown). Moreover, when combining the two genetically modified systems, i.e. the interaction of hmp8 plants with Pst (arvB/hmpX), we observed additive effects in all examined defense responses.

We first detected significantly lower H₂O₂ levels during the oxidative burst in NO-deficient interactions, and removal of NO from both the plant and the pathogen side had an additive effect (Fig. 3). Less H₂O₂ staining in the presence of NOD was also obvious when plants were infiltrated with Glc/Glc oxidase, a H₂O₂-generating system (Fig. 4), indicating that the in planta capability to degrade H₂O₂ was increased by the action of the NOD. This effect might be due to the NO-degrading function of the NOD or possibly by direct degradation of H₂O₂ by NOD. Because the rate of H₂O₂ degradation was identical in leaf extracts from wild-type and hmp8 plants (data not shown), we conclude that direct H₂O₂ degradation through NOD does not take place. Rather, a factor differing in hmp8 and wild-type plants but not in the corresponding extracts might account for the different observation in intact plants and extracts, respectively. This factor might be the concentration of NO, which is delivered by the intact plant continuously but not necessarily by extracts. Following this interpretation, the higher in planta H₂O₂ degradation capability of NO-deficient Hmp plants suggests an inhibitory effect of NO toward H₂O₂-degrading enzymes. In fact, the predominant H₂O₂ scavenging enzymes are catalase and ascorbate peroxidase. Mammalian catalase is reversibly inhibited by NO (Brown, 1995), and it has been shown in vitro that NO inhibits both tobacco catalase and ascorbate peroxidase (Clark et al., 2000). In accordance to previous reports (Delledonne et al., 1998; Foissner et al., 2000), our data reveal that a local burst of NO coincides with the oxidative burst at the site of pathogen infection. Therefore, it is conceivable that this NO burst locally contributes to maintain more sustained and higher H₂O₂ levels that can then either act directly as an antimicrobial oxidant or indirectly by triggering various defense responses (Lamb and Dixon, 1997).

The most striking differences in defense gene expression concerned the induction of pal transcripts, which was significantly attenuated in hmp plants and in the presence of avirulent Pseudomonas expressing HmpX (Fig. 5). This observation confirms pharmacology-based findings demonstrating a reduction of pal expression by NOS inhibitors in soybean cells and pal induction by NO donors and recombinant NOS in soybean and tobacco, respectively (Delledonne et al., 1998; Durner et al., 1998). In addition, the UV-induced expression of pal and chs, the first committed enzyme in anthocyanin biosynthesis, was strongly repressed in hmp plants (Fig. 6). Based on this experimental evidence, NO appears to play a pivotal regulatory role in the signaling processes leading to expression of phenylpropanoid pathway genes.

PAL is the key enzyme for the general phenylpropanoid pathway, and possible outcomes are lignin, anthocyanin, and/or SA biosynthesis. However, despite the strong repression of pal, SA levels only showed a 20% reduction in the presence of Hmp in plants or HmpX in bacteria (Fig. 9). This supports the findings that in Arabidopsis, SA is produced by alternative routes, e.g. by the isochorismate pathway (Wildermuth et al., 2001). The observed induction of pal in the Pseudomonas-Arabidopsis pathosystem might then feed alternative processes like lignification or the production of other phenylpropanoid compounds to a significant extent. If SA is the causative agent of PR-1 induction (Yalpani et al., 1991; Uknes et al., 1992), the rather weak attenuation of SA induction might explain the fact that in these cases the expression of PR-1 is not affected (Fig. 5). The levels of SA might still be above a threshold value necessary for full pathogen-induced PR-1 expression. In the double experiment in which NO is reduced from both the pathogen and the plant, however, SA was reduced to 50% of the usual value and PR-1 expression was clearly delayed, although not fully suppressed. It is worth noting that the expression of PR-1 is also up-regulated in plants challenged with virulent pathogens but slower than in the corresponding incompatible interaction (de Torres et al., 2003). Compared to pharmacological experiments with tobacco and soybean cells (Delledonne et al., 1998; Durner et al., 1998), our results demonstrate a similar tendency of NO involvement in SA and PR-1 production and a strong regulatory role of NO toward the synthesis of phenylpropanoid pathway enzymes like PAL.

When NO is scavenged by Hmp or HmpX alone, dry lesion development is delayed but not eliminated (Fig. 7), and the appearance of microscopic HR lesions is only moderately suppressed (Fig. 8). However, when NO is scavenged by the simultaneous action of Hmp and HmpX, the macroscopic dry HR lesions are yellowish with less pronounced symptoms, and the microscopic HR lesions are significantly reduced (Fig. 8). Therefore, HR lesion development is clearly affected by a reduced NO content in Arabidopsis, and the correlation with SA levels suggests a mediatory role of SA in these processes. This is in accordance with findings that SA is required for induction of the HR in response to bacterial pathogens in soybean (Tenhaken and Rubel, 1997) and that SA is needed for cell death initiation in Arabidopsis lsd mutants (Weymann et al., 1995). The execution of hypersensitive cell death in soybean cells challenged with avirulent P. syringae is strongly diminished by NO scavenging compounds and NOS inhibitors (Delledonne et al., 1998). Furthermore, a poised production of ROI and NO is necessary to trigger the HR, and NO together with H₂O₂, but not , are indicated as the essential players in this process (Delledonne et al., 2001). One possible mechanism for this cooperation could be that NO ensures maintenance of high, persistent H₂O₂ levels that are necessary to trigger the HR. The threshold H₂O₂ levels for HR development in soybean cells is about 6 µM (Levine et al., 1994), and in our experiments, H₂O₂ levels may not have significantly fallen below a comparable threshold value in the cases of Hmp or HmpX action alone. In the double experiment, however, when H₂O₂ levels showed the strongest decrease (Fig. 3), the H₂O₂ threshold might not have been exceeded and, consequently, a significant reduction in HR development could be observed. A similar reasoning could explain why a significant induction of gst transcription still took place under NO deficiency, even in the double experiment (Fig. 5). For gst expression, H₂O₂ threshold levels have been shown to be below the HR value, which are around 2 µM in soybean cells (Levine et al., 1994).

Blockage of the accumulation of NO by NO scavengers or mammalian NOS inhibitors has previously been shown to enhance bacterial growth of avirulent Pseudomonas in Arabidopsis leaves, although to a lower extent in comparison to the growth of virulent strains (Delledonne et al., 1998). In this work, NO scavenging by NOD led to a similar, albeit weaker, tendency of bacterial growth enhancement of avirulent Pst (Table I). Obviously, the removal of NO in these and other recent experiments is not sufficient for a strong growth enhancement of avirulent pathogens, despite the apparent effects on progression of the described defense responses. This behavior is in some respects reminiscent of the dnd (defense, no death) class of Arabidopsis mutants that show the ability to limit pathogen growth in a gene-for-gene resistance without developing an HR (Yu et al., 1998). Dnd1 encodes a cyclic nucleotide-gated ion channel (Clough et al., 2000). In mammals, cyclic nucleotides like cGMP or cADP-Rib are second messengers closely associated with NO signaling (Galione and White, 1994; Schmidt and Walter, 1994), and this has been suggested for plants as well (Pfeiffer et al., 1994; Durner et al., 1998; Clarke et al., 2000). It will be of interest to investigate if generation of NO leads to activation of Dnd1, which contributes to the development of symptoms characteristic of the HR.

The complete scavenging of a highly diffusible and reactive molecule like NO is difficult to achieve for a single protein, and the (physiologically active) reaction of NO with other cellular molecules might to a certain extent still compete with the NOD reaction. This is also reflected by the fact that Hmp-overexpressing plants still emitted about one-half the amount of gaseous NO than wild-type plants (Fig. 1E). Therefore, not all cellular NO-mediated effects might have been fully suppressed by this transgenic approach. Compared to the very similar salicylate hydroxylase (NahG) strategy applied by Gaffney et al. (1993), which addressed the role of the far less reactive signaling compound SA in disease resistance, these data have some limitations due to the physicochemical properties of NO. In general, the possibility that protein overexpression in plants leads to side effects not associated with the physiological process under investigation cannot be fully excluded. For instance, it was demonstrated recently that effects on plant defense responses observed by overexpression of NahG might not necessarily result from a lack of SA accumulation but could partly be a consequence of the presence of the SA degradation product catechol (van Wees and Glazebrook, 2003). Because Hmp converts NO, O₂, and NAD(P)H to nitrate in equistoichiometric amounts, it is conceivable that physiological side effects resulting from oxygen and NAD(P)H consumption or nitrate accumulation exist in hmp-overexpressing plants. Such side effects, however, should be minimal in hmp plants considering the comparatively low NO levels produced in plants (Fig. 1E) and taking into account the use of an inducible vector system, which restricted the action of Hmp to a small experimental window. The parallel tendencies observed in two different genetic backgrounds, i.e. the action of Hmp in Arabidopsis and HmpX in Pseudomonas, further support that the described effects on plant defenses were a direct consequence of NO degradation rather than a result of indirect effects caused by NOD on plant metabolism. Besides its NOD activity, however, it was shown that Hmp-overexpressing E. coli strains are capable of generating ROI (Poole and Hughes, 2000). We can rule out that this metabolic activity took place in transgenic hmp plants during the course of our pathogen and UV experiments due to the findings that transcriptional up-regulation of gst, a sensitive marker for ROI production (Levine et al., 1994), and a positive DAB reaction could not be detected in uninfected but DEX-induced hmp plants (Figs. 3 and 5). The latter statement, however, could not be maintained when hmp plants were exposed to DEX for longer periods of time. About 4 d after transgene induction, we detected elevated levels of ROI in leaves of hmp plants by DAB staining and the expression of gst and pr-1 in the absence of pathogens (data not shown). Obviously, a new physiological situation in hmp plants appeared when plants perpetually accumulated Hmp protein, either caused by the constant removal of NO or by an emerging ROI-generating activity. Interestingly, in a recent transgenic approach, the constitutive overexpression of an alfalfa (Medicago sativa) hemoglobin with putative NO scavenging properties in tobacco similarly resulted in increased basal ROI levels, besides elevated pathogen-induced SA levels and reduced disease symptoms after P. syringae infection (Seregélyes et al., 2003). These findings indicate that a temporally controlled expression rather than constitutive or prolonged overexpression of certain transgenes like (flavo) hemoglobins might be crucial in overexpression studies. The use of the inducible vector system permitted us to use a window of 3 to 4 d after transgene expression to perform pathogen and physiological experiments without undesirable side effects like ROI production.

	MATERIALS AND METHODS

TOP ABSTRACT RESULTS DISCUSSION MATERIALS AND METHODS LITERATURE CITED

 

Generation of Hmp-Overexpressing Arabidopsis

To generate transgenic Arabidopsis overexpressing the Escherichia coli hmp gene, pTA7001, a dexamethasone-inducible expression system, was used (Aoyama and Chua, 1997). The hmp coding sequence was generated by PCR with the primers 5'-CGGCTCGAGATGCTTGACGCTCAAACCATC-3' and 5'-GGACTAGTACGCGGCAATTTAAACCGCGTC-3' using a full-length hmp clone as a template, which was kindly provided by A.M. Gardner (University of Cincinnati). The PCR product was subcloned into pGEMT-Easy (Promega, Madison, WI), sequenced, and introduced into pTA7001 by the use of 5'-XhoI and 3'-SpeI restriction sites. The construct was transformed into Agrobacterium tumefaciens (strain GV3101) and the latter used for plant transformation of Arabidopsis ecotype Col-0 by the floral dip method (Clough and Bent, 1998). After transformation, seeds were harvested from T0 plants and surface sterilized, and positive transformants were selected on phytagar plates supplemented with Murashige minimal organics medium (Life Technologies, Paisley, UK) containing 15 µg L^–1 hygromycin.

Homozygous T3 plants from single insert lines were used for all experiments, and plants were grown at 22°C under a 9-h-light/15-h-dark cycle. For transgene induction, hmp plants were sprayed with a solution of 3 µM DEX in 0.01% Tween 20. Control experiments were performed with wild-type Col-0 plants treated with 3 µM DEX in 0.01% Tween 20 and/or hmp transgenic plants solely sprayed with 0.01% Tween 20. Pathogen infiltrations followed 16 h after DEX/Tween 20 treatment.

Growth of Plant-Pathogens and Infection

Pseudomonas syringae pv tomato carrying the avirulence gene avrB were transformed with a pRK415 broad host vector (Keen et al., 1988) carrying the complete coding sequence of hmpX (Favey et al., 1995) under control of the Lac promoter, which is constitutively active in Pseudomonas. Detailed description of the construction of pRK415-hmpX and maintenance in Pseudomonas is not provided (M. Boccara, C. Mills, J. Zeier, C. Anzi, C. Lamb, R. Poole, and M. Delledonne, unpublished data).

Pst strains were grown overnight at 28°C in King's B medium containing the appropriate antibiotics (concentrations: rifampicin 50 µg L^–1, kanamycin 50 µg L^–1, tetracycline 15 µg L^–1). Bacteria were pelleted, washed three times with 10 mM MgCl₂, resuspended, and diluted in 10 mM MgCl₂ to the desired concentration (generally 2 x 10⁶ cfu mL^–1, for symptom development 5 x 10⁶ cfu mL^–1, for bacterial growth 10⁶ cfu mL^–1). The bacterial solutions were infiltrated from the abaxial side into one-half of a sample leaf using a 1-mL syringe without a needle. Control (mock) inoculations were performed with 10 mM MgCl₂. Macroscopic symptoms were documented 2 d after infection. Bacterial growth was assessed by homogenizing discs originating from infiltrated areas of three different leaves in 1 mL of 10 mM MgCl₂, plating appropriate dilutions on King's B medium containing Rifampicin, and quantifying colony numbers after 2 to 3 d.

UV Treatment of Arabidopsis Plants

Five-week-old Arabidopsis wild-type and hmp8 plants were pretreated with DEX for 16 h and placed into a growth chamber equipped with UV-A light-emitting black light tubes (Phillips TL 8 W/08; Eindhoven, The Netherlands).

Histochemical Staining and Quantification of H₂O₂ Levels and Microscopic HR Lesions

DAB and Trypan blue staining were performed as described by Thordal-Christensen et al. (1997) and Koch and Slusarenko (1990), respectively. Exogenous H₂O₂ was generated by infiltrating 10 µL of 2.5 mM D-Glc and 2.5 units mL^–1 Aspergillus niger Glc oxidase (Calbiochem, San Diego) in 20 mM Na phosphate buffer, pH 6.5, into Arabidopsis leaves.

For quantification of the number of stained pixels inside the infected leaf area, the histogram function of Adobe Photoshop 6.0 (Adobe Systems, Mountain View, CA) was used. Microscopic photographs were reduced to grayscale mode, and all pixels inside the infiltration zone with a gray tone value <125 were quantified. To account for background staining, the corresponding value for an area of equal size inside the noninfected opposite side of the leaf was subtracted from the latter value and the result divided by the total amounts of considered pixels to yield the relative number of stained pixels in percentage.

RNA and Protein Analysis

Total RNA was isolated from Arabidopsis leaves using Trizol reagent (Life Technologies) following the manufacturer's instructions. RNA-blot hybridization (Levine et al., 1994) was performed with probes of the Arabidopsis gst, pal, and PR-1 genes: ATGST (GenBank accession no. U70672), ATPAL1A (X62747), and ATHRPRP1A (M90508). Equal loading was verified by gel staining with ethidium bromide and by hybridization with an rDNA probe.

For protein extraction, three leaves were homogenized with 1 mL of extraction buffer (15 mM HEPES, 40 mM KCl, 5 mM MgCl₂, 1 mM dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, pH 7.6). The mixture was centrifuged for 30 min at 19,000g and 4°C. The supernatant constituted the protein extract. Protein samples were subjected to SDS-PAGE on 10% (w/v) polyacrylamide (Sambrook et al., 1989) and electroblotted to a polyvinylidene difluoride membrane (Hybond-P; Amersham Pharmacia, Little Chalfont, UK). Probing and detection of western blots were performed as described in the ECL western-blotting detection kit (Amersham). The primary antibody raised against E. coli Hmp was kindly provided by Robert Poole (Sheffield, UK) and used at a dilution of 1:3,000. A goat anti-rabbit IgG horseradish peroxidase conjugate (Sigma, St. Louis) was used as the secondary antibody with a dilution of 1:10,000.

Determination of SA Levels

Measurements of SA and SAG essentially followed the protocol of Raskin et al. (1989). Briefly, 0.15 g of frozen leave tissue was homogenized in 1 mL of 90% methanol and extracted for 10 min at 40°C. The mixture was centrifuged for 5 min at 14,000g, and the pellet was extracted for another 10 min at 40°C with 100% methanol. Supernatants from both extractions were combined and dried under a gentle stream of N₂ at 40°C. The residue was resuspended in 1.5 mL of 0.1 M HCl, and 100 ng of o-anisic acid was added as an internal standard. After centrifugation for 10 min at 14,000g, the aqueous solution was extracted three times with 2 mL of cyclopentane/ethylacetate (1:1). The extracts were combined and the solvent removed under N₂ at 40°C. The residue was dissolved in 50 µL of methanol and passed through a solid-phase extraction column filled with ODS-H optimal packing material (Capital HPLC, Broxburn, UK) using 6 mL of methanol. The eluate was dried under N₂ at 40°C, dissolved in 100 µL of methanol, and used for HPLC analysis. For detection of SAG, the aqueous, acidic phase from the first extraction step was heated to 100°C for 30 min to convert the glucoside to free SA. The above protocol was repeated starting with addition of the internal standard.

HPLC analysis was performed using an ODS-H optimal column (10 x 2.1 mm, Capital HPLC) on a Shimadzu (Columbia, MD) LC-5A chromatograph. For separation, a linear gradient from 95% of H₂O/BuOH/HOAc (98.3/1.2/0.5) to 90% acetonitrile/BuOH/HOAc (98.3/1.2/0.5) in 20 min and flow rate of 0.7 mL min^–1 was applied. For detection, a Waters (Milford, MA) 474 scanning fluorescence detector with an excitation wavelength of 300 nm was used. The emission wavelength was switched at 7 min elution time from 365 nm to 405 nm to ensure highest sensitivities for o-anisic and SA, respectively.

NO Degradation Assay

The kinetics of NO degradation were measured electrochemically using an Iso-NO meter (World Precision Instruments, Sarasota, FL). A saturated, 2 mM NO solution was prepared by bubbling 10 mL of NO gas through 5 mL of HEPES buffer (see above). Protein extracts were prepared as described above using six fully grown Arabidopsis leaves and 1 mL of HEPES-extraction buffer (without dithiothreitol). For NO degradation measurements, 1 mL of plant extract was supplemented with 10 µL of 10 mM NADH and the temperature of the solution kept at 24°C in a water bath. An Iso-NO electrode was calibrated according to the manufacturer's instructions and submerged into the protein solution inside a gas-tight vial. Under stirring, 5 µL of NO solution was added, and the time-dependent changes of the NO signal were recorded.

NO Emission Measurements

Rosette leaves were cut from root parts of Arabidopsis plants and immediately floated on deionized water. The leaves from three different plants were placed in a transparent lid container with 2 L of air volume. A constant flow of measuring gas (NO-free air conducted through a custom-made charcoal column) of 1.5 L/min was pulled through the container and subsequently through the chemiluminescence detector (CLD 770 AL ppt; Eco-Physics, Dürnten, Switzerland) by a vacuum pump connected to an ozone destroyer. Light was provided by a 400 W HQi-lamp (Schreder, Winterbach, Germany) above the container. The quantum flux density could be adjusted at 100 µmol m^–2 s^–1 photosynthetic active radiation by a polyester sieve (pore size is 210 µm) on the lid of the container. Air temperature in the container was usually about 23°C in the dark and 23°C to 25°C under light conditions.

NO Detection using DAF2-DA Fluorescence

Arabidopsis leaves were treated with 2 x 10⁶ cfu mL^–1 Pst (avrB) or 10 mM MgCl₂ as described above, and 3 h later, 10 µM DAF2-DA (Sigma) dissolved in 10 mM Tris/KCl, pH 7.2, was infiltrated into the pretreated leaf areas. One hour after DAF2-DA infiltration, leaf areas were analyzed microscopically using a Zeiss Axioskop 2 fluorescence microscope equipped with a confocal laser scanner (LSM 5 PASCAL; Zeiss, Oberkochen, Germany). Leaves were excited with an argon laser (488 nm). DAF2-DA fluorescence was recorded using a channel with a 505- to 530-nm band-pass filter, and autofluorescence of chloroplast was captured with a channel equipped with a 560-nm long-pass filter.

Sequence data from this article have been deposited with the EMBL/GenBank data libraries under accession numbers X58872, X75893, U70672, X62747, and M90508.

	ACKNOWLEDGMENTS

 
We thank Robert Poole (Sheffield, UK) and Anne M. Gardner (Cincinnati) for kindly providing hmp antibodies and the full-length hmp clone, respectively. Werner M. Kaiser (Würzburg, Germany) is gratefully acknowledged for providing the opportunity to perform NO-emission experiments. J.Z. is a Fellow of the Alexander-von-Humboldt Foundation. The corresponding fellowship supervision by Robert Chow (Edinburgh, UK) is also gratefully acknowledged.

Received March 11, 2004; returned for revision May 26, 2004; accepted June 21, 2004.

	FOOTNOTES

 
^[w] The online version of this article contains Web-only data.

Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.104.042499.

^* Corresponding author; e-mail zeier{at}botanik.uni-wuerzburg.de; fax 49 (0)931 8886235.

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