Fast, transient and specific intracellular ROS changes in living root hair cells responding to Nod factors (NFs)

Authors

  • Luis Cárdenas,

    Corresponding authorSearch for more papers by this author
  • Adán Martínez,

    1. Departamento de Biología Molecular de Plantas, Instituto de Biotecnología, Universidad Nacional Autónoma de México, UNAM, Apartado Postal 510-3, Cuernavaca, Morelos 62271, México
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  • Federico Sánchez,

    1. Departamento de Biología Molecular de Plantas, Instituto de Biotecnología, Universidad Nacional Autónoma de México, UNAM, Apartado Postal 510-3, Cuernavaca, Morelos 62271, México
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  • Carmen Quinto

    1. Departamento de Biología Molecular de Plantas, Instituto de Biotecnología, Universidad Nacional Autónoma de México, UNAM, Apartado Postal 510-3, Cuernavaca, Morelos 62271, México
    Search for more papers by this author

(fax 52 + 777 3136600; e-mail luisc@ibt.unam.mx).

Summary

The role of reactive oxygen species (ROS) in root-nodule development and metabolism has been extensively studied. However, there is limited evidence showing ROS changes during the earliest stages of the interaction between legumes and rhizobia. Herein, using ratio-imaging analysis, increasing and transient ROS levels were detected at the tips of actively growing root hair cells within seconds after addition of Nod factors (NFs). This transient response (which lasted up to 3 min) was Nod-factor-specific, as chitin oligomers (pentamers) failed to induce a similar response. When chitosan, a fungal elicitor, or ATP was used instead, a sustained increasing signal was observed. As ROS levels are transiently elevated after the perception of NFs, we propose that this ROS response is characteristic of the symbiotic interaction. Furthermore, we discuss the remarkable spatial and temporal coincidences between ROS and transiently increased calcium levels observed in root hair cells immediately after the detection of NFs.

Introduction

Legumes can acquire nitrogen through a symbiotic interaction with rhizobial bacteria. This process involves a molecular dialogue between the two partners, in which legume roots exude flavonoids that induce the expression of bacterial nodulation genes encoding proteins involved in the synthesis and secretion of specific lipochitooligosaccharides: the so-called Nod factors (NFs). These NFs signal back to the plant root, and trigger several responses such as ion changes (K+, Cl, Ca2+and H+), cytoplasmic alkalinization, calcium oscillations and gene expression that lead to bacterial invasion and nodule formation (Cárdenas et al., 2000; Oldroyd and Downie, 2004).

In plants, reactive oxygen species (ROS) modulate numerous biological processes such as growth, cell cycle, programmed cell death, plant defense, hormone signaling, biotic and abiotic stress responses and development. ROS have also been found to play a key role in regulating polar growth in root hair cells, fucus zygotes and pollen tubes through their ability to regulate calcium channels, which are involved in maintaining the apical calcium gradient (Coelho et al., 2008; Foreman et al., 2003; Jones et al., 2007; Potocky et al., 2007). On the other hand, ATP has also emerged as an important modulator of NADPH oxidases in plant cells (Kim et al., 2006; Song et al., 2006). In fact, extracellular ATP has been found to modulate intracellular calcium and the levels of ROS in root hair cells (Jeter et al., 2004; Kim et al., 2006; Song et al., 2006).

Whereas compelling evidence indicates that NFs are signal molecules secreted by bacteria that induce various plant cellular responses, it remains unknown whether such rhizobial signal molecules may also elicit pathogen-like responses. It has been proposed that plant pathogenesis and symbiosis are variations on a common theme (Baron and Zambryski, 1995). In this regard, previous results indicate that rhizobia might be initially recognized as intruders that somehow evade or overcome the plant defense response. Indeed, several plant defense responses are induced during root nodule ontogeny (Gamas et al., 1998; Parniske et al., 1990; Vasse et al., 1993). In alfalfa, not all infection threads culminate in successful infection; many abort even compatible symbiotic interactions because of the hypersensitive response triggered in the root cortex (Vasse et al., 1993). In peas, the plant defense response has been proven to be far more critical for arresting the advancement of rhizobial infection during incompatible interactions (Perotto et al., 1994). Wisniewski et al. (2000) proposed that diamine oxidase (DAO) could be a potential source for hydrogen peroxide (H2O2), which might be used by cell wall peroxidase (POD) for hardening the glycoprotein matrix (MGP). Together, these data indicate the importance of ROS metabolism during symbiotic interactions (Wisniewski et al., 2000).

Additional evidence for the role of ROS signaling during rhizobia–legume interactions has been described in Medicago truncatula, where a peroxidase gene (rip1) is induced in the presence of NFs from Sinorhizobium meliloti (Cook et al., 1995). This peroxidase has a cis domain that is ROS sensitive (Ramu et al., 2002). It is interesting that rip1 transcription and ROS production co-localize in the same root cells (Ramu et al., 2002). In fact, exogenous H2O2 can induce rip1 expression, suggesting that cis elements (OCS and OBP) in the rip1 gene are required for induction after ROS production. Furthermore, it has been demonstrated that ROS and ethylene are part of the NFs-induced signal cascade involved in nodule development in Sesbania rostrata (D’Haeze et al., 2003).

During the early stages of M. truncatulaS. meliloti symbiosis, the oxidation of nitroblue tetrazolium (NBT) was detected within the infection threads, indicating that O2 is produced during this process (Santos et al., 2001). Furthermore, it is well known that there is an accumulation of H2O2 during the infection process and during nodule senescence (Rubio et al., 2004). Accordingly, it is plausible that rhizobia should have a mechanism to respond against ROS accumulation in plants. In this regard, bacterial ROS scavenging enzymes such as superoxide dismutase, glutathione-S-transferase and catalase seem to play a protective role in the legume–rhizobia interaction (Jamet et al., 2003; Ramu et al., 2002; Santos et al., 2000; Sigaud et al., 1999). The increased activity of these ROS scavenging enzymes in S. meliloti appears to be essential for the regulation of intracellular ROS production during the establishment and maintenance of symbiosis with the plant host (Jamet et al., 2003).

Paradoxically, decreased ROS production in response to treatment with specific NFs has been also reported in M. truncatula (Lohar et al., 2007; Shaw and Long, 2003b). However, these results were obtained several minutes after the addition of NF, and ROS levels were measured not from a single cell, but from whole sectioned roots. These apparently contradictory results prompted us to analyze ROS responses at the subcellular level using single living root hair cells from the responsive root region. In addition, improved cell imaging methodology was used to measure ROS levels within seconds after the addition of NFs. Herein, we used a semi-quantitative ratio-imaging approach to show that intracellular ROS levels transiently increased in single living root hair cells within a few seconds after treatment with NFs. This response was specific for NFs, and was different from that induced by chitosan, a fungal elicitor, ATP or chitin pentamers that constitute the non-active backbone of the NFs.

Results

Growing root hairs exhibit a tip-localized ROS distribution

Single growing root hair cells were loaded with different concentrations of an ROS-sensitive fluorescent dye (CM-H2DCFDA) in order to achieve optimal loading conditions that did not alter growth and root hair morphology. Root hair cells imaged under these conditions presented a tip-localized ROS signal. Figure 1 (inset) shows the co-localization of differential interference contrast (DIC), and the fluorescent image indicates where the ROS distribution extends to in the subapical region. The location of this signal is similar to that previously described in root hairs from Arabidopsis thaliana (Foreman et al., 2003), and hence we considered this to be a typical ROS distribution for an actively growing root hair cell. Furthermore, time-lapse imaging microscopy demonstrated that ROS levels in growing root hair cells remained unaltered in time, with only discrete fluctuations (Figure 1). In contrast, when root hair cells entered the mature stage, the ROS levels were notably decreased and were completely absent in non-growing cells (data not shown).

Figure 1.

 Intracellular reactive oxygen species (ROS) levels observed in growing Phaseolus vulgaris root hairs.
Inset: an overlay of a DIC and the corresponding fluorescent image from a root hair cell; the tip-localized intracellular ROS distribution was revealed using an ROS-sensitive dye (CM-H2DCFDA). The drawing indicates the region of interest (ROI) where the measurement was performed. Red and blue colors indicate high and low intensities, respectively. Scale bar: 15 μm. The graph illustrates the relative ROS values pooled from 15 different individual root hair cells, and shows that the fluorescent signal remained invariable over time.

NFs induce a rapid and specific transient increase in intracellular ROS levels

Actively growing Phaseolus vulgaris root hair cells from root zone II (Heidstra et al., 1994) were analyzed after loading the roots with the ROS-sensitive dye, under conditions previously described. After root hair cells showed a typical ROS distribution (Figure 2a, left-hand image), they were treated with Rhizobium etli NFs (10−9 m). Rapid intracellular ROS elevation at the apical region of the cell was observed; ROS levels started to increase 15 s after the addition of NFs to the growth medium, and reached a maximum level after 1 min (Figure 2a, middle image; Figure S1). Thereafter, the signal decreased to basal levels (Figure 2a, right-hand image). Similar results were obtained when 15 independent root hair cells were analyzed (Figure 2b). ROS elevation occurred transiently within the first 3 min, and subsequently declined to basal intracellular levels that were finally attained approximately 4 min after treatment with NFs (Figure 2b). The ROS signal decreased to below the basal level by 10 min after addition; this coincided with the arrest of root hair growth, and with the typical swelling response observed at the apical dome (data not shown). ROS signals could not be detected in young (emerging) root hair cells from root zone I (data not shown), because this root region has particularly elevated ROS levels (Shin et al., 2005) that mask single-cell measurements.

Figure 2.

 Intracellular reactive oxygen species (ROS) changes in living Phaseolus vulgaris root hairs after treatment with Nod factors (NFs).
(a) The left-hand image shows a growing root hair cell loaded with CM-H2DCFDA dye and presenting a tip-localized ROS signal (t = 0). The middle image depicts the same root hair cell treated with Rhizobium etli NFs (t = 1 min): the intracellular ROS levels rapidly increased within the first 10–15 s, and peaked after 1 min; thereafter, basal ROS levels were achieved after 3 min, as observed in the right-hand image. Red and blue colors indicate high and low ROS concentration levels, respectively. Scale bar: 15 μm.
(b) Time course analysis of ROS production in root hair cells treated with growth medium containing NFs (10−9 m) or chitin oligomers (10−9 m); a control with only growth medium was also included. Images were analyzed for relative intensity values at the tip of each root hair cell. Pooled data from 15 different cells are shown.

The observed transient ROS responses were specifically induced by NFs, as the addition of growth medium alone or growth medium containing chitin oligomers (pentamers) did not induce any response (Figure 2b). Accordingly, chitin oligomers were used as a negative control for the rest of the study.

H2O2 and UV radiation induced similarly increased ROS levels in root hair cells

In order to test if intracellular ROS levels could be artificially increased in root hairs, exogenous hydrogen peroxide (50 μm) was added to the growth medium. Under these conditions, a sustained increase in intracellular ROS levels was observed after 3 min (Figure 3). Intracellular ROS levels were also artificially increased in root hair cells after a 5-s pulse of UV light (340 nm), as previously reported (Dixit and Cyr, 2003; Mackerness et al., 2001) (Figure 3). This continuously increasing ROS signal was similar to that observed after the addition of H2O2. Apparently, root hair cells could only overcome UV and H2O2 treatment within the first 3 min; after this time there was a dramatic increase in intracellular ROS levels (Figure 3).

Figure 3.

 Reactive oxygen species (ROS) changes in response to hydrogen peroxide (H2O2) and UV radiation.
Root hair cells loaded with CM-H2DCFDA dye and treated with 50 μm exogenously applied H2O2 (black squares) or for 5 s with UV radiation at 340 nm (grey circles) were immediately imaged after treatment. These cells responded with sustained increased ROS levels, and both treatments induced similar responses.

In contrast, the transient ROS response induced in root hairs after treatment with NFs (Figure 2) was different from that observed with exogenously applied H2O2, or after UV irradiation (Figure 3). A sustained and increasing ROS signal that eventually saturated the probe was observed instead (Figure 3), suggesting that these cells were dying. At the cellular level, root hair cells treated in this way became vacuolated, and a swelling response was usually observed (data not shown). Collectively, these data confirm that the conditions and the fluorescent dye concentration used adequately probed the levels of intracellular H2O2 and free radicals in living root hair cells under different conditions.

Chitosan induced sustained increases in intracellular ROS levels

In order to assess whether the ROS levels observed in root hairs after treatment with NFs were specific for the symbiotic interaction, a fungal elicitor was added instead. Chitosan, which is widely used to induce a hypersensitive response in plants, was added to root hair cells that were previously treated with the ROS-sensitive dye (Figure 4, left-hand image), and ROS levels were analyzed under the aforementioned conditions. ROS levels were dramatically increased (Figure 4, middle image) by 5 min after treatment with chitosan, and reached their highest level by 9 min (Figure 4, right-hand image). This sustained and increased response was similar to that induced by UV light radiation and H2O2 treatments (Figure 3). It is worth mentioning that besides causing a dramatic increase in intracellular ROS levels, chitosan also markedly reduced cytoplasmic streaming by 10 min after its addition (data not shown). Intracellular ROS levels in root hair cells were entirely different when comparing transient increases after NF treatment (Figure 4b) with sustained increases after elicitor treatment (Figure 4a,b).

Figure 4.

 Differential intracellular reactive oxygen sepcies (ROS) responses obtained with treatment with chitosan and Nod factors (NFs).
(a) The left-hand image depicts a growing root hair cell loaded with CM-H2DCFDA and showing the typical tip ROS distribution (t = 0). The middle image represents the same root hair cell after treatment with chitosan: the cell presented increased ROS levels. The right-hand image illustrates elevated ROS levels observed after 9 min of chitosan treatment. The red color indicates high levels of ROS. Scale bar: 15 μm.
(b) Graph showing that treatment with chitosan induced a continuous increase in intracellular ROS. The ROS level did not return to the initial values (red squares). Conversely, the addition of NFs induced transient ROS induction (black diamonds). Chitosan or NFs were added at the beginning of the series (t = 0).

Ratio-imaging of intracellular ROS levels reveals a tip-focused signal that responds to treatment with NFs, and is inhibited by diphenylene iodonium (DPI)

The ROS-sensitive dye (CM-H2DCFDA) used in the experiments previously described is a single-wavelength dye that does not allow ratio-imaging. In order to account for the variably accessible cytoplasmic volume of root hair cells, we introduced a reference dye (see Experimental procedures for the pseudo-ratio-imaging analysis. This approach was used to depict, in a semi-quantitative way, the subcellular ROS distribution in growing root hair cells. We found that intracellular ROS levels were indeed focused to the apical dome (Figure 5, inset, left-hand image), and were drastically increased in response to NFs with a transient pattern that peaked after 1 min (Figure 5, inset middle image; Figure S2). This response resembled that obtained previously with the non-ratio-imaging approach (Figure 2, middle image), and was very consistent in 10 of 12 cells; moreover, this ratio analysis allowed the visualization of the subcellular distribution before and after treatment with NFs. It is interesting that at the end of the transient response, the intracellular ROS levels remained higher than the initial values observed before treatment with NFs (Figure 5).

Figure 5.

 Root hairs pre-treated with diphenylene iodonium (DPI) did not display the transient intracellular reactive oxygen species (ROS) response.
Root hair cells were treated with a reference dye (Cell Tracker Red) and ROS-sensitive dye (CM-H2DCFDA) for ratio-imaging of the spatial distribution of ROS. Root hair cells that were preliminarily treated with DPI and then challenged with Nod factors (NFs) did not show the transient ROS response (gray trace). Root hair cells under the same conditions without DPI showed the typical transient response after treatment with NFs. This transient response was observed in 10 of 12 cells. The inset shows the typical subcellular distribution before treatment with NFs (left image), at the peak of the response (middle image) and after the transient response (right-hand image). Note that the intracellular ROS levels remained slightly elevated at the end of the experiment. Scale bar: 15 μm.

DPI is an inhibitor of NADPH oxidases and flavin-containing enzymes, and has been widely used to reduce intracellular levels of ROS in plant cells (Foreman et al., 2003; Lohar et al., 2007; Shaw and Long, 2003b). Growing root hair cells have constant intracellular levels of ROS that did not change when chitin pentamers were used as a control (Figure 6). However, with DPI treatment, the intracellular levels of ROS were drastically reduced after 4 min, although they did not completely disappear, and remained at the basal level after 10 min of treatment with DPI (Figure 6). When root hairs showed basal levels of ROS (after 10 min of treatment with DPI) they were challenged with NFs, but specific transient changes in ROS were not observed (Figure 5). The absence of transient ROS induction after treatment with NFs in root hairs previously treated with DPI suggests that the activation of NADPH oxidases is closely linked to the perception and signaling of NFs.

Figure 6.

 Diphenylene iodonium (DPI) gradually reduced the levels of intracellular reactive oxygen species (ROS).
Root hair cells previously treated with DPI were closely monitored for production of intracellular ROS after 4 min (we did not observe a change prior to that time); as depicted, DPI treatment resulted in gradually decreased levels of cytoplasmic ROS after 4 min, but ROS production was not abolished the (black circles). Root hairs treated with pentamers as a control did not show any response (gray diamonds). Inset, a DIC image of a living root hair cell (left-hand image) treated with Mitotracker Green to follow the mitochondrial distribution (right-hand image). Note that the apical region is enriched with mitochondria. Scale bar: 15 μm.

Root hair cells treated with DPI showed arrested growth and, eventually, root hair swelling (data not shown). As organelles such as chloroplasts and mitochondria with highly oxidizing metabolic activity or high electron flow are also important sources of ROS production in plant cells, we explored the subcellular distribution of mitochondria in growing root hair cells using a mitochondria-specific fluorescent probe (Mitotracker Green). Mitochondria were distributed throughout the cell, but were more abundantly localized at the apical region of the root hair cells (Figure 6, see inset), which coincides with the region in which the ROS signal was observed. Thus, we propose that these mitochondria could partially contribute to ROS production in this region, including basal production observed after DPI treatment.

Extracellular ATP modulates intracellular levels of ROS

Extracellular ATP has been described as inducing changes in intracellular levels of ROS (Song et al., 2006); however, the subcellular distribution of these ROS have not been described. Root hair cells previously loaded with ROS-sensitive and reference dyes were challenged with 10 mm ATP. Under these conditions a continuous increase of apical ROS levels was observed (Figure 7), which was eventually accompanied by a higher cytoplasmic accumulation at the tip region that sometimes resulted in cell bursting (data not shown). Furthermore, repeated local delivery of 5 mm ATP in the vicinity of the root hair cell using a blunt needle resulted in transient increases in the intracellular levels of ROS (Figure 7).

Figure 7.

 ATP induced an increase in the levels of intracellular reactive oxygen species (ROS).
Root hair cells loaded with the ROS-sensitive dye and the reference dye were exposed to 10 mm ATP; intracellular ROS increased constantly until reaching a constant value (black circles). Root hair cells under the same conditions, but exposed to rapid 5 mm ATP perfusion (3 s) showed transiently increased ROS levels (gray diamonds). This response was reproduced when cells were perfused again with 5 mm ATP. Arrows indicate when ATP was added.

Discussion

In the present study, the production and distribution of intracellular ROS, as well as the specific response to NFs, were analyzed in growing P. vulgaris root hair cells. Interestingly, ROS levels were dramatically and transiently increased within a few seconds after treatment with NFs. The response was specific for NFs, and was clearly different to that observed after the addition of H2O2, ATP or chitosan, or after exposure to UV radiation.

As ROS can be used as intracellular signals, all organisms have developed several ways to counteract their deleterious effects (Mittler, 2002; Neill et al., 2002; Vieira Dos Santos and Rey, 2006). The ability of cells to suppress ROS toxicity, and to decode the strength and amplitude of the signal, requires many genes involved in regulating the ROS homeostatic response (Mittler et al., 2004). The functions of ROS signaling in eukaryotic cells include the regulation of cell migration, growth modulation, opening and closing of ion channels, gene expression, development and programmed cell death (Foreman et al., 2003; Mittler et al., 2004; Ushio-Fukai, 2006). As ROS easily diffuse across the plasma membrane, and have a short half-life, rapid localization at the subcellular level is crucial for understanding the nature of the signaling events after a given stimulus. As the ROS-sensitive dye (CM-H2DCFDA) is a non-ratiometric and single-wavelength dye, it was necessary to use an additional dye as a reference for ratio-imaging. Using this approach, we were able to register transient ROS changes induced by NFs in a semi-quantitative way. In this work, we showed that normal root hair cells from P. vulgaris present a tip-localized ROS signal when analyzed with a pseudo-ratio-imaging approach. These ROS changes transiently occurred within a few seconds, and lasted up to 3 min, on average, after root hairs were challenged with NFs. In addition, we found that this response was localized to the tip of the growing cell. This apical response suggests an important role for ROS during the perception of NFs, probably in regulating polar growth, as reported in Arabidopsis, pollen tubes and focus zygotes (Cárdenas et al., 2006; Coelho et al., 2008; Foreman et al., 2003; Potocky et al., 2007; Takeda et al., 2008). Transient ROS changes might participate in decoding the rhizobial signal as a part of symbiosis. Although this idea has been previously proposed (Ramu et al., 2002), responses at the subcellular level have not been observed. By using a pseudo-ratiometric approach, it was possible to subcellularly visualize transiently increased ROS levels in root hair cells responding to NFs. Interestingly, intracellular ROS levels remained higher than the initial values. This suggests that increased ROS levels could be necessary for inducing expression of some early nodulins, as previously suggested (Ramu et al., 2002). Early nodulin expression could be involved in morphological changes that anticipate infection thread formation, or in the regulation of infection thread number, thereby allowing bacteria to enter root hair cells when they are able to circumvent the high basal levels of ROS.

Multiple sources for ROS production inside cells (Ushio-Fukai, 2006) have been described. In particular, it is known that mitochondria and chloroplasts are major sources for generation of ROS as a result of intense membrane electron flow (Mittler et al., 2004). Plant NADPH oxidases have been shown to be the main source of ROS during pathogen attack and UV irradiation (Rao et al., 1996; Sagi and Fluhr, 2001, 2006; Torres et al., 2005). In this work, we showed that root hairs from P. vulgaris showed decreased intracellular ROS levels when previously treated with DPI (Figure 6, black trace). However, a basal level is maintained, indicating that other sources such as plant cell wall diamine oxidase and mitochondria could engage in cytoplasmic production of ROS (Wisniewski et al., 2000). In fact, we observed that mitochondria were located more abundantly at the tip region of the root hair (Figure 6, inset), and might potentially contribute to generation of ROS in the apical zone. Further analysis revealed that root hair cells previously treated with DPI and subsequently challenged with NFs failed to respond with transient increases in intracellular levels of ROS (this work). This result suggests that NADPH oxidases are most likely to be the main source of intracellular ROS. The evidence above, in addition to previous reports (D’Haeze et al., 2003; Ramu et al., 2002), suggest that ROS production is intimately related with the NFs perception pathway. Although it has been shown recently that phosphatidyl inositol 3-Kinase (PI3K) and phosphatidyl inositol-specific phospholipase C (PI-PLC) inhibitors suppress ROS production, a clear analysis of subcellular ROS distribution has not yet been provided (Peleg-Grossman et al., 2007). Furthermore, it is not known how early ROS homeostasis is executed at the subcellular level, or what the differences are between symbiotic and pathogenic responses (Levine et al., 1994). As the ROS response was transiently observed after treatment with NFs, root hair cells must not only have a mechanism to increase the production of ROS, but must also have a mechanism to efficiently remove or neutralize them. ROS homeostasis can be achieved by scavenging intracellular ROS or inhibiting their production. The present finding of increased ROS production only seconds after exposure to NFs seems to contradict the findings of Lohar et al. (2007) recently reported in M. truncatula, at 1 h after treatment with NFs. That study found decreased expression of NADPH oxidase genes, which could account for the reduced levels of ROS observed after 1 h (Lohar et al., 2007; Shaw and Long, 2003b). However, it should be noted that the previous experiments were started several minutes after exposure to NFs, and did not explore the immediate response. This consideration leads us to suggest that NFs may both positively and negatively modulate the intracellular levels of ROS. According to our results, NFs might rapidly stimulate NADPH oxidase either directly or indirectly in the early stages. This could be followed later by inhibition, as demonstrated by Lohar et al. (2007). It has been reported that DPI reduces intracellular levels of ROS, and mimics the swelling response induced by NFs (Lohar et al., 2007). It is possible that decreased ROS levels are required later on during the swelling response, but that higher concentrations might be required during the very early stages of the symbiotic process.

It has been reported that an Arabidopsis NADPH oxidase (RBOH) has EF-hand motifs that are involved in calcium binding (Keller et al., 1998; Sagi and Fluhr, 2001), and that ROS can activate calcium channels (Foreman et al., 2003; Mori and Schroeder, 2004). Furthermore, it has been reported that the oscillating growth pattern of root hair cells is coupled to extracellular pH and changes in ROS (Monshausen et al., 2007): this suggests that ROS could be involved in cell wall property changes that allow polarized growth. The observed ROS gradient (this work) in the apical region spatially correlates with the recently described localization of NADPH oxidase at the apical plasma membrane of root hair cells (Takeda et al., 2008). These data support the idea that ROS and calcium are two key interacting signaling elements required to modulate polarized tip growth. It is plausible that the previously reported elevated calcium influx observed after treatment with NFs (Cárdenas et al., 1999; Shaw and Long, 2003a) could be involved in the transient increases in ROS observed in the present study. In general, the EF-hand and calmodulin binding domains in some NADPH oxidases are thought to be well suited for this kind of regulation (Banfi et al., 2004; Tirone and Cox, 2007). In fact, some calmodulins have been shown to be differentially expressed in response to NFs (Camas et al., 2002). However, there may be other mechanisms responsible for downregulating their function, as NADPH oxidases, which are upregulated during a pathogen attack, are modulated by different kinases, acting in a sequential or parallel manner (Benschop et al., 2007). It will be interesting to determine how this fine regulation is accomplished, and whether ROS changes activate calcium responses or vice versa. In this scenario, calcium and ROS play synergistic key roles in modulating the early stages of the symbiotic interaction.

That the signal of the ROS-sensitive dye (CM-H2DCFDA) was decreased, even though it is irreversibly photooxidized, might be the result of at least two factors. Firstly, the ROS-generating mechanism is tip localized, which means that it occurs in a very limited region of the root hair cells, and could be downregulated after the transient response to NFs, as suggested above, with the consequence that there is a reduction in the generation of ROS. Secondly, the limited region with increased ROS levels could be rapidly dissipated by the strong cytoplasmic streaming and diffusion.

In animal cells, ATP induces ROS production by activating NADPH oxidase (Dichmann et al., 2000; Pines et al., 2005). A regulatory role for extracellular ATP in modulating cellular ROS levels via activation of NADPH oxidases has been recently suggested to occur in plant cells (Roux and Steinebrunner, 2007), including root hair cells (Kim et al., 2006; Song et al., 2006). Our results add weight to the idea that ATP induces changes in intracellular ROS levels in root hair cells, as previously suggested (Song et al., 2006), and thus also favors the idea that ATP could be involved in polar growth, as it can increase intracellular levels of ROS. Analysis of the spatiotemporal correlation between extracellular ATP, calcium and ROS changes in the early stages of symbiosis, and also during pathogen attack, will help in the understanding of the signal pathways involved in each response.

Based on the findings of the present study, we propose that the early production and distribution of ROS are closely related to ATP and calcium signaling in legume root hair cells. Indeed, increased levels of ROS (this work) and calcium spatially and temporally coincide after treatment of root hair cells with NFs (Cárdenas et al., 1999; Shaw and Long, 2003a).

Pathogenesis and symbiosis have many common features (Baron and Zambryski, 1995). The precise regulation of ROS production seems to play a key role in successful Rhizobium–legume interaction (Perotto et al., 1994; Santos et al., 2001; Vasse et al., 1993). It is possible that by regulating ROS production to occur at the right time and place, rhizobia are allowed to enter the host plant without triggering a hypersensitive response. A failure to control ROS elevation might provoke an infection thread abortion (Vasse et al., 1993). The response of root hair cells to NFs with a transient ROS signature signal different from that observed after treatment with chitosan, UV and H2O2 suggests that these cells can differentiate symbiotic from pathogenic signals within seconds. The inability of intracellular ROS to reach initial basal levels after the transient response suggests that increased ROS levels could play a role in the cellular responses. This idea is supported by experiments involving treatment with DPI that showed an inhibition of root hair curling and infection thread formation (Peleg-Grossman et al., 2007).

Using a semiquantitative ratiometric approach in living root hair cells, we established that specific transient elevations in ROS occur at the tip within seconds of treatment with NFs. This transient ROS signature could be the earliest downstream signal after a symbiotic or pathogenic molecular stimulus is perceived by the plant. It is important to unravel the mechanisms underlying ROS signaling during the early symbiotic interaction, in addition to the intimate connections between calcium signaling, extracellular ATP, cell wall peroxidase activities and potential cytoskeleton rearrangements triggered by transient Ca2+ elevation, and ROS production.

Experimental procedures

Seed germination

Phaseolus vulgaris cv. Negro Jamapa seeds were surface-sterilized with sodium hypochlorite for 5 min, followed by five rinses with sterile water, subsequent treatment with pure ethanol for 1 min and five more rinses with water, as described in Cárdenas et al. (1995). Sterile bean seeds were transferred to special plates with wet paper towels, covered with foil and then transferred to a growth chamber at 27°C. After 48 h, seeds were germinated and ready to use in our experiments.

Mounting the living root hairs

The 2-day-old seedlings were placed in liquid Fahreus medium at pH 6.0. After 8 h root hairs were usually well adapted to the medium. Intact seedlings containing the growing root hairs were mounted in chambers constructed from large Petri dishes. These were perforated in the center, and the hole was covered with a large glass cover slip glued with silicon. Seedlings were visualized under a Diaphot 300 (Nikon, http://www.nikon.com) inverted microscope with an 40×/1 NA water immersion lens (Nikon). The chamber contained a layer of solid Fahreus medium (phytagel 0.8%), and the seedlings were set over this layer. At this point, growth medium was continuously perfused with a peristaltic pump at 0.2 ml min−1 (Bio-Rad, http://www.bio-rad.com). The total volume in the chamber was maintained in 2.5–3 ml, on average, and the root region was covered with cellophane paper. The solid Fahreus medium and the cellophane paper helped to prevent root movement.

Treatment of root hair cells with an ROS-sensitive (CM-H2DCFDA) and reference dye (CMTPX)

In brief, the ROS-sensitive probe CM-H2DCFDA [5-(and-6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate, acetyl ester] (Molecular Probes, http://probes.invitrogen.com) was dissolved in DMSO (Sigma-Aldrich, http://www.sigmaaldrich.com) and centrifuged for 2 min at 8000 g to remove non-dissolved particles. Then, the dye solution was diluted with Fahreus medium at a final concentration of 30–50 μm and then added to the plate, replacing the original medium. After 15 min, the medium was replaced with free dye and measurements were performed. This procedure was carried out carefully to avoid any mechanical stress to cells. When performing ratio-imaging, the reference dye Cell tracker™ red CMTPX (Invitrogen, http://www.invitrogen.com) was dissolved in DMSO as a stock solution of 30 μm. Special plates containing the seedlings with the growing root hair cells were treated with the CMTPX dye for 2–3 h. After this time, the dyes were clearly located in the cytoplasm, and were excluded from the vacuoles (Figure S3). Then, the ROS-sensitive dye was added and observed 60 min later under the fluorescence microscope.

Treatment of root hair cells with DPI, H2O2, chitosan, UV and ATP

Root hairs were treated with DPI (Sigma-Aldrich), which is an inhibitor of NADPH oxidase and other flavin-containing enzymes. DPI was dissolved in DMSO and used at 40 μm. H2O2 (Sigma-Aldrich) was freshly prepared each time and used at a concentration of 50 μm. Chitosan (Sigma-Aldrich) was prepared according to the method of Hadwiger and Beckman (1980), dissolved in water and then cleaved by exposure to glacial acetic acid for more than 12 h. Then, the fragments were neutralized by adjusting the pH to 6.0 (Hadwiger and Beckman, 1980). The resulting mixture was used at a final concentration of 100 mg L−1 by diluting the stock solution with Fahreus medium. Treatment with UV was automatically performed as follows: the UV light source was set to open for 5 s, and then the shutter was closed so that we could open the path for excitation at 485 nm, take an image and visualize the intracellular ROS level at each point. ATP was prepared as a 0.1 m stock solution in Tris buffer at 180 nm as follows: 45 μl of 1 m Tris base plus 205 μl of dH2O were mixed carefully and then transferred to an Eppendorf tube containing 14 mg of ATP with shaking for 30 s until all of the ATP was dissolved; aliquots were stored at −80°C.

Incubation of root hairs with NFs

Rhizobium etli NFs were purified by HPLC and applied to plant roots as previously described (Cárdenas et al., 1995). Before treatment, NFs were mixed with 0.5 ml of the same medium that the seedlings had been growing in, and were then added to the growing root hairs with a peristaltic pump to replace the NF-free medium. As a negative control, 10−8 m penta-N-acetylchitopentaose (Seikagaku, http://www.seikagaku.co.jp) dissolved in CHAPS {non-denaturing, zwitterionic detergent [3-(3-cholamidopropyl)-dimethylammonio]-1-propane-sulfonate; Sigma-Aldrich} was used.

Image acquisition and processing

All images were acquired with a CCD camera (Sensys; Roper Scientific, http://www.roperscientific.com) attached to a Nikon TE300 inverted microscope with a 40×/1 NA water immersion objective lens. Loaded cells were excited using a xenon illumination source (DG-4; Sutter Instruments, http://www.sutter.com), which contained a 175 W ozone-free xenon lamp (330–700 nm) and a galvanometer for a wavelength switch. The ROS-sensitive dye excitation was 485 nm and the emission was collected at 535 nm (bandpass: 25 nm). The CMTPX dye was excited at 555 nm and the emission was collected at 605 nm (bandpass: 30 nm). The best temporal and spatial resolutions were achieved when cells were exposed for 15–20 ms at a 490-nm excitation wavelength, with a 2 × 2 binning of the CCD camera. Images were acquired each second, which was fast enough to detect immediate changes in ROS levels. These short exposure times were crucial; otherwise photo-oxidation would have contributed a continuous signal increase (Figure S4). All of the filters used were from Chroma Technology (http://www.chroma.com), and emission filters were located in a filter wheel (Sutter Instruments, http://www.sutter.com). The set-up was automatically controlled by MetaMorph/MetaFluor software (Universal Imaging, http://www.moleculardevices.com). Finally, images were prepared for publication using Adobe Photoshop software (http://www.adobe.com).

Acknowledgements

This work was supported by grants from Dirección General de Asuntos del Personal Académico (DGAPA), Universidad Nacional Autónoma de México, Nos IN228903 (LC) and IN204305 (CQ); CONACyT 58323 (LC), 42560-Q (CQ) and 42562-Q (FS). We thank Liliana Martinez, Maria Luisa Barroso, Olivia Santana and Noreide Nava for technical support. We also thank Dr Otto Geiger, Chris Wood and Peter K. Hepler for their critical reading of the manuscript.

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