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Molecular Plant Advance Access originally published online on October 31, 2007
Molecular Plant 2008 1(2):218-228; doi:10.1093/mp/ssm016
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© The Author 2007. Published by Molecular Plant Shanghai Editorial Office in association with Oxford University Press on behalf of CSPP and IPPE, SIBS, CAS.

Nitric Oxide in Plants: Production and Cross-talk with Ca2+ Signaling

Angélique Besson-Barda, Cécile Courtoisa, Adrien Gauthiera, Jennifer Dahana, Grazyna Dobrowolskab, Sylvain Jeandrozc, Alain Pugina and David Wendehennea,1

a Unité Mixte de Recherche INRA 1088/CNRS 5184/Université de Bourgogne, Plante–Microbe-Environnement, 17 rue Sully, BP 86510, 21065 Dijon cedex, France
b Institute of Biochemistry and Biophysics, Polish Academy of Sciences, 02–106 Warsaw, Poland
c Unité Mixte de Recherche UHP INRA 1136, Interactions Arbres/Micro-Organismes, Université Henry Poincaré Nancy I, BP 239, 54506 Vandoeuvre cedex, France

1 To whom correspondence should be addressed. E-mail wendehen{at}dijon.inra.fr, tel. +33-3–80–69–37–22, fax +33-3–80–69–32–26.


   Abstract
TOP
 Abstract
INTRODUCTION
 ORIGINS OF NO IN...
 DISCUSSION
 METHODS
 SUPPLEMENTARY DATA
 
Nitric oxide (NO) is a diatomic gas that performs crucial functions in a wide array of physiological processes in animals. The past several years have revealed much about its roles in plants. It is well established that NO is synthesized from nitrite by nitrate reductase (NR) and via chemical pathways. There is increasing evidence for the occurrence of an alternative pathway in which NO production is catalysed from L-arginine by a so far non-identified enzyme. Contradictory results have been reported regarding the respective involvement of these enzymes in specific physiological conditions. Although much remains to be proved, we assume that these inconsistencies can be accounted for by the limited specificity of the pharmacological agents used to suppress NO synthesis but also by the reduced content of L-arginine as well as the inactivity of nitrate-permeable anion channels in nitrate reductase- and/or nitrate/nitrite-deficient plants. Another unresolved issue concerns the molecular mechanisms underlying NO effects in plants. Here, we provide evidence that the second messenger Ca2+, as well as protein kinases including MAPK and SnRK2, are very plausible mediators of the NO signals. These findings open new perspectives about NO-based signaling in plants.


   INTRODUCTION
TOP
 Abstract
 INTRODUCTION
ORIGINS OF NO IN...
 DISCUSSION
 METHODS
 SUPPLEMENTARY DATA
 
Nitric oxide (NO) is a gaseous lipophilic free radical first identified as a biological signaling molecule by mammalian biologists. The roles of NO as a modulator of neurotransmission, vascular tone and as an effector in immune responses are well recognized (Schmidt and Walter, 1994). By the late 1990s, plant biologists had started to pay attention to NO. It was undoubtedly an era of great enthusiasm for NO as a plant signaling molecule and, today, it is known to play a crucial role in the regulation of physiological processes ranging from development to adaptation to biotic and abiotic stresses (Supplemental Table 1). There are several NO-producing systems in plants—enzymatic or non-enzymatic ones—and current views on NO enzymatic production tend to fall into two main origins: nitrite-dependent sources and L-arginine-dependent sources. Because of problems inherent to the methods and plant materials used in investigating NO production, as well as to the likely limited specificity of pharmacological agents, there is still some controversy about the source of NO in a defined physiological process. In addition, while proteomic and transcriptomic strategies have led to the identification of numerous NO target genes and proteins (Polverari et al., 2003; Lindermayr et al., 2005, 2006; Grün et al., 2006; Belenghi et al., 2007), the molecular mechanisms underlying its effects remain poorly understood. This review focuses on the description of NO sources in the light of recent findings and discusses emerging data supporting a key role for Ca2+ in mediating NO-based signals in plants.


   ORIGINS OF NO IN PLANTS
TOP
 Abstract
 INTRODUCTION
 ORIGINS OF NO IN...
DISCUSSION
 METHODS
 SUPPLEMENTARY DATA
 
Arginine-Dependent NO Synthesis
In mammals, NO is generated by the family of nitric oxide synthase (NOS) enzymes. All NOSs catalyze the NADPH-dependent formation of NO and L-citrulline from L-arginine (Furchgott, 1995; Li and Poulos, 2005). What is the situation in plants? There is no clear homologue of animal NOS in the genome of A. thaliana (The Arabidopsis Genome Initiative, 2000). Nevertheless, several studies provide evidence arguing for the existence of a NOS-like enzyme in plants. First, there have been a number of reports on the presence of plant proteins immuno-reacting with anti-bodies raised against mammalian NOS (Barroso et al., 1999; Ribeiro et al., 1999). However, proteomic identification of the candidate proteins led to the identification of NOS-unrelated proteins, thus proving the poor reliability of such an immunological-based strategy (Butt et al., 2003). Second, NOS functional activities (measured by following the conversion of radioactive L-arginine to radioactive L-citrulline) have been detected in plant tissue extracts and purified organelles, including peroxisomes and mitochondria (Barroso et al., 1999; Guo et al., 2005; Corpas et al., 2006). Accordingly, inhibitors of mammalian NOS have been widely and successfully used to suppress NO production in plants challenged by biotic and abiotic stresses (Delledonne et al., 1998; Durner et al., 1998; Lamotte et al., 2004; Mur et al., 2005; Arnaud et al., 2006; Zhang et al., 2007; Zhao et al., 2007; Zottini et al., 2007). Here, too, although the data are compatible with the occurrence of a NOS-like enzyme in plants, they remain questionable. Indeed, so far, there is no direct experimental evidence that the radioactive products detected when assessing plant NOS activity in vitro is indeed L-citrulline (Crawford et al., 2006). Furthermore, taking it for granted that NOS inhibitors are specific in plants seems incongruous.

On the basis of these findings, an emerging hypothesis is that the plant enzyme displaying NOS-like activity is structurally different from classical mammalian NOS. Such a possibility has been verified by Guo et al. (2003), who reported the identification of AtNOS1—an A. thaliana member of a novel family of putative NOS unrelated to eNOS, nNOS and iNOS. The protein was shown to produce NO from L-arginine in mitochondria and was found to be associated with NO production in A. thaliana plants exposed to abscisic acid (ABA, Guo et al., 2003, 2005), lipopolysaccharides (LPS) and virulent pathogens (Zeidler et al., 2004). Furthermore, NO levels were found to be lower in the Atnos1 mutants impaired in AtNOS1 expression (Guo et al., 2003). However, AtNOS1 ability to catalyze a NOS-like activity was recently questioned (Crawford et al., 2006; Zemojtel et al., 2006). Rather, it has been proposed that AtNOS1 (re-named AtNOA1 for nitric oxide associated 1; Crawford et al., 2006) might act as a GTPase implied in mitochondrial biogenesis (Zemojtel et al., 2006). Because mitochondria are strong sources of NO, this scenario may explain the reduced level of NO in the Atnos1 mutant.

Another important aspect to be considered concerns the involvement of polyamines (PAs) in NO synthesis. Tun et al. (2006) recently reported that adding the polyamines spermidine and spermine to seedlings of Arabidopsis caused rapid production of NO in the elongation zone of the root tip and in primary leaves, especially in the veins and trichomes. Quite obviously, the quickness of the PA-dependent NO production speaks in favour of an enzymatic source. This conversion might be carried out by unknown enzymes or by PA oxidases (Yamasaki and Cohen, 2006). The latter are not known to generate NO in animal systems but some PA oxidases possess an enzymatic mechanism different from that of the otherwise homologous animal enzymes (Binda et al., 2002). Strictly speaking, the finding that PAs cause NO synthesis should be discussed in the light of putative side effects of mammalian NOS inhibitors in plants. Indeed, one would expect that these pharmacological compounds might also affect the activity of arginase and arginine decarboxylase—two key enzymes of the PA biosynthetic pathway using L-arginine as a substrate. In other words, the suppression of NO synthesis by mammalian NOS might be related to an inhibition of PA biosynthesis and subsequently of PA-induced NO production rather than a direct effect on a putative NOS-like enzyme. Future work should investigate this assumption.

Nitrite-Dependent NO Synthesis
Nitrite is emerging as a main substrate for NO synthesis in plants, and enzymatic as well as non-enzymatic routes for nitrite-dependent NO production have been described. Enzymatic routes mainly involved NR—a key enzyme of nitrogen assimilation. NR catalyses the reduction of nitrate to nitrite using NAD(P)H as an electron donor (Lea, 1999) but can also generate NO from nitrite both in vitro (Yamasaki and Sakihama, 2000) and in vivo (Rockel et al., 2002), this second activity representing only a small part (about 1%) of its normal nitrate-reducing capacity (Rockel et al., 2002; Planchet et al., 2005). Biochemical as well as genetic-based approaches using NR-deficient mutants with reduced nitrite content indicate that NR is the main enzymatic NO source in ABA signaling and during hypoxia (Desikan et al., 2002; Dordas et al., 2004; Bright et al., 2006). Besides NR, a tobacco root-specific plasma membrane-bound nitrite:NO reductase (NI-NOR) was shown to catalyze the reduction of apoplastic nitrite into NO (Stöhr et al., 2001). NI-NOR activity might be coordinated with those of a plasma membrane-bound NR (PM-NR) reducing apoplastic nitrate to nitrite. The identity of NI-NOR is currently unknown. Other sources include a yet unidentified enzyme catalyzing a mitochondrial electron transport-dependent reduction of nitrite to NO (Planchet et al., 2005) and an apoplastic non-enzymatic conversion of nitrite to NO occurring at acidic pH in the presence of reductants such as ascorbic acid (Bethke et al., 2004).

Nitrite- or L-arginine-Dependent Pathway?
Contrasting data regarding the enzymatic source of NO produced in particular physiological contexts have been reported. For example, both NR and NOS-like enzymes have been shown to catalyze NO synthesis in A. thaliana guard cells and tobacco cell suspensions exposed to ABA and elicitors of defence responses, respectively (Desikan et al., 2002 versus Guo et al. 2003 and Zhang et al., 2007; Lamotte et al., 2004 versus Planchet et al., 2006). These inconsistencies have been attributed to the limited specificity of mammalian NOS inhibitors but need re-evaluation in the light of new findings.

First, Modolo et al. (2006) recently reported that the commonly used A. thaliana NR-deficient double mutant nia1 nia2, which has been used to assess the involvement of NR in NO synthesis, has much lower L-arginine content in leaves (almost 10 times lower) as compared to wild-type plants. Furthermore, the same authors observed a significant NO production when nia1 nia2 leaves were infiltrated with L-arginine. Taken together, these results indicate that mutants impaired in NR expression lack substrates (that is nitrite and L-arginine) to produce NO from the nitrite- but also the L-arginine-dependent pathways. Therefore, data based on the use of NR- or nitrate/nitrite-deficient plants should be cautiously interpreted.

Second, studies based on NR- or nitrate/nitrite-deficient plants and cell suspensions do not take into account the key role of nitrate-permeable anion channels in signaling processes. This fact is exemplified by the analysis of the NO source in cryptogein signaling. Cryptogein is a 10-kDa proteinaceous elicitor produced by the oomycete Phytophthora cryptogea triggering defence responses in tobacco (Ricci et al., 1989). When applied to tobacco plants and cell suspensions, cryptogein induces within 5 min a Ca2+-dependent NO synthesis sensitive to mammalian NOS inhibitors but insensitive to NR inhibitors (review by Garcia-Brugger et al., 2006; see also Foissner et al., 2000; Lamotte et al., 2004). We previously reported that cryptogein triggers a rapid and massive efflux of nitrate, leading to a loss of intracellular nitrate content of about 60% within 1 h (Wendehenne et al., 2002; Gauthier et al., 2007). Interestingly, the nitrate efflux was shown to be a necessary step for the mediation of cryptogein-induced early and late events, including Ca2+ influx, increases in [Ca2+]cyt and subsequent Ca2+-dependent events, namely plasma membrane depolarization, increase in ROS production, gene expression and cell death (Wendehenne et al., 2002; Gauthier et al., 2007). In a recent paper, Planchet et al. (2006) reported that NO production by cryptogein was completely absent in ammonium-grown tobacco cell suspensions totally devoid of nitrate, indicating that NO synthesis was catalyzed through a nitrate/nitrite-dependent pathway rather than an L-arginine-dependent mechanism, as previously inferred from pharmacological studies (Foissner et al., 2000; Lamotte et al., 2004). To complete this investigation, in the present study, we analyzed other cryptogein-induced events in ammonium-grown cells deprived of nitrate versus nitrate-grown cells (Figure 1). As expected, and in accordance with Planchet et al. (2006), the elicitor-mediated nitrate efflux (Figure 1A) and NO production (Figure 1B) were strongly reduced in ammonium-grown cells compared with nitrate-grown cells. However, both ROS production and rises in [Ca2+]cyt were also almost completely inhibited in ammonium-grown cells (Figure 1C and 1D). In contrast, the activity of the Mitogen-Activated Protein Kinase (MAPK) SIPK (Salicylic acid-Induced Protein Kinase), which was previously shown to be induced independently of the cryptogein-induced nitrate efflux in nitrate-grown cells, was still induced in ammonium-grown cells exposed to the elicitor (Supplemental Figure 1). Based on these observations and the key role of nitrate-permeable anion channels in cryptogein signaling, it is conceivable that all the signaling events acting downstream of nitrate efflux are strongly impaired in nitrate-deprived cells. Therefore, these data raise the question of the importance of NR as a source of NO synthesis in cryptogein signaling: is the absence of NO production in nitrate-deprived cells related to the absence of NR substrates (nitrate/nitrite), or is it associated with the absence of a nitrate efflux and the related changes in [Ca2+]cyt required for NOS-like mediated NO synthesis? The question is still opened and deserves further investigation.


Figure 1
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  		Figure 1. Cryptogein-Induced Early Events in Ammonium-Grown Tobacco Cell Suspensions.

(A) Time course of nitrate efflux induced by cryptogein (25 nM) in ammonium-grown Nicotiana tabacum cv Xanthi cells deprived of nitrate (cry NH4+) and in nitrate-grown Nicotiana tabacum cv Xanthi cells (cry NO3–). Nitrate content was determined as described by Wendehenne et al. (2002). Each value represents the mean ± SD of nine measurements (three replicates per experiment performed three times).

(B) NO production induced by cryptogein (25 nM) in ammonium-grown Nicotiana tabacum cv Xanthi cells deprived of nitrate (cry NH4+) and in nitrate-grown Nicotiana tabacum cv Xanthi cells (cry NO3–). NO production was measured using the intracellular NO-sensitive fluorophore diaminofluoresceine-diacetate (DAF-2DA), as reported by Lamotte et al. (2004). Each value represents the mean ± SD of nine measurements (three replicates per experiment performed three times).

(C) Cryptogein-induced [Ca2+]cyt increase in ammonium-grown cells deprived of nitrate (cry NH4+) and in nitrate-grown cells (cry NO3–). [Ca2+]cyt measurement was performed using apoaequorin-transformed Nicotiana plumbaginifolia cells expressing aequorin addressed in the cytosol, as described by Lamotte et al. (2006). Controls, omitted for clarity, did not induce changes in [Ca2+]cyt during the time of experiment. Results are from one of three representative experiments.

(D) H2O2 production induced by cryptogein (25 nM) in ammonium-grown Nicotiana tabacum cv Xanthi cells deprived of nitrate (cry NH4+) and in nitrate-grown Nicotiana tabacum cv Xanthi cells (cry NO3–). Production of H2O2 was detected by chemiluminescence, as reported by Lamotte et al. (2004). Each value represents the mean ± SD of nine measurements (three replicates per experiment performed three times).

 
NO-Mediated Cellular Signals: A Ca2+ Story
In animals, numerous studies point to a specific role for NO in controlling Ca2+ homeostasis. Pharmacological, biochemical and electrophysiological approaches have shown that NO modulates the activity of plasma membrane as well as intracellular Ca2+-permeable channels. Strictly, almost all types of Ca2+ channels appear to be regulated by NO (Clementi, 1998). NO impacts on their activity directly through S-nitrosylation—the reversible formation of a covalent bound between a cysteine residue and an NO group—or indirectly (Stamler et al., 2001; Ahern et al., 2002). The indirect means involve cGMP, produced following the NO-induced activation of soluble guanylate cyclase, and/or cyclic ADP-ribose (cADPR)—a Ca2+-mobilizing metabolite that is synthesized from NAD+ by ADP-ribosyl-cyclase (Willmott et al., 1996; Hanafy et al., 2001). Accumulating evidence suggest that cADPR mediates Ca2+ release by activating the intracellular Ca2+ channels ryanodine receptors (RYR) in mammals but also in plants (Allen et al., 1995; Fliegert et al., 2007).

The concept that NO also modulates Ca2+-permeable channels in plant cells was supported by the observation that the NO released by NO donors induced a transient rise in [Ca2+]cyt in Vicia faba guard cells and in tobacco cell suspensions (Garcia-Mata et al., 2003; Gould et al., 2003; Lamotte et al., 2004, 2006). Moreover, NO scavengers and mammalian NOS inhibitors reduced the increase in [Ca2+]cyt triggered by hyperosmotic stress or elicitors of defense responses, including cryptogein and endopolygalacturonase I from Botrytis cinerea (Gould et al., 2003; Lamotte et al. 2004, 2006; Vandelle et al., 2006). Figure 2 gives an example of such experiments (see also Lamotte et al., 2006). Treatment of transgenic Nicotiana plumbaginifolia cells expressing the Ca2+ reporter aequorin addressed in the cytosol with the NO donor diethylamine-NONOate (DEA/NO) was followed by a rapid and transient biphasic elevation in [Ca2+]cyt (Figure 2A). Experiments based on the use of the NO scavenger cPTIO demonstrated that the first increase in [Ca2+]cyt was caused by the DEA/NO solubilization buffer whereas the second elevation was specifically due to NO (Lamotte et al., 2006). Depending on the experiments, the second elevation reaches 350–500 nM (Figure 2A and 2B). The transient aspect of NO-induced [Ca2+]cyt changes suggests that Ca2+-ATPase and other Ca2+ transporters (as well as Ca2+-buffering compounds) are active within the cell membranes after NO treatment. Interestingly, a second treatment with DEA/NO 30 min after a first treatment with the NO donor induced a delayed and reduced [Ca2+]cyt rise compared with the first one, indicating that the same cells are poorly responsive to repeated NO treatments (Figure 2B). It is unlikely that the ineffectiveness of the second NO treatment was caused by a breakdown of ion gradient regulation by NO, since a subsequent exposition to hyper-osmotic stress led to a typical biphasic increase in [Ca2+]cyt (Gould et al., 2003; Lamotte et al., 2006). The inefficiency of the second NO treatment may reflect the inactive state of components of the pathway(s) mediating the NO-dependent [Ca2+]cyt increase. Interestingly, it should also be noted that treatments with DEA/NO did not elicit increases in nuclear free Ca2+ concentration in transgenic tobacco cells expressing aequorin addressed to the nucleus (Lecourieux et al., 2005). This latter finding highlights a specific role of NO in governing the cytosolic Ca2+ homeostasis.


Figure 2
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  		Figure 2. Nitric Oxide Induces an Elevation in [Ca2+]cyt in Nicotiana plumbaginifolia Cells.

(A) Changes in [Ca2+]cyt in apoaequorin-transformed Nicotiana plumbaginifolia cells in response to DEA/NO. 8-br-cADPR (250 µM) was added to cell suspensions 10 min prior to DEA/NO (50 µM). Treatment with the DEA/NO solubilization buffer (control cells), omitted for clarity, induced the first increase of [Ca2+]cyt but not the second one, which was shown to be specific to NO (Lamotte et al., 2006). 8-br-cADPR itself, omitted for clarity, did not induce changes in [Ca2+]cyt during the time of experiment. Results are from one of six representative experiments.

(B) Ineffectiveness of a second DEA/NO application in inducing changes of [Ca2+]cyt in apoaequorin-transformed Nicotiana plumbaginifolia cells. A second DEA/NO (50 µM) treatment was applied 30 min after the first treatment (time 0) with the NO donor (50 µM). Sorbitol (250 mM) was added to the cell suspensions 55 min after the beginning of the experiment. Results are from one of three representative experiments.

 
Inhibitors of plasma membrane and intracellular Ca2+ permeable channels have both been found to inhibit NO-induced increases in [Ca2+]cyt (Garcia-Mata et al., 2003; Gould et al., 2003; Lamotte et al., 2004, 2006; Vandelle et al., 2006). Thus, depending on the physiological context, NO might promote an influx of Ca2+ from the extracellular space and/or mobilization of Ca2+ sequestered in intracellular Ca2+ stores. Although the identity of the Ca2+ -permeable channels involved in that process remains unknown, all pharmacological-based studies unanimously point out RYR-like channels as main targets for NO. Analyses of the putative involvement of cADPR in NO signaling have strongly reinforced this assumption. Indeed, NO-induced accumulation of the pathogenesis-related (PR)-1 transcripts in tobacco leaves was suppressed in the presence of the cADPR-selective antagonist 8-bromo-cADPR (Klessig et al., 2000). In agreement with these data, vacuum infiltration of nanomolar concentrations of cADPR in tobacco leaf disks triggered the expression of the PR-1 gene, which was suppressed by RYR inhibitors (Durner et al., 1998). Naturally, these data do not conclusively prove that NO activates RYR-like receptors, but collateral evidence supports that cADPR has a dominant role in generating NO-dependent Ca2+ fluxes: the NO-mediated Ca2+ transient influx is reduced by almost 40% by 8-bromo-cADPR (Figure 2A; see also Lamotte et al., 2006). Finally, it should be mentioned that the possibility that NO might also influence the activity of inositol 1,4,5-triphosphate receptors has been suggested (Vandelle et al., 2006).

Together with cADPR, the involvement of protein kinases in mediating NO-induced changes in [Ca2+]cyt has become an interesting object of study. Inhibitors of protein kinases, such as K252a and staurosporine, indeed reduced the [Ca2+]cyt rises triggered by NO in Vicia faba guard cells and tobacco cell suspensions, indicating that protein kinases might be downstream effectors of NO action on [Ca2+]cyt (Sokolovski et al., 2005; Lamotte et al., 2006). Lamotte et al. (2006) tentatively identified these protein kinases by analyzing the protein kinase activities of protein extracts from N. plumbaginifolia cells exposed to the NO donor DEA/NO—a treatment leading to an increase in [Ca2+]cyt (see above). A representative in-gel kinase assay is shown Figure 3A (see also Lamotte et al., 2006). Treatment with NO resulted in the activation of a 42-kDa protein kinase within 5 min. Its activation was observed with DEA/NO concentrations as low as 50 µM (data not shown; Lamotte et al., 2006). The NO-induced 42-kDa protein kinase was identified as NtOSAK (Nicotiana tabacum Osmotic-Stress-Activated protein Kinase)—a member of the plant SNF (Sucrose Non Fermenting) 1-related protein kinase type 2 (SnRK2) family (Mikolajczyk et al., 2000; Kelner et al., 2004; Lamotte et al., 2006). Originally, NtOSAK was identified as a hyperosmotic and salt stress-activated protein kinase, suggesting that it may play an important role in osmotic-stress signaling (Mikolajczyk et al., 2000). Accordingly, NO was shown to be required for NtOSAK activation in tobacco cell suspensions exposed to hyperosmotic stress (Lamotte et al., 2006).


Figure 3
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  		Figure 3. NO Promotes the Activation of the Protein Kinases NtOSAK and SIPK in Nicotiana plumbaginifolia Cells.

(A) Activation of protein kinases in apoaequorin-transformed Nicotiana plumbaginifolia cells exposed to DEA/NO. Cell suspensions were treated with DEA/NO solubilization buffer (control) or 2 mM DEA/NO, and aliquots of the cultured cells were taken at the indicated times. The NO scavenger cPTIO (500 µM) was added to cell suspensions 10 min prior to DEA/NO, and aliquots were taken 30 min later. Protein kinase activities were analyzed in cell extracts by in-gel kinase assay with HIIIS as a substrate, as described by Lamotte et al. (2006). cPTIO itself did not induce protein kinase activation (data not shown). The 42-kDa protein kinase was identified as NtOSAK; its activation was induced in response to micromolar concentrations of DEA/NO (see the text; Lamotte et al., 2006). The results shown are one of five representative experiments.

(B) Immunological analysis and Ca2+ dependence of the NO-activated 48-kDa protein kinase. Protein extracts from untreated apoaequorin-transformed Nicotiana plumbaginifolia cells or exposed to DEA/NO (2 mM) for 30 min were subjected to a Western blot analysis using polyclonal antibodies that are specific to the MAPK-activated form. Ruthenium red (RR, 5 µM) and La3+ (500 µM) were added to cell suspensions 10 min prior to DEA/NO. When used alone, these compounds did not induce MAPK activation (data not shown). The results shown are one of three representative experiments.

(C) Effects of La3+ on the NO-induced activation of the 48-kDa protein kinase and NtOSAK in apoaequorin-transformed Nicotiana plumbaginifolia cells. La3+ (500 µM) was added to the cell suspensions 10 min prior to DEA/NO (2 mM). The cells were exposed to the donor for 15 min and then analyzed for protein kinase activity by in-gel kinase assay with HIIIS as a substrate. When used alone, La3+ did not induce protein kinase activation (data not shown). The results shown are one of three representative experiments.

(D) Immunoprecipitation of the NO-activated 48-kDa protein kinase with the anti-SIPK antibodies. Protein extracts from untreated apoaequorin-transformed N. plumbaginifolia cells or exposed to DEA/NO (2 mM) for 30 min were immunoprecipitated with anti-SIPK antibodies. The resulting immunocomplexes were analyzed by in-gel kinase assay with HIIIS as a substrate. The results shown are one of three representative experiments.

 
In addition to NtOSAK, NO induced the activation of a second protein kinase with a molecular mass of 48 kDa (Figure 3A). Its activity peaked at 30 min before returning to the basal level and was detected only at high DEA/NO concentrations (that is mM ranges). Because the in-gel kinase assay used in this study allows the detection of the activity of MAPKs, we therefore analyzed whether the NO-induced 48-kDa protein kinase might be a MAPK. For this purpose, we tested the cross-reactivity of polyclonal antibodies raised against a phosphorylated form of human ERK1 (Extracellular signal-Regulated Kinase 1) and ERK2. These antibodies specifically react with the activated form of plant ERK-related MAPKs (Zhang and Liu, 2001). Immunostaining revealed one band that was strongly detected in protein extracted from NO-treated N. plumbaginifolia cells for 30 min (Figure 3B). The reactive band was poorly immunodetected in control cells. This band has a relative molecular mass of 48 kDa, consistent with the in-gel kinase assay (Figure 3A). Thus, the NO-induced 48-kDa protein kinase is likely to be a MAPK. SIPK is a tobacco 48-kDa MAPK that is activated in response to pathogens and osmotic stress (see above; Mikolajczyk et al., 2000; Zhang and Liu, 2001). To verify that the NO-induced 48-kDa MAPK could correspond to SIPK, we immunoprecipitated cell extracts from untreated and NO-treated cells with specific anti-SIPK polyclonal antibodies, and analyzed the resulting immunocomplexes with the in-gel kinase assay technique. As shown in Figure 3D, the NO-induced 48-kDa MAPK was immunoprecipitated by the anti-SIPK antibodies, thus demonstrating that it is SIPK. To investigate whether Ca2+ is required for the 48-kDa protein kinase activation by NO, we tested the effects of inhibitors of plasma membrane Ca2+-permeable channels (lanthanum, La3+) and RYR (ruthenium red). The Ca2+ surrogate La3+, which blocks the NO-induced Ca2+ influx and therefore reduces the subsequent [Ca2+]cyt elevation (Lamotte et al., 2004, 2006), completely suppressed the activation of the 48-kDa kinase by NO, as revealed by Western blotting (Figure 3B) and in-gel kinase assay (Figure 3C). By contrast, the RYR inhibitor did no affect the activation of the 48-kDa kinase (Figure 3B). Thus, the activation of SIPK could be preceded by a rise in [Ca2+]cyt triggered by the NO-dependent activation of plasma membrane Ca2+-permeable channels. The scenario is bound to be different for NtOSAK, whose activation is insensitive to La3+ (Figure 3C) and intracellular Ca2+-permeable channel inhibitors (Lamotte et al., 2006).

The demonstration that artificially generated NO stimulated MAPKs including SIPK was previously reported (Clarke et al., 2000; Kumar and Klessig, 2000; Pagnussat et al., 2004; Zhang et al., 2007). However, the key role of Ca2+ in that process has not been investigated so far. The observation that a convergence of Ca2+- and NO-signaling pathways might occur at the MAPK level was similarly observed in neuronal cells by Lee et al. (2000). It should be noted that NO has been shown to activate SIPK through a salicylic acid (SA)-dependent pathway in tobacco leaves (Kumar and Klessig, 2000). In contrast, we observed that SIPK remained fully active in response to NO when similar experiments were performed in transgenic tobacco cell suspensions unable to accumulate SA (data not shown). This discrepancy might be related to the biological model (e.g. plants versus cell suspensions) and the source of NO (e.g. injection of a recombinant rat neuronal NOS in the extracellular spaces of tobacco leaves versus DEA/NO in the present study).

Finally, although the finding that NO activates NtOSAK and MAPKs enrich our understanding of how NO exerts its effects, their putative involvement in NO-induced [Ca2+]cyt rises needs to be established. We assume that NtOSAK is an interesting candidate; the analysis of its implication in the changes of [Ca2+]cyt triggered by NO is under investigation.

The data discussed above propose that NO triggers cellular events in plant cells by causing an increase in [Ca2+]cyt. This process has been shown to occur in ABA, hyper-osmotic and elicitor transduction pathways (Garcia-Mata et al., 2003; Gould et al., 2003; Lamotte et al., 2004, 2006; Vandelle et al., 2006). Several studies have also dealt with a role for Ca2+ in initiating NO production. As a result, the idea that NO produced through NOS-like enzymes requires a rise in [Ca2+]cyt is increasingly accepted (Delledonne et al., 1998; Lamotte et al., 2004; Corpas et al., 2006; Vandelle et al., 2006). Recent work by Ali et al. (2007) has significantly advanced our knowledge about the nature of the molecular mechanisms underlying this process. These authors report that the Ca2+-dependent NOS-like-mediated NO production occurring in response to LPS in A. thaliana guard cells is brought about by the activation of the plasma membrane cyclic nucleotide gated channel CNGC2. Supporting this conclusion, plants without functional CNGC2 (dnd1: defence no death 1 mutants) lack inward plasma membrane Ca2+ currents and failed to produce NO in response to LPS. Interestingly, the hypersensitive response (HR), which is normally reduced in the dnd1 plants inoculated with avirulent pathogens, was partially restored by the NO donor SNP. These latter findings highlight a role for NO produced through a Ca2+-dependent process in mediating the HR, in agreement with previous pharmacological-based studies (Delledonne et al., 1998; Lamotte et al., 2004).

To sum up, the evidence discussed here documents the complexity of the interactions between NO and the second messenger Ca2+. On the one hand, NO produced through an L-arginine-dependent pathway is strictly Ca2+-dependent. Identification of CNGC2 as a component of this process is of utmost interest. On the other hand, NO is emerging as a very plausible initiator of the Ca2+ signal and might mediate part of its effects through cADPR and protein kinases. Clearly, this picture offers an interesting perspective of the NO-linked messengers that play a role in transducing signals. Naturally, future work will have to clarify the tight interplays between NO, Ca2+, cADPR, and protein kinases but the findings so far are certainly promising.


   DISCUSSION
TOP
 Abstract
 INTRODUCTION
 ORIGINS OF NO IN...
 DISCUSSION
METHODS
 SUPPLEMENTARY DATA
 
The explosive growth in the area of research related to NO over the past few years has been amazing, rapidly extending to all aspects of plant physiology. Notwithstanding this statement, the fact of the matter is that we have not yet been able to identify a plant enzyme displaying a NOS-like activity. This issue, together with the ongoing debate relative to the source of NO in particular physiological context, is a main problem faced by plant biologists working in the field of NO research. Clearly, biochemical purification of the enzyme displaying NOS-like activity is a main priority. The recent finding of PAs as putative substrates for NO synthesis has shed some light on this issue and, although speculative, might explain the effectiveness of mammalian NOS inhibitors in suppressing NO synthesis.

The details about how NO exerts its effects at the molecular level remain unclear. The discovery of the occurrence of tight interplays between NO, Ca2+, cADPR, and protein kinases has provided an important focal point to future research in this area. Perhaps one of the most compelling biological questions is how the NO/Ca2+ pathways will guide the cells toward a specific response. The identification of NO as well as Ca2+ protein targets, for instance S-nitrosylated proteins or new Ca2+-dependent protein kinases, in specific cellular contexts should help to answer this question. With the recent development of proteomic approaches to study S-nitrosylation reactions (Lindermayr et al., 2005, 2006; Belenghi et al., 2007), we can confidently hope that part of these perspectives are now at hand.


   METHODS
TOP
 Abstract
 INTRODUCTION
 ORIGINS OF NO IN...
 DISCUSSION
 METHODS
SUPPLEMENTARY DATA
 
Cell Culture
Nicotiana tabacum cv Xanthi cell suspensions were cultivated as previously described (Lamotte et al., 2004). Briefly, cell suspensions were maintained in Chandler's medium on a rotary shaker (150 rpm, 25°C) under continuous light (photon flux rate 30–40 µmol m–2 s–1). Cells were maintained in the exponential phase and subcultured 1 d prior to utilization. Transgenic Nicotiana plumbaginifolia cell suspensions expressing apoaequorin were cultivated in the dark on a rotary shaker (150 rpm, 25°C), as described by Lamotte et al. (2006). For experiments, 8 d old transgenic tobacco cell suspensions were used.

Ammonium-grown cell suspension cultures totally devoid of nitrate were grown on Chandler's medium with small modifications as described by Planchet et al. (2006): 1.5 mM NH4Cl instead of NH4NO3 and KNO3. Cells were sub-cultured three times (once every 3 d) in a fresh Chandler's medium devoid of nitrate before experiments.

Cell Treatments
Cells were washed by filtration and resuspended at 0.1 g fresh weight mL–1 in the H10 suspension buffer (175 mM mannitol; 0.5 mM CaCl2 ; 0.5 mM K2SO4 ; 10 mM HEPES ; pH 5.75), and equilibrated in the same buffer for 2 h at 24°C on a rotary shaker (150 rpm) before treatments. Typically, 6 x 105 cells/0.1 g fresh weight occur in cell suspensions.

All chemicals were purchased from Sigma-Aldrich (Saint-Quentin-Fallavier, France) except carboxy-PTIO (cPTIO) and coelenterazine, from Calbiochem (Schwalback, Germany), and DEA/NO, from Cayman chemical (Ann Arbor, USA). The chemicals were dissolved in water.

DEA/NO was dissolved in NaOH 0.01 M as a stock solution stored on ice and daily prepared. To initiate the release of NO, an aliquot of the stock alkaline solution of DEA/NONOate (DEA/NO) was dissolved in phosphate buffer 100 mM pH 7.2 maintained at room temperature to give a 2 or 100-mM final concentration, depending on the experiment. Cells were treated with DEA/NO (50 µM or 2 mM final concentration) 20 s after dissolving the stock alkaline solution of DEA/NO in the phosphate buffer. As control, cells were treated with a same volume of DEA/NO solubilization buffer (NaOH 40 µM in phosphate buffer 100 mM pH 7.2) without DEA/NO. For each assay, a fresh phosphate buffer solution of DEA/NO was prepared from the stock alkaline solution of the donor.

Cryptogein was purified according to Gauthier et al. (2007) and prepared as a 10-µM stock solution in water.

Extracellular Nitrate Content Analysis
At the indicated time of cryptogein treatment, aliquots of 2 mL of cells (0.2 g fresh weight) were filtrated and the resulting extracellular medium was collected. The NO3– concentration in the extracellular medium was determined using a NO3– colorimetric assay kit (Alexis Biochemicals), as described by Wendehenne et al. (2002).

NO Production
NO accumulation was determined using the fluorophore 4,5-diaminofluorescein diacetate (DAF-2DA, Sigma-Aldrich), as described by Lamotte et al. (2004). After filtration, Nicotiana tabacum cv Xanthi cell suspensions were incubated in the H10 suspension buffer containing 20 µM DAF-2DA for 1 h in the dark at 24°C on a rotary shaker (150 rpm) and then rinsed twice with fresh suspension buffer to wash off excessive fluorophore. Cells were then transferred into 24-well plates (Costar, Corning Incorporated, Corning, NY, USA) containing 1 mL of cells per well, and treated with cryptogein in the dark. NO production was measured using a 24-well reader fluorometer (Fluoroskan Ascent fluorometer, Labsystems, Helsinki, Finland) with 485 nm excitation and 510 nm emission filters. Fluorescence was expressed as relative fluorescence units. For each treatment, measurements of NO production over time were performed on the same batch of cells.

H2O2 Production
AOS production was determined by chemiluminescence, as described by Lamotte et al. (2004). Twenty minutes after adding cryptogein to tobacco cell suspensions, duplicate aliquots of 250 µL were collected from the batch of treated cell suspensions and mixed with 300 µL of 10 mM Hepes buffer, pH 6.5, 175 mM mannitol, 0.5 mM CaCl2, 0.5 mM K2SO4, and 50 µL of 0.3-mM luminol. Chemiluminescence measured using a luminometer (Lumat LB9507, Berthold, Bad Wildbad, Germany) was integrated and expressed in nanomoles of H2O2 per gram fresh weight of cells.

Analysis of Cytosolic Free Calcium Concentrations
In vivo reconstitution of aequorin was performed by addition of 1 µM coelenterazine to cells in the H10 suspension buffer for at least 3 h in the dark (150 rpm, 24°C). Coelenterazine is a prosthetic group of aequorin required for its full Ca2+-binding activity. The bioluminescence of 250-µL aliquots of cells (maintained in a cuvette) was recorded continuously at 1-s intervals using a digital luminometer (Lumat LB9507, Berthold, Bad Wildbad, Germany). At the end of the experiments, residual functional aequorin was quantified by adding 300 µL of lysis buffer (10 mM CaCl2; 2% Nonidet P40, v/v; 10% ethanol, v/v) and monitoring the resulting increase in luminescence. Luminescence data transformation into cytosolic Ca2+ concentration was calculated as previously described (Lamotte et al., 2004).

In-Gel Kinase Assay
After treatment, Nicotiana plumbaginifolia cells (0.25 g) were harvested by filtration, frozen in liquid nitrogen, and ground in a mortar. Preparation of protein extracts was performed as previously reported (Lamotte et al., 2006). Protein extracts (30 µg) were electrophoresed in 10% SDS-polyacrylamide gels embedded with 0.14 mg mL–1 histone IIIS (HIIIS) or with 0.20 mg mL–1 myelin basic protein (MBP). Then, SDS was removed by washing the gels for 1 h with the washing buffer (50 mM Tris-HCl pH 8.0; 20% 2-propanol). The gels were then equilibrated for 1 h in buffer B (50 mM Tris-HCl pH 8.0; 5 mM 2-β-mercaptoethanol). Subsequently, proteins were denaturated for 1 h with 6 M guanidine-HCl in buffer B, and allowed to re-nature overnight at 4°C in buffer B containing 0.04% Tween 40. The gels were then equilibrated for 30 min at room temperature in 10 mL of the reaction buffer (40 mM HEPES pH 7.5; 0.5 mM CaCl2 ; 20 mM MgCl2 and 2 mM DTT) and then for 1.5 h in the reaction buffer supplemented with 25 µM ATP and 15 µCi of {gamma}-32P-ATP (Amersham). The reaction was stopped by transferring the gels into 5% (w/v) trichloroacetic acid and 1% (w/v) sodium phosphate. The unincorporated {gamma}-32P-ATP was removed by extensive gel washing in the same solution for at least 2.5 h. Finally, the gels were dried onto 3MM paper (Whatman) and exposed to X-Omat AR film (Kodak). Prestained molecular markers (Bio-Rad) were used to calculate the apparent size of kinases.

Immunoprecipitation Assay
Briefly, proteins from crude extracts (100 µg) of untreated and NO-treated Nicotiana plumbaginifolia cells were incubated with 5 µg polyclonal anti-SIPK antibodies and 20 µL of packed volume of protein A-agarose (Sigma-Aldrich, Saint-Quentin-Fallavier, France) in immunoprecipitation buffer (20 mM Tris pH 7.5; 2 mM EDTA; 2 mM EGTA; 50 mM β-glycerophosphate; 100 µM Na3VO4; 2 mM DTT; 500 µM phenylmethylsulfonyl fluoride; 1 µM leupeptin; 1 µM pepstatin; 1% Triton X-100 and 150 mM NaCl) at 4°C for 12 h on a rocker. Agarose-bead complexes were pelleted by brief centrifugation, washed four times with immunoprecipitation buffer, and resuspended in 20 µL of sample buffer. The samples were then heated at 95°C for 4 min. After brief centrifugation, the supernatant was analyzed by in-gel kinase assays using HIIIS as a substrate.

Western Blot Analysis
Protein samples (30 µg) were separated on 10% SDS-polyacrylamide gels and transferred to nitrocellulose membranes (Hybond-ECL, Amersham, Piscataway, NJ). Membrane blocking and detection were performed according to the instructions supplied with the ECL Plus kit (Amersham, Piscataway, NJ). Polyclonal primary antibodies raised against the phosphorylated form of human ERK1 and ERK2 (Phospho-p44/42 Map kinase Antibody, Cell Signaling Technology n° 9101) were used at a dilution 1:1000. These antibodies detect endogenous levels of ERK-related MAPK only when catalytically activated by phosphorylation at Thr202 and Tyr204. Anti-SIPK antibodies were used at a final concentration of 0.2 µg/mL. The goat anti-rabbit IG conjugated to horseradish peroxidase (Sigma) was used at a dilution of 1:25 000.


   SUPPLEMENTARY DATA
TOP
 Abstract
 INTRODUCTION
 ORIGINS OF NO IN...
 DISCUSSION
 METHODS
 SUPPLEMENTARY DATA
 
Supplementary Data are available at www.mplant.oxfordjournals.org


   Acknowledgements
 
The panorama of the field offered here has been simplistic and we apologize to those colleagues whose work we did not review. We are indebted to Prof. D.F. Klessig (Boyce Thompson Institute for Plant Research, Ithaca, USA), who provided anti-SIPK antibodies. We thank Annie Buchwalter for helpful discussions. Foundings: Ministère de l'Education Nationale, de la Recherche et de la Technologie; Agence Nationale de la Recherche (BLAN07-2–184783); Institut National de la Recherche Agronomique (SPE 1088a); Conseil Régional de Bourgogne (HCP 189); Ministère des Affaires Etrangères (EGIDE Polonium, grant 11545WG); Polish Ministry of Education and Science (grant PBZ-KBN-110/PO4/2004). No conflict of interest declared.

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