In MIMS, a semipermeable membrane directly faces a cell suspension in a temperature-adjustable translucent reaction chamber (Figure 1a; Fock and Sültemeyer, 1989). Dissolved gases diffuse through the membrane into a capillary before entering the ion source of a benchtop mass spectrometer. In RIMS (Figure 1a), however, intact plant leaves or small plants are incubated in a translucent chamber that is attached to the mass spectrometer via a thin restriction capillary. In combined MIMS/RIMS assay, NO and other gases (i.e. O2, CO2, NO2, etc.) from a sample pass either the membrane (MIMS) or the restriction capillary (RIMS), and then directly evaporate into the ionization chamber of a benchtop mass spectrometer, thus allowing sensitive levels of detection.
Figure 1. Schematic diagram of the experimental setup for MIMS/RIMS-based NO measurements (a), and verification of signal specificity by the addition of NO donors (b), injection of NO-saturated water (c), and scavenging of the NO signal by PTIO (b,c).
(a) In MIMS, a cell suspension in an 8–10-ml reaction chamber is circulated over a thin (50 µm) teflon membrane by a magnetic stirrer. Dissolved gases, such as NO, diffuse through the membrane and evaporate into the ionization chamber of a mass spectrometer. In RIMS, a metal bellows pump ensures rapid and efficient mixture of the 120-ml gas phase, which includes the volume of a leaf cuvette (8 cm × 8 cm × 0.4 cm). Before entering the mass spectrometer, NO and other gases pass a restriction capillary (inner diameter: 0.1 mm; length: 2 m). A three-way valve serves to switch between the two sample chambers.
(b) Changes in the abundance of mass 30 (as measured by MIMS) after addition of 180 µm of the NO-releasing compounds SNAP and GSNO into 8 ml of O2-free buffer (HEPES/NaOH, pH 7.8). The time of injection is indicated by an open arrow.
(c) For calibration of the NO signal, the sample chamber was filled with 10 ml of water and aerated with N2 for up to 5 min until the concentration of O2 was nearly zero. At the times denoted by the open arrows, 5 µl of NO-saturated water (corresponding to 1.9 mm NO at 20°C) was added, resulting in a final NO concentration of 0.95 µm. Numbers give the abundance (units at m/z = 30) for two extremes of the received NO signal, which assigns 1 abundance unit to 10 or 13 pmol of NO.
In (b) and (c), signal specificity was further validated by the addition of the NO scavenger PTIO (150 µm) at the times indicated by the filled arrows. The dotted lines indicate zero NO.
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The specificity of the NO signal (m/z = 30) was evaluated using MIMS in three independent tests. (i) Addition of NO-releasing compounds, e.g. S-nitroso-N-acetyl-dl-penicillamine (SNAP) and S-nitroso-l-glutathione (GSNO), caused an immediate rise in the NO signal (Figure 1b). Note that although same concentrations (180 µm) of the NO-donors were applied in Figure 1(b), the rate of NO release was about five times higher with SNAP than it was with GSNO (Figure 1b). This finding might explain the higher efficiency of SNAP to activate distinct physiological responses in various biological systems or cell types (Durner et al., 1998). (ii) Injection of small aliquots of NO-saturated water into 10 ml of H2O resulted in distinct signals of m/z = 30 within less than 6 sec (Figure 1c). Such a procedure can be used for simple calibration of the NO signal, which, for our system, revealed that 1 abundance at m/z = 30 corresponded to 10–13 pmol of NO. (iii) Addition of specific NO scavengers, such as 2-phenyl-4,4,5,5-tetramethylimidazolinone-3-oxide-1-oxyl (PTIO), caused a rapid decrease in signal intensity to nearly zero NO (Figure 1b,c). Together, these data clearly show that the mass spectrometric technique allows for the specific and sensitive determination of NO, and that the system is fast enough to permit real-time measurements of changes in NO levels (Figure 1b,c).
Online detection of NO from various organisms by MIMS/RIMS assay
It has been suggested recently that various plants (Garcia-Mata and Lamattina, 2003; Rockel et al., 2002; Yamasaki et al., 1999), bacteria (Ji and Hollocher, 1988), and fungi (Yamasaki, 2000) can produce and then emit gaseous NO into the environment from nitrite, either via the non-enzymatic reduction of apoplastic nitrite (Bethke et al., 2004), or in a side reaction catalyzed by nitrate reductase (NR; Rockel et al., 2002). The NO-releasing activity of NR was facilitated at high nitrite levels and low oxygen concentrations (Rockel et al., 2002). In fact, we observed considerable reduction (more than 50%) in NO yield in the presence of 21% (v/v) O2 (data not shown). Therefore, the subsequent experiments were all performed at low oxygen concentrations (under 1%, v/v).
To validate the suitability of MIMS/RIMS to detect NO emissions from different biological sources, a variety of tissues from diverse organisms was treated with nitrite at a low oxygen level, and was assayed for the release of gaseous NO into the environment. Upon nitrate addition, extracellular NO levels continuously increased in cell cultures of mouse, higher plants, algae, and cyanobacteria, and also in suspended fungal mycelia, tobacco leaves, and whole Arabidopsis plants (Table 1; Figure 2).
Table 1. Organisms for which nitrite-induced NO release has been demonstrated using adopted MIMS/RIMS assay
|Mammalian cell cultures|
| Mouse macrophages||MIMS|
| Tobacco tissue culture||MIMS|
| Parsley tissue culture||MIMS|
| Soybean tissue culture||MIMS|
| Tobacco leaves||RIMS|
| Arabidopsis plants||RIMS|
| C. reinhardtii||MIMS|
| Pythium sp.||MIMS|
| Botrytis sp.||MIMS|
| Fusarium sp.||MIMS|
| Synechocystis PCC6803||MIMS|
| Synechococcus PCC7942||MIMS|
Figure 2. Validation of MIMS/RIMS-based NO assay.
A low (approximately 0.5%, v/v) oxygen level by means of nitrite (20 mm)-induces NO release in cell suspensions of C. reinhardtii (trace A), soybean (trace B), Fusarium sp. (trace C), and in a tobacco leaf (trace D). For trace D, one leaf of a tobacco plant was sprayed with 20 mm NaNO2, and the subsequent NO release from the leaf was assayed using RIMS. When a sheet of paper was used instead of the tobacco leaf, emission of NO could not be detected. During the assays, temperature was kept at 20°C. Arrows denote the times of nitrite addition.
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Like most of the other techniques used to quantify NO, MIMS/RIMS allows the detection of only extracellular NO. Bearing in mind that NO is a highly active molecule and that only part of the endogenously produced NO may be released as a gas, the MIMS/RIMS technique, although being indicative, is unlikely to detect small increases in the level of intracellular NO. For this purpose, DAF-2 DA-based NO detection, combined with microscopical techniques, needs to be employed (Foissner et al., 2000; Gould et al., 2003; Pedroso et al., 2000). This methodology, however, displays a variety of other disadvantages (see Introduction).
During optimal nitrogen assimilation of plants, cytoplasmic NR catalyzes the reduction of nitrate to nitrite. The latter compound is translocated into the chloroplasts, where it is reduced to NH4+ by nitrite reductase. Thus, under optimal conditions, nitrite and NR do not come together in significant amounts to allow for a major production of NO by NR. However, the nitrite concentration in the cytoplasm could increase to a significant level in the absence of an electrochemical gradient across the chloroplastic envelope. This is when photosynthetic electron transport is impaired (Shingles et al., 1996), for example, at night (Wildt et al., 1997), or when plant cells are attacked by necrotizing pathogens (Agrios, 1997). Alternatively, at least part of the NO that is emitted from nitrite-treated plant tissues (Table 1; Figure 2) may result from the non-enzymatic reduction of nitrite in the apoplast (Bethke et al., 2004).
It is remarkable that nitrite is able to induce the emission of NO also in phytopathogenic fungi (Table 1; Figure 2). Because NO can easily diffuse between cells, it is possible that in attacking fungi, plant-derived nitrite elicits the formation of NO, which might then serve to further support the plant's defense response (Delledonne et al., 1998; Durner et al., 1998).
In addition to nitrite treatment, plant cells can also release NO when exposed to pathogens (Delledonne et al., 1998; Durner et al., 1998). In this case, NO likely is produced in a reaction that is catalyzed by a variant form of the P-protein of the mitochondrial glycine decarboxylase complex. Interestingly, the biochemical properties of variant P resemble those of mammalian NO synthases (Chandok et al., 2003).
To investigate whether MIMS/RIMS technology is suited to also record the emission of NO in plant–pathogen interactions, suspension-cultured tobacco and soybean cells were treated with avirulent Pseudomonas syringae pv. tomato or virulent P. syringae pv. glycinea, respectively, and assayed for NO release using MIMS.
As shown in Figure 3, in the incompatible interaction between tobacco cells and P. syringae pv. tomato, there was a rapid MIMS-detectable NO burst that was maximal after approximately 1 h of treatment, and followed by a prominent second burst of NO from the 4- through 8-h time point (Figure 3, trace A). In the compatible interaction between soybean cells and P. syringae pv. glycinea, both timing and extent of the early NO burst were similar to the one detected in the incompatible tobacco–P. syringae pv. tomato interaction. The second NO burst, however, was much less pronounced (Figure 3, trace B). These results are in line with earlier findings from compatible and incompatible plant–pathogen interactions obtained using an oxyhemoglobin-/methemoglobin-based NO assay (Delledonne et al., 1998). Together with these data, the results in Figure 3 support the assumption that the second NO burst is one likely key event in determining avirulence in plant–pathogen interactions.
Figure 3. Nitric oxide emission in plant–pathogen interactions.
Cell suspension cultures of tobacco (trace A) and soybean (trace B) were incubated at time zero with avirulent P. syringae pv. tomato or virulent P. syringae pv. glycinea, respectively.
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Fungicide-induced NO release from plants and fungi
It has been speculated recently that the enhanced pathogen resistance of tobacco induced by the novel strobilurin fungicide F 500® (common name: pyraclostrobin; Herms et al., 2002), as well as some aspects of the F 500®-induced ‘physiological side-effect’ in plants, might be because of F 500®-induced NO release (Köhle et al., 2002).
MIMS analysis revealed that in tobacco cell cultures, F 500® in fact elicited a huge emission of NO after approximately 5 min (Figure 4a), which steadily increased over the 24-h detection period (Figure 4b), while respiratory O2 uptake was dramatically inhibited (Figure 4c). In a similar manner, cultured soybean cells also exhibited F 500®-dependent NO release, which was maximal after 5 h and remained essentially unchanged over the next 19 h (Figure 4d). In intact tobacco leaves, F 500® caused an immediate emission of NO that could still be detected at the fifth day post-treatment (Figure 4e). The latter result fits with the observation that F 500® rapidly enters plant leaves where its effects last for a prolonged period (Stierl et al., 2002).
Figure 4. MIMS/RIMS assay of F 500®-induced NO release, and alterations in respiration rate of plants and fungi.
Tobacco (a–c), soybean (d), and Botrytis sp. (f,g) cell suspensions were treated with F 500® (5 µm, final concentration) at time zero (indicated by the filled arrow in (a)). External NO (a,b,d,f) and O2 (c,g) were determined using MIMS. In (a), specificity of the NO signal was confirmed by addition of the NO scavenger PTIO (150 µm, final concentration) at the time denoted by the open arrow. In (b,c,f,g), con stands for control measurements with samples to which the same concentration of solvent (DMSO) was added. (e) On two tobacco plants, one leaf each was treated with an EC formulation of F 500® (0.6 mm) or with a placebo formulation void of F 500®. NO release was determined using RIMS immediately or after another 5 days. SHAM, salicylhydroxamic acid.
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Interestingly, within 10 min after its application, F 500® was able to induce NO release in various fungi also (data not shown), including Botrytis sp. (Figure 4f). In the latter, F 500®-induced NO production was accompanied by an approximately 50% inhibition of respiration (Figure 4g). Further treatment with salicylhydroxamic acid, an artificial chemical inhibitor of alternative and F 500®-resistant respiration, led to an additional evolution of NO (Figure 4f).
Together, these MIMS/RIMS data point to NO as an important player in both F 500®-induced disease resistance and F 500®-caused ‘physiological side-effect’ in plants. The results obtained also indicate that fungus-generated NO could be involved in triggering plant defense responses to pathogen attack. Finally, with O2 consumption as an example, the data show that MIMS/RIMS technology allows investigations as to the relationship between gaseous parameters of primary metabolism and NO formation (Figure 4b,c,f,g).
As mentioned earlier, NO in plants can be produced from nitrite either via non-enzymatic reduction in the apoplast (Bethke et al., 2004), or in a side reaction catalyzed by NR (Garcia-Mata and Lamattina, 2003; Rockel et al., 2002; Yamasaki et al., 1999). A NR-like enzyme in the plasma membrane of root cells, which uses nitrite as a substrate (Stöhr et al., 2001), a P-protein-like enzyme (Chandok et al., 2003), and a recently identified plant NO synthase (Guo et al., 2003) can also contribute to NO production. Furthermore, NO in plants can be produced non-enzymatically through light-mediated conversion of NO2 by carotenoids (Millar and Day, 1997). Thus, an important issue in NO signal transduction research in plants relates to the source of NO under different physiological conditions.
With nitrite-induced NO release as the example, Figure 5 demonstrates that upon subsequent addition of 14N and 15N-labeled nitrite to the same tobacco cell culture, MIMS is able to distinguish between subsequently released 14NO and 15NO isotopomeres. Therefore, isotope tracing experiments, combined with MIMS/RIMS analysis, may be helpful in identifying the source of gaseous NO in cells. Future experiments with 15N-arginine-loaded cells may provide physiological evidence for mammalian NO synthase-like enzyme activity in plants (Chandok et al., 2003; Guo et al., 2003).
Figure 5. 14N/15N-isotope tracing.
14N- and 15N-nitrite (each at 20 mm) were subsequently added to the same tobacco cell suspension at the times indicated by the filled and open arrows, respectively. The subsequent emission of 14NO or 15NO was simultaneously detected using MIMS assay. We failed to detect any NO production in the absence of cells under the conditions used.
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The isotope tracing – MIMS/RIMS – combination represents a major advance in mass spectrometry-based NO detection and is an advantage over other techniques used to monitor NO production, including electron paramagnetic resonance (Caro and Puntarulo, 1999) and photoacoustic laser spectroscopy (Leshem and Pinchasov, 2000; Mur et al., 2003).