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As nitric oxide (NO) is a key messenger in many organisms, reliable techniques for the detection of NO are essential. Here, it is shown that a combination of membrane inlet mass spectrometry (MIMS) and restriction capillary inlet mass spectrometry (RIMS) allows for the fast, specific, and non-invasive online detection of NO that has been emitted from tissue cultures of diverse organisms, or from whole plants. As an advantage over other NO assays, MIMS/RIMS discriminates nitrogen isotopes and simultaneously measures NO and O2 (and other gases) from the same sample. MIMS/RIMS technology may thus help to identify the source of gaseous NO in cells, and elucidate the relationship between primary gas metabolism and NO formation. Using RIMS, it is demonstrated that the novel fungicide F 500® triggers NO production in plants.
Recently, a spectrofluorometric assay, based on the binding of NO to 4,5-diaminofluorescein diacetate (DAF-2 DA), was shown to be suited for the direct measurement of NO from plant cells in suspension culture (Tun et al., 2001). However, when monitoring spatio-temporal aspects of NO production with DAF-2 DA and confocal laser scanning (Foissner et al., 2000) or epifluorescence microscopy (Gould et al., 2003; Pedroso et al., 2000) in plant tissue samples, these need to be wounded during sample preparation. Therefore, they might display unmeant NO emissions (Foissner et al., 2000). In addition, DAF-2 DA-based estimations of NO are often affected by differences in dye loading between different tissue types or specific organelles (Foissner et al., 2000).
Greater accuracy in the measurement of NO must be based on direct assays, such as spin trapping electron paramagnetic resonance (Caro and Puntarulo, 1999; Huang et al., 2004; Pagnussat et al., 2002), photoacoustic laser spectroscopy (Leshem and Pinchasov, 2000; Mur et al., 2003), or mass spectrometry (Lewis et al., 1993). Eleven years ago, these authors reported the direct and simple measurement of NO emissions from mammalian cell cultures by a mass spectrometric assay referred to as membrane inlet mass spectrometry (MIMS; Lewis et al., 1993). However, many biologists have glossed over this report. As a further development of the technique, we here report that a combination of MIMS and restriction capillary inlet mass spectrometry (RIMS) allows for the direct, fast, specific, and non-invasive online detection of NO from both liquid suspensions (MIMS) and the gaseous phase (RIMS).
Results and discussion
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.
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
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
In the MIMS assays, addition of nitrite in the absence of respective cells did not cause detectable NO production. In the RIMS experiments, spraying leaves or plants with water or equimolar concentrations of phosphate did not elicit detectable NO release. In all the experiments, concentration of O2 was <1% (v/v).
Mammalian cell cultures
Tobacco tissue culture
Parsley tissue culture
Soybean tissue culture
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.
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).
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).
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).
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).
Because NO is an important signal in diverse organisms, reliable techniques for its detection are required. MIMS/RIMS represents an advanced method for the non-invasive, fast, and specific detection of NO in cell cultures, tissue samples, and whole plants. The MIMS/RIMS setup is easy to handle and calibrate. Furthermore, it only requires a low-cost benchtop mass spectrometer and allows real-time measurements of extracellular NO under non-invasive conditions. Although being less sensitive than photoacustic laser spectroscopy, MIMS/RIMS technology offers a number of advantages: (i) NO measurements are possible from gaseous and liquid phases; (ii) in addition to NO, other gases (i.e. CO2, O2, NO2) can be measured simultaneously, and even the analysis of more complex gases (e.g. ethylene, methyl jasmonate) is possible with the same experimental setup; (iii) isotope tracing experiments (e.g. with 14N/15N) can be performed in order to elucidate the origin of extracellular NO in leaves and aquatic suspensions; and (iv) the simple handling of the MIMS/RIMS system makes it an excellent candidate for routine NO measurements, as well as for automation processes. Thus, on a long-term basis, the MIMS/RIMS technology might help to shed further light into the enigmas of the chemistry and biology of the NO key signal of life.
15NaNO2 was obtained from Chemotrade Inc. (Leipzig, Germany); 14NaNO2, salicylhydroxamic acid, and PTIO were purchased from Sigma-Aldrich (Taufkirchen, Germany). F 500® was synthesized by BASF AG and provided as an EC formulation of 250 g l−1 (BASF Code# BAS 500 01 F). A placebo formulation void of F 500® (BASF Code# BAS 500 00 F) was used for control treatments.
Mouse macrophage cell cultures RAW264.7 (TIB-71) were purchased from the American Tissue Culture Collection, and grown at 37°C in 150-ml tissue culture flasks in HEPES (25 mm)-buffered Roswell Park Memorial Institute (RPMI) 1640 medium enriched with l-glutamine. After formation of a continuous monolayer, cells were removed from the bottom of the culture flask by gentle agitation and subjected to centrifugation (3000 g, 5 min). The pellet was re-suspended in 10 ml of fresh growth medium.
Parsley, tobacco (cv. Xanthi nc), and soybean (cv. Williams 82) tissue cultures were grown as described by Kauss et al. (1992), Hennig et al. (1993), and Levine et al. (1994), respectively. Four to six-day-old cell cultures were washed on a funnel and suspended in fresh growth medium at a density of approximately 80 mg ml−1. Fifty-milliliter aliquots of cell suspension were allowed to adapt in 250-ml Erlenmeyer flasks by shaking for at least 1 h at 120 r.p.m.
Chlamydomonas reinhardtii (strain 11–32b) was obtained from the Sammlung für Algenkulturen (Göttingen, Germany); Synechocystis sp. PCC6803 and Synechoccocus sp. PCC7942 were purchased from the Pasteur Culture Collection (Paris, France). C. reinhardtii and the cyanobacteria were grown in high salt minimal medium (Sueoka et al., 1967) and BG-11 medium (Allen, 1968), respectively. Cells were kept under continuous light at 30°C in air enriched with 5% (v/v) CO2. Cells were centrifuged (5000 g, 5 min) and concentrated in assay buffer containing 50 mm Bis–Tris-Propane/HCl (pH 8.0). Aliquots of cell suspension were adjusted to a final chlorophyll concentration of 5–10 µg ml−1, which was determined according to Porra et al. (1989) before use in the experiments.
Pythium sp., Botrytis sp., and Fusarium sp. mycelia were grown in synthetic CDN (Czapek Dox Normal) medium (Cooper et al., 1984) at room temperature, and were used in the assays after 7–10 days.
Tobacco (Nicotiana tabacum cv. Xanthi nc) plants were grown at 22°C with 60% relative humidity in a 16-h light cycle, and used in the experiments at 6–8 weeks. Arabidopsis (ecotype Columbia) plants were grown for 4–6 weeks at an 8-h photoperiod and at 22°C with 60% relative humidity before use in the assays.
Pseudomonas syringae pv. tomato (strain DC3000) and P. syringae pv. glycinea (AvrC) were grown at 30°C in King's B medium (King et al., 1954) for 1 day. After centrifugation (5000 g, 5 min), bacterial cells were washed and re-suspended to approximately 35 × 106 cfu ml−1 in 10 mm MgCl2. 1 × 107 bacteria were used for infection of tobacco or soybean cell cultures.
NO and O2 assay
Ten-milliliter aliquots of tissue culture were transferred to the aquatic sample chamber of the mass spectrometer (model HP5970B; Figure 1). Upon respective treatments, external NO and/or O2 was continuously monitored using MIMS.
In the experiments with intact tobacco leaves, these were inserted into the leaf/plant cuvette of the mass spectrometer (Figure 1a). The release of NO was determined using RIMS. In case of Arabidopsis, a whole plant was transferred to the leaf/plant cuvette of the mass spectrometer and used for RIMS analysis.
This research was supported by a grant to U.C and D.F.S. from BASF. We thank Bernhard Brüne, Raimund Tenhaken, and Matthias Hahn for providing mouse macrophage, soybean, and Botrytis sp. cell cultures, respectively. We also appreciate provision of Pythium sp. and Fusarium sp. mycelia suspensions by John Speakman. Luis Mur is thanked for valuable information on photoacoustic laser spectrometry.