Molecular Plant Advance Access originally published online on December 13, 2007
Molecular Plant 2008 1(2):270-284; doi:10.1093/mp/ssm020
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Measuring NO Production by Plant Tissues and Suspension Cultured Cells
a Department of Plant Biology, Faculty of Agronomy, Mendel University of Agriculture and Forestry, Zemedelska 1, Brno 613 00, Czech Republic
b Department of Plant and Microbial Biology, University of California, Berkeley, CA 94720–3102, USA
1 To whom correspondence should be addressed. E-mail jan.vitecek{at}seznam.cz, fax +420 545 133 025, tel. +420 545 133 344.
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Abstract |
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 TOP Abstract INTRODUCTIONRESULTSDISCUSSIONMETHODS |
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We describe an inexpensive and reliable detector for measuring NO emitted in the gas phase from plants. The method relies on the use of a strong oxidizer to convert NO to NO2 and subsequent capture of NO2 by a Griess reagent trap. The set-up approaches the sensitivity for NO comparable to that of instruments based on chemiluminescence and photoacoustic detectors. We demonstrate the utility of our set-up by measuring NO produced by a variety of well established plant sources. NO produced by nitrate reductase (NR) in tobacco leaves and barley aleurone was readily detected, as was the production of NO from nitrite by the incubation medium of barley aleurone. Arabidopsis mutants that overproduce NO or lack NO-synthase (AtNOS1) also displayed the expected NO synthesis phenotype when assayed by our set-up. We could also measure NO production from elicitor-treated suspension cultured cells using this set-up. Further, we have focused on the detection of NO by a widely used fluorescent probe 4-amino-5-methylamino-2',7'-difluorofluorescein (DAF-FM). Our work points to the pitfalls that must be avoided when using DAF-FM to detect the production of NO by plant tissues. In addition to the dramatic effects that pH can have on fluorescence from DAF-FM, the widely used NO scavengers 2-phenyl-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (PTIO) and 2-(4-carboxyphenyl)-4,4,5,5-tetramethylimidazoline-1-oxyl-3-oxide (cPTIO) can produce anomalous and unexpected results. Perhaps the most serious drawback of DAF-FM is its ability to bind to dead cells and remain NO-sensitive.
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INTRODUCTION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
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There is growing interest in the biology of NO in plants and many articles have appeared during the last ten years that have focused on NO and its synthesis (Lamattina et al., 2003). There are several routes for NO synthesis in plants, including its non-enzymatic production from nitrite (Yamasaki, 2000; Bethke et al., 2004) and enzymatically by NR (Dean and Harper, 1986; Kaiser et al., 2002; Rockel et al., 2002; Stöhr and Ullrich, 2002) and nitric oxide synthases (Guo et al., 2003). The existence of bona fide plant nitric oxide synthases remains controversial, however, and plant biologists have therefore relied on more indirect methods for establishing whether and where NO is produced by specific cells and tissues.
Many methods have been developed for NO detection and they capitalize on the high diffusibility of NO as well as its broad spectrum of chemical reactivity. In general, methodological approaches to detect NO in plants can be classified based on the level of contact of the detection system with the biological material. The detection of NO in the gas phase is currently attracting much attention, especially because of the spatial separation of the biological material from the detector. Further, these systems can be utilized for real-time measurement of NO synthesis. Instruments based on chemiluminescence detectors showing high specificity and sensitivity have been used to measure NO production from whole plants or excised organs as well as by cell suspension cultures (Rockel et al., 2002; Planchet et al., 2005; Planchet and Kaiser, 2006; Planchet et al., 2006). The detection limit of chemiluminescence instruments is about 20–50 pmol NO and they are commonly designed to quantify NO in the range 2–4000 ppb (Byrnes et al., 1996).
As an alternative to chemiluminescence measurement, laser photoacoustic spectroscopy was recently applied to measurement of NO produced by plants (Leshem and Pinchasov, 2000; Mur et al., 2005), having a detection limit of around 20 pmol NO (Mur et al., 2005). Mass spectroscopy was also utilized to detect NO in the gas phase (Conrath et al., 2004). This method is also capable of direct sampling of NO from the aqueous phase using a restriction capillary (Conrath et al., 2004), or via a semi-permeable membrane (Bethke et al., 2004). Carbon-NO selective electrodes have also been developed that allow NO detection in solution, and a limit of detection of less than 1 nM NO can be achieved with this approach (Zhang and Broderick, 2000). Further, electron paramagnetic resonance was utilized to detect NO released from plant organs (Dordas et al., 2003; Xu et al., 2004; Modolo et al., 2005). In general, the detection limit for electron paramagnetic resonance is about 10 nM (Nagano, 2002).
Whereas these approaches have the sensitivity and specificity required to make precise measurements of NO synthesis, many are expensive and some have a high demand on workspace. In this manuscript, we describe a simple and inexpensive method for routine measurement of NO. The principle underlying this method relies on well understood principles of NO chemistry. CrO3 is used as a strong oxidizer to convert NO to NO2, which, in turn, is captured by a trap containing Griess reagent. The NO2 that is trapped can be easily visualized by color changes in the Griess reagent and can be quantified by spectrophotometry.
Several methods have been developed for measuring NO that rely on the interaction of a detecting molecule with the biological material. These include the use of hemoglobin to quantify NO produced by suspension-cultured cells (Delledonne et al., 1998) and from callus culture (Zhao et al., 2004), and various types of fluorescent probes. Indeed, fluorescent probes are the most widely utilized for NO detection in plants. Among the most commonly used fluorescent probes is diaminofluorescein (DAF) and its membrane-permeable diacetate form (Kojima et al., 1998). DAF has been widely used to localize the site of NO production in plant cells and tissues (Correa-Aragunde et al., 2004; Gabaldón et al., 2005; Hu et al., 2005), and to quantitate the production of NO by suspension-cultured cells in vitro (Krause and Durner, 2004). In each of these studies, proof that DAF was measuring localized NO accumulation was given by the ability of the NO scavenger cPTIO to quench the fluorescence signal from DAF. Although these experiments demonstrated the localization of NO production in plant tissues, there are reasons to urge caution in the application of this method. One reason for caution in the use of dyes such as DAF is that their utility may be limited by the local biological environment. DAF does not react directly with NO but with the intermediate of NO oxidation, dinitrogen trioxide (N2O3) (Kojima et al., 1998) (Scheme 1). For example, reducing agents such as ascorbate and glutathione that are abundant in plant tissues (e.g. Foyer and Noctor, 2005) can scavenge compounds such as N2O3 and NO2 (Rodriguez et al., 2005) and potentially limit NO detection by DAF.
The highly acidic nature of the plant apoplast and the presence of reducing agents in the plant cell wall can also prove a complicating feature for the detection of NO production because this type of environment can lead to the non-enzymatic production of NO from nitrite (Bethke et al., 2004). In their study of NO production by tracheary elements of Zinnia elegans, Gabaldón et al. (2005) showed that mature tracheary elements that are likely to be non-living are a source of fluorescent signals that these authors associate with a burst of NO immediately prior to cell death. Non-living plant cells can be a source of fluorescence signals from DAF, and a non-enzymatic source of NO was suggested to be one component in the asymmetric production of NO in gravi-responding soybean roots (Hu et al., 2005). That soybean roots were producing NO non-enzymatically in the apoplast was supported by experiments showing that inhibitors of neither NOS nor NR completely abolished the fluorescent signal from DAF in the gravitropic bending of the root (Hu et al., 2005).
These reports led us to investigate the response of the DAF derivative DAF-FM (Kojima et al., 1999) to changes in the local environment both in vitro and in vivo in plant cells and tissues, as well as the effectiveness of PTIO and cPTIO as scavengers of the fluorescent signal from DAF-FM. In this manuscript, we report on data that confirm the observations made by others working mainly with mammalian cells (e.g. Gomes et al., 2006), emphasizing that caution must be exercised when using fluorescent probes in experiments with living cells and tissues and that data obtained with these dyes must be interpreted conservatively.
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RESULTS |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
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Characteristics of the Oxidizer Column NO Detector
Details of the set-up for the detection of NO with the oxidizer column are described in ‘Materials and Methods’. Briefly, a stream of humidified air is pumped over a sample in an enclosed sample chamber (Figure 1). This stream is first passed through a trap containing Griess reagent to remove traces of HNO2 and passed over a column containing the strong oxidizing agent CrO3. The air stream then goes through a second Griess reagent trap that collects any NO that was oxidized to NO2. Virtually all HNO2 or NO2 were removed from the gas phase by traps 1 and 2 (not shown). To establish that this simple bench-top system can quantitatively and specifically measure low concentrations of NO gas, several controls were performed.
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Detection of NO using our set-up varies according to the rate of airflow through the system (Figure 2A). The amount of NO captured was determined when air containing 9.9 ppm NO was allowed to flow at rates of between 5 and 40 ml min–1. Airflow in these experiments was regulated using a computer-regulated series of mass flow controllers (Libourel et al., 2006). There is a statistically significant increase in the amount of NO captured as the air flow increases, and, at 40 ml min–1 (the limiting flow rate of the system), more than 60% of the added NO is oxidized and trapped in the second Griess reagent trap.
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Next, we examined the effect of NO concentration on the efficiency of NO capture using an airflow of 40 ml min–1 and NO concentrations of between 10 ppb and 8 ppm. As the data in Figure 2B show, more than 90% of NO is captured at concentrations below 1 ppm but the amount of NO captured above this concentration drops sharply. There is about a 20% decline in the amount of NO captured by the set-up as NO concentrations increase from 1 to 8 ppm (Figure 2B).
During operation, the oxidizer column became hydrated, as demonstrated by a color change from brick-red to yellow. The column showed a constant recovery rate for NO until more than two-thirds of the column packing became hydrated, allowing about 3 h of continuous measurement at an airflow rate of 40 ml min–1. Using pre-columns with molecular sieves or spargers with a saturated solution of CaCl2 to remove water vapor from the gas phase did not extend column life, but rather caused failure of NO detection.
The sensitivity of the system was also determined by diluting NO gas using our mass flow controller system (Libourel et al., 2006) to concentrations in the range of 10–50 ppb and measuring the amount of NO captured in the second Griess trap. These low concentrations of NO were pumped through the system at 40 ml min–1. A detection limit of between 1 and 2 ppb was calculated (3xsignal/noise ratio) representing 0.1–0.2 nmol h–1 NO under standard pressure and temperature (101.3 kPa and 25°C, respectively) and flow rate specified above.
We also determined the specificity of the oxidizing/trapping method by introducing a range of volatile, low molecular mass nitrogen-containing compounds into the system at a flow rate of 40 ml min–1. Among the compounds tested that gave no signal were cyanide, azide and ammonia. Nitrous oxide, on the other hand, gave a positive signal in the second Griess trap, but only at concentrations of around 1000 ppm.
Measuring NO Produced from Biological Sources
To establish that the oxidizer column set-up can function to quantitate NO produced from living plant tissues, we selected two experimental systems that have previously been shown to be sources of NO. We chose tobacco (Nicotiana benthamiana) leaves to demonstrate the reduction of nitrate to NO by NR because N. benthamiana leaf tissues have high levels of inducible NR (Yamamoto-Katou et al., 2006). Excised N. benthamiana leaves infiltrated with a buffered solution of 50 mM KNO3 were placed in the sample chamber of the oxidizer set-up immediately after excision. Air was passed over the tissue for 1 h at 40 ml min–1. Figure 3A shows that more than 50 nmol NO g–1 h–1 was produced by KNO3-treated leaves. Because leaf tissue was infiltrated with KNO3 buffered to pH 7.5, negligible amounts of HNO2 were produced.
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We have previously shown that aleurone tissue from barley grain can produce NO from added KNO2 (Bethke et al., 2004). We incubated ten barley aleurone layers for 24 h, removed the incubation medium to the sample chamber of the oxidizer column set-up and added 125 µM KNO2. Because the aleurone layer secretes organic acids that acidify the incubation medium to a pH of around 4, both HNO2 and NO were trapped (Figure 3B). After 30 min of incubation, 2.2% of the added KNO2 was trapped as HNO2 in the first Griess trap and 5.9% was captured in the second trap as a result of NO oxidation on the column. It resulted in the production of 5.6 and 15.0 nmol ml–1 h–1 of HNO2 and NO, respectively (Figure 3B). These values compare favorably with those obtained previously (Bethke et al., 2004).
Barley aleurone layers also synthesize NR in response to added nitrate (Ferrari and Varner, 1969, 1970). We used our oxidizer set-up to establish whether this tissue also produces NO from nitrate. Fifty-five aleurone layers were incubated in 50 mM KNO3 in the sample chamber and NO synthesis was monitored after 1 and 2 h (Figure 3C). After 2 h incubation with nitrate under aerobic conditions, aleurone layers produce 0.025 nmol NO layer–1 h–1.
NO Synthesis by Arabidopsis Mutants
We examined whether the Arabidopsis mutants Atnox1 (NO overproducing) and Atnos1 (NOS knockout) differ from wild-type plants in the amount of NO released to gas phase. Leaves were excised from wild-type, Atnox1 and Atnos1 plants and placed in the sample chamber of our oxidizer column set-up and NO production was measured for 1 h at a flow rate of 40 ml min–1. As the data in Figure 4 show, wild-type Arabidopsis leaves produce 0.4 nmol NO g–1 h–1, whereas nox1 plants produce almost twice as much. The effect of knocking out the AtNOS1 gene is very dramatic, as no NO production could be detected.
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Enhanced NO Synthesis by Tobacco Cell Suspensions Treated with Elicitor
We also tested the ability of our oxidizer column set-up to detect synthesis of NO from suspension cultured plant cells. We examined the response of suspension-cultured tobacco (N. tabacum) to the elicitor cryptogein. Twenty milliliters of suspension-cultured tobacco cells (cell density 0.1 g ml–1) were incubated in the sample chamber of a fish tank compressor-based set-up and an airflow of 40 ml min–1 was established for 1 h. Cells were mock treated with either H2O or 50 nM of cryptogein. Cryptogein brought about a 2.3-fold increase in NO production relative to mock-infected cells (Figure 5A).
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), cryptogein (
), representative data from three measurements are shown.We also compared NO production from these cells using the oxidizer column with data obtained using the NO-responsive dye DAF-FM. Samples (50 µl) withdrawn from the mock- or cryptogein-treated cell suspension were incubated with membrane-impermeable DAF-FM (1 µM) for 10 min, diluted and filtered immediately, and the fluorescence measured. Synthesis of NO in response to cryptogein increased rapidly during the first 100 min of incubation and thereafter reached a plateau (Figure 5B). Mock-treated cells showed low levels of NO synthesis, although there was evidence of a slight stimulation of NO within the first 60 min of incubation.
DAF-FM Fluorescence with NO is Time- and pH-Dependent
Oxygen-free, deionized water was sparged with NO to obtain a saturated solution that was then diluted to final concentrations of 190 nM NO and 380 nM NO in a reaction mixture containing 1 µM DAF-FM. The oxygen concentration in the reaction mixture was estimated to be 
260 µM based on the solubility of oxygen in water (Lide and Frederikse, 1995) and the volumes of the reaction mixture and air in the reaction vessel. The reaction of NO with DAF-FM was followed for up to 120 min (Figure 6A). For both concentrations of NO, relative fluorescence (493 nm excitation, 514 nm emission) increased rapidly during the first 10 min of incubation and reached a steady slow increase at 40 min. After 60 min, there was a minimal (15%) increase in fluorescence.
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), total fluorescence in the presence of NO (Relative fluorescence also changed when 1 µM DAF-FM was allowed to react with NO (300 nM) at varying pH (Figure 6B and 6C) and buffer concentrations (Figure 6C). With increasing pH from 5.5 to 9, there is no change in DAF-FM fluorescence, but there is gradual quenching of the fluorescence signal from DAF-FM plus NO, the degree of quenching being more pronounced above pH 7 (Figure 6B). NO-dependent fluorescence from 1 µM DAF-FM with 300 nM NO was higher in the presence of 2-(N-morpholino)ethanesulfonate (MES) at pH 5.8 in the reaction mixture than with 3-(N-morpholino)propanesulfonate (MOPS) or phosphate at pH 7.0 (Figure 6C) matching the data in Figure 6B. There was the same level (
0.05) of gradual quenching of fluorescence with all buffers as their concentration increased from 5 to 50 mM (Figure 6C). Quenching of DAF-FM fluorescence was not found in control samples without NO (data not shown).
Nitrite May Interfere with NO Measurement Using DAF-FM
We used two different concentrations of nitrite in order to test possible interference of nitrite with NO measurement using DAF-FM. Figure 7 shows that there was no significant increase in fluorescence from 1 µM DAF-FM at 0.5 µM nitrite in water (pH 5.6), but, as the pH was lowered below 2.5, there was a linear increase in fluorescence. At 100 µM nitrite, however, a linear increase in fluorescence was observed, with 1 µM DAF-FM in unbuffered water (pH 5.6).
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dashed line).The NO Scavengers PTIO and cPTIO Enhance DAF-FM Fluorescence In Vitro
PTIO and cPTIO are widely used scavengers of NO in biological experiments. We therefore examined the kinetics of DAF-FM fluorescence with NO in the presence of these scavengers. As Figure 8A shows, there is an increase in fluorescence from 1 µM DAF-FM in the presence of 190 nM NO and 380 nM NO, but this signal is not quenched in the presence of either 5 or 100 µM PTIO. Indeed, in the presence of 5 µM PTIO, there is a very rapid increase in fluorescence from DAF-FM and 380 nM NO so that the reaction goes to completion within 1 min and the signal is higher than that obtained in the absence of PTIO. Similar kinetics were observed when 100 µM PTIO was added to a reaction mix of 1 µM DAF-FM and 380 nM NO except that the fluorescence signal was lower than that obtained with 5 µM PTIO by about 30%. cPTIO also dramatically increased fluorescence from DAF-FM and NO in a similar manner (data not shown).
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Recently, cPTIO was shown to mask fluorescence of DAF in addition to enhancing the reaction of DAF with NO (Arita et al., 2006). To establish whether the lowering of the fluorescence signal from DAF-FM by higher concentrations of PTIO is due to fluorescence absorption, we allowed 1 µM DAF-FM to react with 380 nM NO for 1 h before adding 5–1000 µM PTIO (Figure 8B). Whereas 5 µM PTIO did not cause a decrease in fluorescence, 100 µM PTIO absorbed about 18% of initial fluorescence. On the other hand, if PTIO was present before addition of NO solution, 100 µM PTIO caused about a 30% decrease in fluorescence compared with fluorescence in the presence of 5 µM PTIO (Figure 8A).
Dead Cells Can Fluoresce Brightly when Loaded with DAF-FM Diacetate
Many cell types in plants are dead at functional maturity, such as the water conducting elements of the xylem and the starchy endosperm of cereal grains. We were therefore interested in the behavior of DAF-FM in tissues that had dead cells. Cereal aleurone layers that have both dead and living cells at functional maturity were used as an experimental system to test the ability of plant tissues to load DAF-FM diacetate (DAF-FM DA), the membrane permeant form of DAF-FM, and the in vivo fluorescence response of DAF-FM to NO. Aleurone layers loaded with DAF-FM DA were incubated in a sample chamber constructed on a microscope slide having an inlet and outlet that allowed the addition of appropriate dyes and NO donors.
Figure 9A–9D shows a series of images of the same aleurone layer viewed under different conditions of illumination and incubation. Figure 9A is a phase contrast image of an aleurone layer showing aleurone cells surrounded by a thick cell wall and thin-walled subaleurone cells adhering to the surface of the aleurone layer. Figure 9B shows an image of an aleurone layer loaded with DAF-FM DA for 15 min and observed by fluorescent light. Very low fluorescence from the cells of the aleurone layer is seen, confirming our previous observations with isolated aleurone protoplasts (Beligni et al., 2002). When an NO donor (500 µM diethylamine NONOate - DEANO) is added to the aleurone layer and the fluorescence image captured, angular cells fluoresce brightly (Figure 9C), but the thick-walled aleurone cells remain weakly fluorescent. The angular fluorescent cells in Figure 9C are non-living subaleurone cells and they can be seen faintly in the phase-contrast image (Figure 9A). When fluorescein diacetate (FDA) is added to the sample (Figure 9D), fluorescein accumulates in the cytosol and nucleus of aleurone cells, indicating that these are living cells, but there is no increase in fluorescence from subaleurone cells.
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The images shown in Figure 9D demonstrate that aleurone cells are alive because they can accumulate FDA and hydrolyze it to brightly fluorescent fluorescein. The weak fluorescence from DAF-FM DA (Figure 9B and 9C) suggests that these cells did not load the dye efficiently because they showed no fluorescence in the presence of the NO donor DEANO, even though the adjacent dead subaleurone cells were brightly fluorescent. We adopted several strategies to enhance the loading of DAF-FM DA into living aleurone cells. Prolonged incubation of the tissue with DAF-FM DA (up to 120 min), and the incorporation of antioxidants (butylated hydroxytoluene, dithiothreitol and ascorbate) to prevent formation of oxidized groups (e.g. aldehydes) which may immobilize DAF-FM DA because of binding to amino groups, failed to enhance dye uptake.
Plant tissues, especially secretory glands such as cereal aleurone, produce a spectrum of esterases and these enzymes (Fincher, 1989) could de-esterify DAF-FM DA. Schroeder and colleagues (Kuchitsu et al., 2002) have used eserine (an esterase inhibitor from plants) to inhibit the activity of extracellular esterases when loading esters of fluorescent dyes. We were unable to enhance the uptake of DAF-FM DA when eserine was added to the incubation medium, neither were we able to enhance dye uptake when the aleurone layers were pretreated with 10 mM HCl - a procedure also known to inhibit extracellular hydrolase activity in this tissue (Varner and Mense, 1976).
As a positive control for the above experiments, we used the abaxial epidermis of tobacco leaves treated with cryptogein. Both epidermal and stomatal guard cells of tobacco showed weak fluorescence after loading with DAF-FM DA (Figure 9F). That background fluorescence showed no time-dependent increase indicates that there was no synthesis of NO without cryptogein treatment. After treatment with cryptogein, the fluorescence was especially pronounced in stomatal guard cells and also increased in the cytoplasm, nuclei and plastids of epidermal cells (Figure 9G). A further increase in fluorescence of these cells was observed after addition of FDA, confirming that the cells that displayed a strong response to cryptogein were viable (Figure 9H).
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DISCUSSION |
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
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The Oxidizer Column-Based NO Detector
Using commonly available equipment and chemicals, we have constructed a simple bench-top detection system for the quantitation of NO in the gas phase. The system relies on the ability of strong oxidizing agents such as CrO3 to oxidize NO to NO2, and the formation of NO2– and NO3– from NO2 in aqueous solution. The Griess reagent is used to detect the presence of NO2– by the formation of an azo dye product having an absorption maximum of 545 nm. The system is capable of detecting HNO2 and NO2, but, because of the high solubility of NO2 in the aqueous phase and its low volatility, we conclude that our system has high specificity for HNO2.
Our NO detection system was designed to be of high sensitivity and the limit of detection ranges from 1 to 2 ppb of NO (equal 0.1–0.2 nmol h–1 under the above-mentioned conditions) and the response is linear up to 1 ppm NO. This sensitivity approaches that obtained with chemiluminescence (Byrnes et al., 1996) and laser photoacoustic (Mur et al., 2005) instruments. Chemiluminescence (Rockel et al., 2002; Planchet et al., 2005; Planchet and Kaiser, 2006; Planchet et al., 2006) and laser photoacoustic approaches (Leshem and Pinchasov, 2000; Mur et al., 2005) have been successfully used to detect NO emitted from plants, but the cost and availability of these instruments may be out of reach for many researchers. Our set-up is simple and sensitive and has the further advantage that it achieves spatial separation of NO detection from the biological material being sampled, to minimize interference with the detection of NO. One possible source of interference concerns the ability of biological material to readily volatilize NO. If experimental conditions are carefully monitored, such as by ensuring efficient gas exchange in the sample chamber and by selecting the appropriate biological material, problems with NO volatilization can be easily avoided.
When testing for the selectivity of the detector for NO, we found it to be insensitive to a range of other nitrogen-containing compounds, including ammonia, azide, and cyanide. Only nitrous oxide (N2O) elicited a positive signal at concentrations of around 1000 ppm. Though plants can produce N2O as a byproduct of NO3– assimilation, the production of N2O reported by others did not exceed 1 µmol g–1 h–1 (e.g. Dean and Harper, 1986; Smart and Bloom, 2001) (equivalent to 10 ppm under our experimental conditions) so that N2O is unlikely to interfere with NO measurement in our flow-through system.
Plant tissues produce background amounts of NO ranging from 0.05 to 0.3 nmol g–1 h–1 in sunflower leaves (Rockel et al., 2002) to about 1 nmol g–1 h–1 in tobacco leaves (Mur et al., 2005). Production of NO may be increased markedly by exposure to anoxia (up to 170 nmol g–1 h–1 in sunflower leaves (Rockel et al., 2002)), nitrate treatment (up to 10 µmol g–1 h–1 in soybean leaves (Harper, 1981; Klepper, 1987)), or by pathogen infection (40 nmol g–1 h–1 in tobacco leaves (Mur et al., 2005)). Clearly, based on this published information, our oxidizer column set-up is well suited for the quantitation of NO production from plants under a wide range of conditions.
A proof-of-concept series of experiments were carried out to test the functionality of our NO detector. They included experiments with tobacco leaves, cereal aleurone layers, Arabidopsis mutants compromised in their ability to make NO, and tobacco cell suspension cultures. Leaves of tobacco (Yamamoto-Katou et al., 2006) and soybean (Harper, 1981; Klepper, 1987) contain high levels of nitrate-inducible NR, and, in the case of soybean, NR activity leads to NO production. KNO3-treated leaves of N. benthamiana synthesized more than 50 nmol NO g–1 h–1 - an amount that was readily detected by our set-up.
Barley aleurone layers also produce NO when treated with nitrite and we confirmed the data of Bethke et al. (2004) that up to 15 nmol NO ml–1 h–1 was produced when aleurone incubation medium was incubated with 125 µM nitrite. HNO2 is also detected in the first trap of our set-up when nitrite is added to aleurone medium. HNO2 is formed by the protonation of NO2– and, since it is volatile, it is found in the first trap (Figure 1). The synthesis of NO in the aleurone system has been shown to result from reduction of nitrite at low pH. The reductants for this reaction are likely the procyanidins that are present in the testa that adheres to the cereal aleurone and which are released into the incubation medium. Combined with the low pH of the aleurone incubation medium (pH 3–4) achieved by the release of organic acids (Macnicol and Jacobsen, 1992; Drozdowicz and Jones, 1995), conditions are reached that lead to the formation of HNO2 and thus NO (Scheme 2). The synthesis of NO from nitrite is now believed to be of widespread biological importance, especially in mammals. Indeed, it is thought that nitrite may be the largest storage form of NO in animal tissues (Gladwin et al., 2006).
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Barley aleurone layers also have nitrate-inducible NR activity and, using our set-up, we have shown that NR is also a likely source of NO in this tissue. The ability of nitrate to induce NR synthesis in aleurone layers was demonstrated by Ferrari and Varner (1969, 1970). Interestingly, their assay for NR relied on the release of nitrite from KNO3-treated aleurone layers incubated in ethanol under anoxic conditions. Under their experimental conditions, NR was induced under aerobic conditions, but, when layers were transferred to anoxia and ethanol, nitrite was released. Nitrite is not released under aerobic conditions; rather, as we show, NO is one of the products made in response to nitrate when layers are incubated in air. The observation that release of nitrite from aleurone layers occurs under anoxia is interesting in the context of NO synthesis, because the aleurone apoplast provides conditions for the formation of HNO2 and NO from nitrite.
Having established that NO produced by NR or non-enzymatically could be detected with our set-up, we turned our attention to its synthesis by other sources. Arabidopsis NOS (AtNOS1) has been shown to be required for the synthesis of NO and Atnos1 mutants are impaired in their ability to synthesize NO (Crawford and Guo, 2005; Crawford et al., 2006). Recently, doubts have been cast about the identification of AtNOS1 as a bona fide nitric oxide synthase gene because AtNOS1 protein has not been proven to synthesize NO in vitro (Zemojtel et al., 2006). In addition to Atnos1 mutant, we have included Atnox1 in our experiments, as this mutant was shown to overproduce NO (He et al., 2004). Using the oxidizer column-based NO detector, we were able to discriminate the NO synthesis among wild-type, Atnox1, and Atnos1 plants. Our experiments do not help to resolve the dilemma about the actual role of AtNOS1 in NO synthesis. We merely show that used Arabidopsis mutants are of the expected phenotype with regard to release of NO to the gas phase.
There is growing evidence that NO synthesis is altered during plant pathogen interactions (Klessig et al., 2000; Delledonne et al., 2002; Wendehenne et al., 2004; Mur et al., 2006). We tested the response of the fungal elicitor cryptogein (Blein et al., 1991) in suspension-cultured tobacco (N. tabacum cv Xanthi) cells. We chose this experimental system because it would allow us to test whether NO synthesis by suspension-cultured cells could be detected with our system, while simultaneously asking about the response of tobacco cells to cryptogein.
A rapid burst of NO production in response to cryptogein treatment was demonstrated using fluorescent probes (Lamotte et al., 2004) - a result that has been questioned, as sampling of the gas phase above the cell suspension with a chemiluminescence detector showed no detectable NO synthesis 2 h after cryptogein addition (Planchet and Kaiser, 2006; Planchet et al., 2006). We show NO synthesis by cryptogein-treated suspension-cultured cells of tobacco using two methods. Both the oxidizer set-up and DAF-FM showed an increase in NO production. Both methods showed low background NO production by suspension-cultured cells and this increased significantly when cells were exposed to the elicitor. In addition to confirming previous work on the effects of cryptogein on the hypersensitive response in tobacco (Foissner et al., 2000; Lamotte et al., 2004), the data from the oxidizer column set-up confirm that this system can be successfully used to measure NO synthesis by suspension-cultured cells.
DAF-FM-Based NO Detection
Because DAF is widely used to localize NO in plant tissues, we examined several of the parameters of the dye's response. We show that the reaction between DAF-FM and NO are both time- and pH-dependent - observations that have implications when NO production by plant tissues is being measured.
The reaction of NO at a concentration of 100 nM with 1 µM DAF-FM in water was found to take tens of minutes to go to completion in a simplified chemical system (Figure 6A). The oxidation of NO (Scheme 1) is the likely limiting step for the formation of fluorescent DAF-FM triazole (DAF-FM-T) (Kojima et al., 1998). This process is a third-order reaction (Mayer and Hemmens, 1997) and explains why it can be expected to be very slow at nanomolar NO concentrations. This chemistry may strongly affect real-time detection of NO. Even if the amount of NO produced is constant, it may take some time for the reaction with DAF-FM to be completed.
The formation of DAF-FM-T is also pH-dependent (Figure 6B and 6C), having substantially lower yield above pH 7 (Figure 6B). Dinitrogen trioxide (N2O3), which nitrosates DAF-FM (Scheme 1), is sensitive to nucleophilic attack (Lancaster Jr, 2003) and this may explain why water, and especially hydroxyl anions which are regarded to be stronger nucleophilic agents, compete with DAF-FM in the nitrosation reaction (summarized in Scheme 2).
Further, there was a decrease in NO-dependent fluorescence of DAF-FM when buffer concentration was increased in the reaction mixture, regardless of the buffer used (Figure 6C). Considering that the concentration of MOPS buffer (10–100 mM) used to dilute the reaction mixture prior to fluorescence measurement did not affect the final fluorescence, quenching is likely a result of an increase in the ionic strength of the reaction mixture.
Many processes in plants are accompanied by changes in the pH of the apoplast and cytosol, such as sucrose and ion uptake from the apoplast, acidification of the wall during elongation growth, the hypersensitive reaction, etc. (Buchanan et al., 2002). When measuring NO production by DAF-FM under conditions in which the pH of the sample is likely to change, the detection reaction should preferably be carried out in an environment in which the pH can be controlled. The shift of pH often cannot be avoided during in vivo experiments; therefore, the effects of fluctuating pH must be taken into account when interpreting the experimental data.
We have also focused on the interaction of nitrite with DAF-FM, as it is a potential source of NO in plants (Yamasaki, 2000; Bethke et al., 2004) and a source of nitrosating agents (N2O3 and NO+) (e.g. Grossi and Montevecchi, 2002). Plants can contain high concentrations of nitrite (around 10 µM), which can increase to around 1–5 mM under anoxic conditions (Rockel et al., 2002) or following application of N fertilizers to soil. A fluorescent product can only be formed from 1 µM DAF-FM and 0.5 µM nitrite (a level which can originate from NO oxidation in vivo) when the pH of the reaction mixture is below 2.5; however, at higher nitrite concentrations (100 µM NaNO2), a fluorescence signal can be obtained in unbuffered water (pH 5.6) (Figure 7). Despite the low pKa of nitrous acid (3.2), about 0.5 % of the total nitrite is in the form of nitrous acid in solution under these conditions. Nitrous acid may produce N2O3 (Grossi and Montevecchi, 2002) to nitrosate DAF-FM. Furthermore, the presence of reducing compounds in plant tissues may result in NO production from nitrous acid (Yamasaki, 2000; Bethke et al., 2004), as summarized in Scheme 2. The data in Figure 7 and the work of others imply that even under physiological conditions, in which the extracellular space in plants has a pH in the range of 4–5.5, and in which nitrite concentrations are often at high micromolar level, nitrite may give rise directly to the formation of NO or it may induce nitrosation of DAF-FM via N2O3 without presence of NO.
Scavenging molecules, such as PTIO and cPTIO, have been widely used to help confirm the production of NO and its localization in plant cells and tissues (Neill et al., 2002; Guo and Crawford, 2005; Gomes et al., 2006), and to establish the involvement of NO in biological responses (Beligni et al., 2002; Garcia-Mata and Lamattina, 2002; Pagnussat et al., 2002; Correa-Aragunde et al., 2004; Creus et al., 2005; Tun et al., 2006). Because of the widespread use of PTIO and cPTIO in biology, we examined the kinetics of DAF-FM fluorescence with NO in the presence of these scavengers. Other researchers have shown that PTIO and cPTIO enhance fluorescence of DAF when it was combined with NO donors (Espey et al., 2001; Arita et al., 2006); these data should be interpreted with caution because of the continuous decomposition of the NO donor in the reaction mixture. We therefore chose to study the kinetics of DAF-FM in a system with pure NO. We show that at concentrations of PTIO (cPTIO) used in this work, the reaction with NO went to completion in 1 min in accordance with the rate of NO oxidation by PTIO (cPTIO) (Akaike et al., 1993). The unexpected increase in DAF-FM fluorescence in the presence of NO and PTIO (cPTIO) at concentrations of 100 µM may be a cause for concern. PTIO and cPTIO in the range of 100 µM are commonly regarded to be effective for NO removal in vivo from plant tissues (Beligni et al., 2002; Garcia-Mata and Lamattina, 2002; Gould et al., 2003; Guo and Crawford, 2005). However, the intrinsic oxidation of NO in vivo is likely to be faster in the presence of biological compounds that can oxidize NO, such as heme proteins (Wendehenne et al., 2001), compared with our well defined in vitro experimental system. There is likely a concerted action of PTIO or cPTIO and biological oxidants in vivo that strongly favors the conversion of NO to NO2, lowering the amount of N2O3 formed and causing a low rate of nitrosation of DAF-FM, leading to a negligible increase in fluorescence.
The enhancement of the conversion of DAF to fluorescent DAF triazole (DAF-T) in vitro was ascribed by Arita et al. (2006) as being the result of faster conversion of NO to NO2 and subsequently N2O3 by cPTIO than from the oxidation of NO with oxygen (Miles et al., 1996). Yamasaki and colleagues (Arita et al., 2006) also showed that high cPTIO concentrations (5 mM) mask the fluorescence of DAF-T. Our results demonstrate experimentally that absorption of the fluorescence signal of DAF-FM-T by PTIO concentrations in the range of 50–1000 µM does not fully explain PTIO action. The concerted action of NO oxidation and scavenging should lead to a faster decrease in DAF-FM-T formation according to kinetic calculations based on the data of Goldstein et al. (2003) by increasing PTIO concentrations more than the observed fluorescence decrease. We therefore favor the reaction mechanism postulated by Goldstein et al. (2003). They propose that, in vitro, PTIO reacts efficiently with NO2. The resulting PTIO+ likely rapidly oxidizes DAF-FM to DAF-FM free radical, which can subsequently couple readily with NO to form the nitrosation product (Jourd'heuil, 2002). The reactions that NO, PTIO, or cPTIO and DAF-FM undergo are summarized in Scheme 2.
To investigate the interaction of DAF-FM with dead cells, the barley aleurone layer - a well defined model system consisting of living cells and adhering dead sub-aleurone - was used. When treated with the NO donor DEANO, a rapid increase in fluorescence from dead subaleurone cells was observed in DAF-FM DA-loaded aleurone layers (Figure 9C). These observations serve to indicate an important principle about the use of DAF-FM DA and point to pitfalls in its application in plants, as dead cells can bind DAF-FM. Second, the observation that dead cells fluoresce in the presence of DAF-FM DA implies that dye was likely hydrolyzed by extracellular esterases, because DAF-FM DA does not react with NO. Surprisingly, the fluorescence of living aleurone cells did not increase with DEANO treatment, even when eserine or HCl was applied to inactivate extracellular hydrolases. The weak fluorescence from living aleurone cells suggests either that they take up DAF-FM DA very poorly, do not hydrolyze the dye to the NO-reactive form, or that the dye is modified in the cytosol to a form which does not bind NO.
Mature plant tissues consist of a mix of living and non-living cells, including the water-conducting elements of the xylem (Buchanan et al., 2002). It is perhaps not surprising that some researchers have reported that these non-living tissues of plants appear to be a source of NO (Gabaldón et al., 2005). These data may reflect the non-enzymatic synthesis of NO in the apoplast of plant cells (Yamasaki, 2000; Bethke et al., 2004) and the ability of the dye to adsorb onto dead cells.
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TOPAbstractINTRODUCTIONRESULTSDISCUSSIONMETHODS |
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Chemicals
DAF-FM, DAF-FM DA, DEANO, and cPTIO were purchased from Molecular Probes (Eugene OR, USA). PTIO was obtained from MP Biomedicals (Illkirch, France). Stock solution of DAF-FM in dimethylsufoxide was kept at –20°C. Aliquots of stock solution of DEANO in water were kept at –80°C. Working solution was prepared immediately before use and kept in the dark on ice for not longer than 1 h. Cryptogein was a generous gift from Prof. Vladimir Mikes (Dept of Biochemistry, Faculty of Science, Masaryk University, Brno, Czech Republic). Griess reagent (cat No. LC13280–1; 4% sulfanilamide, 0.2% N-(1-naphthyl)-ethylenediamine, 10% H3PO4) was purchased from LabChem Inc. (Pittsburgh PA, USA). N2O was prepared according to Mattson et al. (2002).
Oxidizer Column Preparation
Column packing was prepared using a modification of the procedure described by Klepper (1979): 100 ml of 3-mm Pyrex glass beads were coated with 4 ml of a solution containing 4 g of potassium dichromate and 1.5 ml of 98% sulfuric acid dissolved in 25 ml of water. Coated beads were dried at 80°C until the color changed from yellow to brick-red. Dried beads were packed into a glass column (interior cross-section: 1.5 cm, length 30 cm), stoppered with glass wool plugs and rubber stoppers (containing an injection needle as an input or output for gas) on each end.
Flow-Through Detector
Two spargers, each made of 15-ml glass test tube, stoppered with a two-holed rubber bung with Peek pipes (gas inlet and outlet) containing 1 ml of Griess reagent, were connected to the oxidizer column by PVC tubing so that the first sparger was placed before the oxidizer column and the second one after the column. Air was humidified with a sparger in H2O placed immediately before the sample chamber. Airflow was controlled using gas tanks and computer-operated mass flow controllers (Libourel et al., 2006). Where indicated, a fish tank compressor was also used to pump air through the system. A ‘T’-pipe was connected to the fish tank compressor in order to split the air flow between the flow-through system and the outlet tube. A pressure meter was inserted immediately after the T-junction on the flow-through system branch. A pinchcock on the outlet tube allowed setting of the pressure. A constant flow rate (±10%) monitored at the end of the flow-through system was achieved by maintaining appropriate pressure (10–20 kPa).
Before operation, each oxidizer column was purged with moist air at a flow rate of 40 ml min–1 for 5 min. To measure NO production, sampling of the gas was carried out for 10–60 min in order to achieve reasonable color intensity of the Griess reagent in the traps. After disconnecting the sample chamber, at least 100 ml of pure moist air was passed through the detector in order to trap the amount of nitrogen oxides remaining in the dead volume of the apparatus. The absorbance of the Griess reagent was assayed at 545 nm after 10 min (Griess reagent needs about 10 min in order to develop full color intensity). A calibration curve of nitrogen equivalents was made using nitrite as standard. Background response of each column was carefully monitored and subtracted from the response of the sample.
The system was tested using NO (212 or 198 ppm in nitrogen; Praxair, San Ramon CA, USA) dilutions in nitrogen (99.999%; Praxair) mixed with inlet air at the desired flow rate prior to entering the first trap of the flow detector (for technical details, see Libourel et al., 2006). In experiments in which a fish tank compressor was used to pump air through the system, a defined amount of NO at a concentration of 
1 ppm in a rubber-plug-stoppered flask was utilized. This test flask was placed into the system instead of a sample chamber and airflow equal to at least ten times that of the volume of the flask increased by 100 ml necessary for purging of the dead volume (see above) was pumped.
KNO3 Treatment of Tobacco Leaves
Tobacco (N. benthamiana) was grown in a growth chamber (25°C, relative humidity 80%, 16-h photoperiod). The youngest fully expanded leaves of 7 week old tobacco plants were used for the experiments. Excised leaves were weighed and syringe-infiltrated with a solution of 50 mM KNO3 buffered with 100 mM potassium phosphate (pH 7.5) containing 1% N-butanol (Nicholas et al., 1976) and immediately placed in the sample chamber (kept in the dark at 25°C) of the oxidizer set-up. Air was passed over the leaf for 1 h at 40 ml min–1.
Production of NO by Aleurone Layer Medium
Barley (Hordeum vulgare cv. Himalaya) aleurone layers were prepared according to Beligni et al. (2002). Ten isolated aleurone layers were incubated in 3 ml of 10 mM CaCl2 with shaking at 60 rpm in darkness at 25°C for 24 h. Two milliliters of the incubation medium was placed in a test tube, kept in the dark at 25°C, and placed between the humidifier and the first trap of the oxidizer set-up. The tip of the gas inlet pipe was inserted below the liquid surface in the test tube. Potassium nitrite was added to a final concentration of 125 µM and air was passed through the sample at 40 ml min–1 for 30 min.
Production of NO by Aleurone Layers
Fifty-five barley aleurone layers prepared as above were placed into 10 ml of 10 mM CaCl2 buffered with 2 mM acetate to pH 4.8 in a 50-ml Ehrlenmayer flask placed on a shaker (200 rpm). This flask was capped with a silicon stopper containing inlet and outlet injection needles and connected prior to the first trap of the oxidizer set-up. After determination of background production of NO, KNO3 was added to the final concentration of 50 mM. Gas sampling was carried out at flow rate of 40 ml min–1 every 45 min for 15 min.
Detection of NO from Arabidopsis Leaves
Arabidopsis plants: Col 0, Atnox1 (NO overproducing (He et al., 2004)) and Atnos1 (NOS knockout (Guo et al., 2003; Crawford et al., 2006)) were grown in a climatized chamber. Cultivation conditions were set as follows: 16-h light period, light intensity 130–150 µE m–2 s–1, temperature 22°C during light period and 20°C during dark period and relative humidity 70–75%. Freshly excised Arabidopsis leaves (0.5 g) were placed into the sample chamber of the oxidizer detector. The sample chamber was kept in ambient light at 25°C. Air was passed over the leaf sample at 40 ml min–1 for 1 h.
Synthesis of NO by Tobacco Cell Suspension
Tobacco (N. tabacum cv Xanthi) suspension-cultured cells were grown in Chandler's medium (Chandler et al., 1972) on a rotary shaker (120 min–1) at 25°C and 12-h light period. Sub-cultivation was carried out once a week, when cells were transferred into fresh medium. Suspension density was set to 0.1 g ml–1. Twenty milliliters of the cell suspension were put into a 50-ml Ehrlenmayer flask placed on the rotary shaker (120 min–1, ambient light, 25°C). Cryptogein was added to a final concentration of 50 nM after a 2-h pre-incubation period. An equivalent amount of water was used as a mock treatment. The flask was stoppered with a silicone plug and mounted into the flow-through set-up. Air passed over the shaken cell suspension at 40 ml min–1 for 1 h.
Nitric Oxide Solutions
Double-distilled water was boiled and purged with N2 (30 ml min–1) for at least 30 min. Five milliliters of oxygen-free water were saturated with NO gas (212 or 198 ppm in nitrogen; Praxair, San Ramon CA, USA) at a flow rate of 15 ml min–1 for 15 min. The NO solution was kept in a tightly closed container. Calculations based on NO solubility in water (Koppenol, 1998) indicate that the saturated solution (23°C, atmospheric pressure) contains 422 or 394 nM NO.
Nitric Oxide Detection with DAF-FM
All reactions were carried out in tightly closed 1.5-ml Eppendorf tubes. Ten-times concentrated working stock solution (10–100 µM) of DAF-FM was prepared by dilution of the dimethylsulfoxide stock solution (1 mM) with water. Where desired, ten-times concentrated buffer was added. The reaction was started by addition of NO or nitrite solution to the desired final concentration. The total volume of the reaction mixture was 100 µl. For determination of reaction kinetics, the total volume was increased to 1 ml in order to ensure enough samples for fluorescence measurement. After incubation in darkness at 25°C, the reaction mixture was diluted 12.5–25 times with 10 mM MOPS buffer, pH 7.0 (or 100 mM where necessary to buffer highly acidic reaction mixtures; concentration of MOPS from 10 to 100 mM did not influence the fluorescence response). Fluorescence was measured using either a Hitachi F4500 spectrofluorimeter (Hitachi High Technologies America, San Jose CA, USA) at excitation and emission wavelengths of 493 and 514 nm, respectively, or an RF 551 Shimadzu spectrofluorimeter (Shimadzu Europe, Duisburg, Germany) at excitation and emission wavelengths of 498 and 517 nm, respectively. The measured response was converted to relative values. The highest value measured in a particular experiment was taken as 100%. That value ranged from 2500 to 4400 arbitrary units with the Hitachi F4500 and from 140 to 1000 arbitrary units with the Shimadzu RF 551.
Nitric Oxide Detection In Vivo
The preparation of tobacco (N. tabacum cv Xanthi, 7 week old plants grown at 25°C, 80% humidity and 16-h light period) abaxial epidermis and the detection of cryptogein-induced NO burst was carried out as described (Foissner et al., 2000). Unless otherwise indicated, ten barley aleurone layers (see above for preparation) were incubated in 3 ml of 10 mM CaCl2 solution containing 5 µM gibberellic acid in a 25-ml Ehrlenmayer flask with shaking at 60 rpm for 6 h. Where indicated, antioxidants (butylated hydroxytoluene, dithiothreitol, and ascorbate) were added to the incubation solution to a final concentration of 1 mM. After incubation, aleurone layers were cut into halves and transferred into a dye loading solution containing 10 mM CaCl2 and 10 µM DAF-FM DA. Where indicated, the esterase inhibitor eserine was added into the loading solution to a final concentration of 300 µM, or dye loading was carried out in solution without CaCl2 but containing 10 mM EGTA.
DAF-FM DA loading of tissue was carried out for 5–120 min. For aleurone layers, loaded tissue was transferred to a chamber assembled on a microscope slide that allowed addition of different solution to the tissue. Images were taken using an Axiophot 373 (Zeiss, Thornwood, NY, USA) microscope equipped with a standard fluorescein optical filter set and captured using a CCD camera (MicroPublisher 5MPix color camera, QImaging, Burnaby, BC Canada).
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Acknowledgements |
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We thank David King for use of the Shimadzu spectrofluorimeter, Dr Jianzhong Liu and Dr Barbara Baker for providing us with Arabidopsis mutants, Martina J
zová for excellent technical assistance and the staff of the Bioimaging Center at UC Berkeley. The authors gratefully acknowledge the US National Science Foundation, the International Programs Division of NSF, the University of California College of Natural Resources, and the Czech Ministry of Education (project 1P05ME757) for their generous support. No conflict of interest declared.
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