JXB Advance Access originally published online on September 24, 2004
Journal of Experimental Botany 2004 55(408):2523-2531; doi:10.1093/jxb/erh266
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RESEARCH PAPER |
Oxygen radicals produced by plant plasma membranes: an EPR spin-trap study
Mojovi
12
i1
eljko Vuini3,*
1Faculty of Physical Chemistry, University of Belgrade, Studentski trg 12–16, YU-11000 Belgrade, Yugoslavia (Serbia and Montenegro)
2Maize Research Institute ‘Zemun Polje’, POB 89–Zemun, YU-11081 Belgrade, Yugoslavia (Serbia and Montenegro)
3Center for Multidisciplinary Studies of the Belgrade University, Ul. Kneza Vieslava 1a, POB 33, YU-11030 Belgrade, Yugoslavia (Serbia and Montenegro)
* To whom correspondence should be addressed. Fax: +381 11 3055 289. E-mail: vucinic{at}ibiss.bg.ac.yu
Received 5 June 2004; Accepted 25 July 2004
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Abstract |
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Plant plasma membranes are known to produce superoxide radicals, while the production of the hydroxyl radical, previously detected in complex plant tissues, is thought to occur in the cell wall. The mechanism of production of superoxide radicals by plant plasma membranes is, however, under dispute. It is shown, using electron paramagnetic resonance spectroscopy with a 5-diethoxyphosphoryl-5-methyl-1-pyrroline N-oxide spin-trap capable of differentiating between radical species, that isolated purified plasma membranes from maize roots produce hydroxyl radicals besides superoxide radicals. The results argue in favour of superoxide production through an oxygen and diphenylene iodonium-sensitive, NADH-dependent superoxide synthase mechanism, as well as through other unidentified mechanism(s). The hydroxyl radical is produced by an oxygen-insensitive, NADH-stimulated mechanism, which is enhanced in membranes in which the superoxide synthase is incapacitated by substrate removal or inhibition.
Key words: DEPMPO, diphenylene iodonium, EPR, hydroxyl, KCN, maize, plasma membranes, spin-trap, superoxide
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Introduction |
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The initial steps in many of the processes in plants, such as signalling, nutrient uptake, stress response, growth, and development, are achieved via activated oxygen and other radical species, some of them accompanied by an externally measurable oxidative burst considered to be a component of the plant defence responses (Wojtaszek, 1997
; Levine, 2002; Vianello and Macrí, 1989). The production of extracellular activated oxygen species during such an oxidative burst in plants may be mediated by the activity of enzymes located in the plasma membrane. The involvement of a plasma membrane oxidase, catalysing the reduction of molecular oxygen to superoxide at the expense of NAD(P)H, similar to the inducible NADPH oxidase in the plasma membrane of mammalian phagocytes (Segal and Abo, 1993), was observed as an early response to pathogen attack (Wojtaszek, 1997). The second mechanism for apoplastic 
production in plants, which can be discriminated from the phagocyte-type oxidase by high sensitivity to inhibition by KCN, is provided by the NAD(P)H oxidizing activity of peroxidase localized in the cell wall (or at the plasma membrane) (Askerlund et al., 1987). The extremely reactive •OH radical can be generated in a Fenton reaction when and H2O2 are available. However, it has recently been demonstrated that •OH radicals are released by maize coleoptiles, and that this is mediated by an enzymatic reaction, suggested to be a cell wall peroxidase (Liszkay et al., 2003).
Studies of purified plasma membranes demonstrated the presence of NAD(P)H-superoxide synthase activity in elicitor-treated rose cells (Auh and Murphy, 1995) as well as in non-treated plants in rose cells (Murphy and Auh, 1996), bean hypocotyls (Van Gestellen et al., 1997), and maize roots (Vuleti et al., 2003a), with characteristics similar to the neutrophil NADPH oxidase. Diphenylene iodonium (DPI) has been used frequently as an inhibitor of the membrane-bound NAD(P)H oxidase, although different degrees of inhibition and different concentrations were effective in the case of plasma membranes obtained from different plant species (Bérczi and Møller, 2000; Murphy et al., 2000). On the other hand, the involvement of plasma membrane peroxidase activity in active oxygen species generation was demonstrated in plasma membrane preparations from cauliflower inflorescences (Askerlund et al., 1987), radish plasma membrane (Vianello and Macrí, 1989), and wheat plasma membrane (Qui et al., 1995). Thus, it has not been unequivocally determined whether superoxide is produced by the superoxide synthase or peroxidase(s). There are no reports, that the authors know of, which show the production •OH radicals by isolated plant plasma membranes.
Most of the measurements of radicals were performed by coupling them to coloured or chemiluminescent compounds, for example, during the nitroblue tetrazolium reduction or the stimulation of lucigenin luminescence. However, such methods suffer from questionable specificity and difficult quantification. Comparative methodological studies (Murphy et al., 1998) demonstrated that test reactions commonly used for the assay of production by plant cells or their components can lead to divergent results, even when applied to the same experimental system. Such shortcomings can be bypassed by using EPR spin-trapping, a technique based on a reaction in which the transient radical species reacts with specific nitrone or nitroso spin-traps to yield more persistent nitroxide spin adducts. These are readily detected by EPR spectroscopy qualitatively, whereby different species can be distinguished, and with great sensitivity (EPR detection of the superoxide radical with DEPMPO is 40 times more sensitive than spectrophotometric analysis with cytochrome c) (Roubaud et al., 1997). It has been applied in numerous studies on animal systems and rarely in plants. The authors are aware of only one report dealing with isolated plant plasma membranes (Qui et al., 1995), where a spin-trap of questionable sensitivity and specificity has been used to show that plant plasma membranes do produce the superoxide radical.
In this report, the production of oxygen radical species by isolated plasma membranes from maize roots was studied using the recently developed DEPMPO spin-trap (Frejaville et al., 1995). DEPMPO is capable of differentiating between different oxygen radical species ( and •OH) and adducts are far more stable than adducts to the commonly used spin-trap DMPO, and its use has already become more frequent in a relatively short time (Vasquez-Vivar et al., 1998; Chamulitrat, 1999; Stolze et al., 2000) even under in vivo conditions (Liu et al., 1999). It is demonstrated that both hydroxyl and superoxide radicals are being produced by plant plasma membranes through the participation of different enzymatic mechanisms. This was partly enabled by measuring oxygen-sensitive plasma membrane systems in oxygen permeable tubing instead of quartz capillaries or flat cells, an important improvement in trapping oxygen radicals in systems which consume oxygen.
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Materials and methods |
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Plasma membrane isolation
Inbred line VA35 of maize (Zea mays L.) was used. The seeds, germinated for 3 d on water, were grown for 14 d on modified Knopp solution (Hoagland and Arnon, 1950
), the concentrations of 
and 
being 10.9 mM and 7.2 mM. The plants were grown in a controlled environment, illuminated with 190 µmol m–2 s–1 photosynthetically active radiation, under a 12 h light/dark regime at 22/18 °C and a relative humidity of 70%. Plasma membrane isolation was performed by partitioning in a two-phase system as described by Larsson (1985). Cut roots were ground in cold grinding buffer (250 mM sucrose, 3 mM EDTA, 50 mM TRIS-HCl, pH 7.5, 1 mM DTT, and 10% w/v glycerol) with a chilled mortar and pestle. The microsomal fraction obtained from the homogenate (10 min at 12 000 g and 30 min at 100 000 g) was washed and suspended in phase buffer (5 mM K-phosphate buffer, pH 7.8, 330 mM sucrose, and 3 mM KCl) for plasma membrane purification by three steps of phase partitioning in a two-phase system [6.5% (w/w) dextran T 500, 6.5% (w/v) polyethylene glycol 335, 330 mM sucrose, 3 mM KCl, 5 mM K2HPO4, pH 7.8]. The final upper phase was diluted (2 mM TRIS-HCl, pH 7.5, and 250 mM sucrose) and centrifuged, the resulting pellet, containing purified plasma membranes, was resuspended in the same buffer to give a final concentration of 3–4 mg protein ml–1 and stored at –60 °C.
Sample preparation
Spin-trap DEPMPO (5-diethoxyphosphoryl-5-methyl-1-pyrroline N-oxide) was purchased from Alexis Biochemical, Lausen, Switzerland. It has been used without purification since the control EPR spectra of buffer solutions with NADH showed no appreciable EPR signal. DPI and SOD (from bovine erythrocytes) was purchased from Sigma, and NADH from BDH, only freshly prepared solutions being used to avoid auto-oxidation. Authentic DEPMPO/OH and DEPMPO/OOH spin adducts were generated by hydroxyl and superoxide generating systems, respectively. Hydroxyl radicals were generated in a Fenton reaction system consisting of 0.5 mM H2O2 and 75 µM Fe2+, the stock solution of FeCl2 being prepurged with N2 to ensure that only Fe2+ was present in the reaction system. Superoxide radicals were generated in a hypoxanthine/xanthine-oxidase system (HX/XO) consisting of 0.4 M hypoxanthine and 0.4 iu ml–1 xanthine oxidase (Sigma) dissolved in a 50 mM HEPES buffer (Frejaville et al., 1995). Standard membrane sample preparation consisted of 2.8 mg protein ml–1 of membranes, 3.3 mM NADH, 240 mM DEPMPO in 50 mM HEPES buffer, pH 7.5. All preparations, except the Fenton system, included 100 µM of chelating agent DETAPAC (Pou et al., 1994) to ensure that the trace metal impurities (especially iron) were removed. All samples were prepared under atmospheric conditions except where specified otherwise. The results presented were obtained from four isolations, with three independent measurements for a treatment.
EPR measurement
Samples were drawn into a 10 cm long gas-permeable Teflon tube (wall thickness 0.025 mm and i.d. 0.6 mm; Zeus industries, Raritan, NJ) and folded into 2.5 cm long segments to improve the signal-to-noise ratio (Swartz et al., 1986). EPR spectra were recorded at room temperature using a Varian E104-A EPR spectrometer operating at X-band (9.3 GHz) using the following settings: modulation amplitude, 2 G; modulation frequency, 100 kHz; microwave power, 10 mW; scan range, 200 G; scan time, 1–4 min. Spectra were recorded and analysed using EW software (Scientific Software). Spectral simulations were performed using WINEPR SimFonia (Bruker Analytische Messtechnik GmbH).
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Results |
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Figure 1 shows EPR spectra of DEPMPO spin adducts obtained in radical generating systems (Fenton and HX/XO). EPR spectrum of an adduct from the Fenton reaction (Fig. 1A) is typical for a DEPMPO/OH adduct (Frejaville et al., 1995
; Vasquez-Vivar et al., 2000). It also contains some traces of an additional radical (most probably DEPMPO/H adduct) barely visible at the wings of the spectrum, a detailed analysis of this adduct, however, being beyond the scope of this publication. The EPR spectrum of an adduct from the HX/XO reaction (Fig. 1B) is a typical 12-line spectrum of the DEPMPO/OOH adduct (Frejaville et al., 1995; Vasquez-Vivar et al., 2000), which can easily be simulated using a routinely employed set of parameters (Vasquez-Vivar et al., 1997). However, even in Fig. 1B traces of the DEPMPO/OH adduct can be discerned, and the spectrum in Fig. 1C cannot be simulated if only the DEPMPO/OOH exists, demonstrating the transformation of the OOH into the OH adduct. Although it has been claimed that such a transformation does not occur (Frejaville et al., 1995; Vasquez-Vivar et al., 1999), experimental results in several papers suggests the existence of this transformation (Frejaville et al., 1995; Chamulitrat, 1999; Vasquez-Vivar et al., 2000). The problem probably lies in various experimental procedures. Most measurements have been performed immediately after the initiation of the reaction, or in the subsequent 10 min, when the relative participation of OH adduct is small (compare Fig. 1B and C). The kinetics recorded over a longer period of time (Fig. 2) shows that the transformation does occur (as in the case of DMPO), but at a much slower rate.
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(1)=0.85, aH
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Figure 3A and B show EPR spectra of radicals produced by membranes in the presence of NADH and trapped by DEPMPO. A comparison of Fig. 3A and B shows that the same type of spectrum is observed, only the number of radicals increases with time. Spectra indicate to more than one type of spin adduct. Figure 3C shows that all the principal characteristics of adducts that DEPMPO has trapped in the membrane can be obtained by simple addition of the spectra obtained in radical generating systems (Fig. 1A, B). For all practical purposes, the most important feature of the spectra is the central four lines where the inner doublets originate from OH, and the outer from OOH (Chamulitrat, 1999
). This could indicate that the membrane produces these two radicals, but one should be careful in interpreting the spectra in Fig. 3A and B. The spectra in Fig. 1 show that the DEPMPO/OH adduct can be obtained by transformation converting the DEPMPO/OOH adduct into DEPMPO/OH. Although this is likely to occur in the membrane preparation as well, there is a profound difference between the spectral features of Figs 1B and 3A. Control experiments with the HX/XO free radical-generating system in HEPES buffer, together with DETAPAC and NADH but without any membranes present, did not show any significant difference compared with the system presented in Fig. 1B (data not presented). In the membrane preparation, the signal of both adducts is visible from the very moment the spin-trap is added to the membrane system (Fig. 3A), but not in the XA/XAO system (Fig. 1B) which demonstrates that, at least in part, the signal of the •OH radical comes from the independent •OH radical production mechanism. The intensity of EPR peaks of DEPMPO/OOH and DEPMPO/OH spin adducts in Fig. 3B (peak intensity ratio around 0.8) should not be considered as a quantitative measure for the production of respective radical species by the plasma membrane. Based on results shown in Figs 1 and 2, the intensity of the DEPMPO/OOH peak reflects the production of the radical, whose signal is continuously diminished through conversion of the DEPMPO/OOH adduct into DEPMPO/OH. Conversely, the intensity of the DEPMPO/OH adduct is the superposition of the membrane production •OH and the amount of DEPMPO/OH adduct originating from the conversion of DEPMPO/OOH. Based on experiments in the absence of NADH (Fig. 5B) the estimated contribution of these two processes after 50 min to the overall amount of DEPMPO/OH is around 70:30%.
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Figure 4 shows the influence of oxygen on the production of radicals. Figure 4B shows the EPR spectrum of the membrane preparation in the glass capillary after 25 min (this spectrum has the optimal S/N, i.e. the highest production of radicals, after which S/N decreases). Apparently, the limited oxygen supply influences the amount
radicals more than the •OH radical (the ratio is around 0.6). An apparently poor signal in the glass capillary is partly the result of the fact that, in the case of the glass capillary, the EPR signal originated from a single 2.5 cm section of capillary, while in case of the gas-permeable tube, four 2.5 cm long segments were placed into the sensitive volume of the EPR cavity, i.e. the EPR signal also reflects the amount of sample in the cavity. Figure 4C is a system where everything is prepurged with N2, and recorded in the gas-permeable tube, while maintaining the N2 atmosphere in the cavity. Here the ratio of OOH/OH adducts is around 0.3 which is even less than in Fig. 4B, which is to be expected since the membranes had insufficient oxygen from the onset of the reaction. This indicates that the presence of oxygen predominantly affects the production of the radical, while production of •OH radicals is less affected by the anaerobic conditions. Figure 4D shows that upon the reintroduction of oxygen, membranes start to produce radicals. The rate of production is faster than in the air (DEPMPO/OOH peak increases five times in just 30 min) supporting the previous conclusion about the requirement of oxygen. The peak of the DEPMPO/OH adduct also slowly increases upon the introduction of oxygen, probably due to the transformation of the OOH adducts. The ratio of OOH/OH adducts is 0.8 after 30 min in O2, i.e. the same as in the standard membrane preparation (Fig. 3B).
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To differentiate between the possible mechanisms which could contribute to the overall production of the two radical species, radical quenching or differential inhibition of superoxide synthase and peroxidases was performed (Fig. 5). Figure 5B shows that, in the absence of NADH, there is virtually no production of DEPMPO/OOH adduct and the EPR spectrum resembles the one obtained in the Fenton reaction (Fig. 1A). The presence of SOD in the preparation eliminates the signal of the DEPMPO/OOH adduct from the spectra (Fig. 5C) and partially decreases the peak intensity of the DEPMPO/OH adduct. Addition of DPI (Fig. 5D) significantly reduces the amount of DEPMPO/OOH, while the spectral intensity of the DEPMPO/OH adduct increases (Fig. 6). The ratio of OOH/OH in the membrane preparation in the presence of 100 µM DPI is 0.25 (the ratio depends on the DPI concentration, Fig. 7B). The addition of KCN into the standard membrane preparation (Fig. 5E) reduces the overall intensity of all EPR peaks roughly by a factor of two, the ratio of the two peaks being 0.6. In the case of heat-denaturated membranes (Fig. 5F) a small quantity of DEPMPO adduct, similar to OH, can be observed (albeit greater than in buffer and NADH). A similar result has been obtained using heat-killed lymphocytes (Chamulitrat, 1999
).
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The kinetics of the production of
and •OH is similar in the case of membranes with NADH present (Fig. 6A). However, the aforementioned transformation of the OOH adduct into the OH adduct which increases the apparent rate of production of •OH, should be borne in mind. A difference in kinetics could be observed in the system with no NADH present (Fig. 6B), in which case DEPMPO/OOH was practically undetectable, DEPMPO/OH was observed, albeit decreased in intensity (which is obvious from the spectra, Fig. 5B). The most interesting observation in the kinetics was in the case when NADH was present together with DPI. Production of was diminished and slow, while the production of •OH was significant and increased with time.
The dependence of spectral intensities of OOH and OH adducts on NADH concentration (Fig. 7) showed no sign of saturation of production up to at least 3.3 mM, while •OH production was almost saturated at NADH concentrations above 0.2 mM.
The results obtained with different DPI concentrations (Fig. 8) demonstrated the opposite effect of DPI on the production of •OH and . Increasing the concentration of DPI up to 0.2 mM induced increasing inhibition of production, being approximately 50% at 0.2 mM DPI. From the spectra shown in the upper part of Fig. 8 it is obvious that a change in the spectral form of the DEPMPO/OOH adduct is visible at lower DPI concentrations (20 and 50 µM), a rapid diminution of the DEPMPO/OOH adduct peak being observed at higher DPI concentrations. Contrary to superoxide, the production of •OH increased with increasing DPI concentration.
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Discussion |
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The results presented here, for the first time, provide clear evidence for the independent production of two radicals (
and •OH) by plasma membranes using a spin-trap capable of their differentiation. The presence of SOD in the membrane preparation completely eliminates the presence of the DEPMPO/OOH adduct and only diminishes the intensity of the DEPMPO/OH adduct. Although the DEPMPO spin-trap was developed a decade ago and proved useful in animal systems, it has not been used in the investigations of plants, nor has simultaneous detection of these two radicals been attempted. For example, in the study of wheat root plasma membranes (Qui et al., 1995), a Tiron semiquinone spin-trap (reportedly sensitive only to ) has been used. On the other hand, a 4-POBN spin-trap suitable only to trap •OH radicals has been used in studies of cultured rice cells (Kuchitsu et al., 1995) and maize coleoptiles (Liszkay et al., 2003). Although these results show that transformation of DEPMPO/OOH into DEPMPO/OH does occur even in the HX/XO system, the advantage of DEPMPO over the more frequently used DMPO is that transformation is slow, thus enabling the reliable differentiation of adducts. However, reliable differentiation of different mechanisms of radical production can be achieved only through the application of inhibitors or activators.
The use of gas-permeable Teflon tubes in EPR spectroscopy is not new, these were first introduced some 25 years ago (Plachy and Windrem, 1977), and subsequently extensively used in EPR oxymetry studies (Swartz and Glockner, 1989). However, their potential usefulness somehow seems to have been overlooked by those working with spin-traps. The use of gas-permeable tubes appears to be essential when dealing with oxygen-consuming systems, because kinetic studies of radical production and the decay of trapped product performed in glass capillaries or flat cells are hampered by the fact that they are conducted in an environment of progressively decreasing amounts of oxygen. This, at least in part, could be an explanation for the widely varying values (more than an order of magnitude) for the rate constants reported for the formation of spin-adducts as well as for their decay under seemingly similar conditions (see discussion in Keszler et al., 2003). Vuleti et al. (2003b) have recently reported the production of radical species by maize root plasma membranes using the EMPO spin-trap in glass tubes and obtained indications for the production of both the superoxide and hydroxyl radicals. Due to the above-mentioned limitations of glass capillaries which were used in those measurements, no clear cut conclusions could be reached.
The dependence of production on NADH concentration shows no sign of saturation at concentrations of 3.3 mM in this study. This is contrary to the results obtained previously with the same plasma membrane preparations (Km=0.019 mM) measured by monitoring nitroblue tetrazolium reduction (Vuleti et al., 2003a) or the results obtained by lucigenin luminescence (Km=0.159 mM NADH) with rose cell plasma membranes (Murphy and Auh, 1996). Similar unsaturated kinetics, in the concentration range up to 0.3 mM NADH, was obtained in a much more complex system of maize coleoptiles, when looking at the oxidation of externally added NADH (Liszkay et al., 2003). On the other hand, in the EPR study of plant plasma membranes (Qui et al., 1995), the spin-trap signal amplitude (assumed to represent production) showed saturation kinetics at NADH concentrations higher than 4 mM. If the possible effect of different species, object, and level of complexity are excluded, the fact remains that this study's own membrane preparations, from the same plant species and variety isolated in the same way show marked differences (one order of magnitude in NADH concentration) depending upon the method of measurement.
Unlike the production of , the production of the •OH radical showed saturation at NADH concentrations around 0.2 mM. Saturation kinetics of •OH radical production by maize coleoptiles demonstrated half the maximal rate at approximately 1 mM NADH, obtained by EPR measurements (Liszkay et al., 2003). Corresponding measurements with purified horseradish peroxidase yielded a slightly lower value (0.7 mM NADH). What is obvious from the kinetics of NADH concentration dependence of the two adducts in these experiments is that different mechanisms of NADH-dependent production are involved in their formation. Thus, it would seem that, at lower concentrations of NADH, both radicals are produced in a similar manner, while at higher concentrations of NADH an additional superoxide-producing mechanism is activated. Perhaps the answer to the above-mentioned discrepancy noted in spin-trapping measurements, as opposed to tetrazolium or lucigenin superoxide measurements should be sought in the ability of the spin-trap to interact with an intermediate in the additional superoxide-producing mechanism activated at higher NADH concentrations. This point requires clarification in the future.
The observed inhibition of the DEPMPO/OOH adduct at higher DPI concentrations (approximately 50% >100 µM) is similar to what has been measured previously with nitroblue tetrazolium measurements on similar maize root plasma membrane preparations (0.1 mM DPI produced 40% inhibition) (Vuleti et al., 2003a), and what has been measured in partially purified superoxide synthase from bean plasma membranes (0.1 mM produced 73% inhibition) (Van Gestellen et al., 1997). The oxidative burst in plant cells was, however, shown to be inhibited by low concentrations of DPI (rose cells, 2 µM DPI 50% inhibition) (Bolwell et al., 1998), and this has been taken as an indication of the involvement of a phagocyte-type NADPH oxidase, in which such low concentrations were effective in inhibition. Inhibition of the neutrophil NADPH oxidase by DPI occurs through a single-electron reduction of DPI to a free radical, a reduced redox centre in the oxidase serving as an electron donor to DPI (O'Donnell et al., 1993). DPI is generally considered an inhibitor of flavin-containing redox enzymes, although it also strongly inhibits a partially purified NADH oxidase from rose cells that shows a fluorescence spectrum characteristic of pterins with no flavin spectrum present (Murphy et al., 2000). Fluorescence spectra of these membranes show that they contain both flavin and pterin moities (M Mojovi et al., unpublished results). On the other hand, it was demonstrated that DPI could also be an inhibitor of the peroxidase-catalysed H2O2 production by French bean cells (40 µM DPI for 50% inhibition) (Bolwell et al., 1998) and purified horseradish peroxidase activity (10 µM DPI for 50% inhibition) (Frahry and Schopfer, 1998), suggesting that DPI can form a free radical by reaction with peroxidase–haem. These results demonstrate that the specificity of DPI is limited, and should not be taken at its face value (Bérczi and Møller, 2000). However, the fact remains that KCN, inhibiting the peroxidases, does not inhibit completely the production of either the superoxide or hydroxyl radicals, and that DPI exhibits a selective partial inhibition of the superoxide production. The OOH adducts also show a clear dependence on the presence of oxygen, while the OH adduct is oxygen insensitive. Combined together, these findings argue in favour of the presence of superoxide synthase in this study's preparations. However, this is not the only source of superoxide radicals.
To conclude, it is obvious from the results obtained using an improved experimental approach that the plasma membrane preparations in this study are capable of producing both the superoxide and hydroxyl radical species. While there have been previous reports on the capacity of various plasma membranes to produce superoxide, no evidence is known that demonstrates their capacity to release hydroxyl radicals in plants. Based on the comparison of their NADH concentration, inhibitor and oxygen dependence, it is clearly shown that different enzymatic mechanisms are underlying such production. It seems possible that the presence of NADH in the membrane preparation enhances the production of free radical species through energization of the NADH-coupled redox plasma membrane systems. Inhibition of the endogenous systems through substrate removal (in the present case oxygen) or inhibitor blocking gives rise to enhanced production of potent hydroxyl radicals.
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Acknowledgements |
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This work was supported by grants OI-1934 and 1928 from the Ministry of Science and Environmental Protection of Republic of Serbia. The authors would like to acknowledge the generous gift of Teflon tubing from Zeus industries, Raritan, NJ.
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Footnotes |
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Abbreviations: DEPMPO, 5-diethoxy-phosphoryl-5-methyl-1-pyrroline N-oxide; DETAPAC, diethylenetriamine pentaacetic acid; DPI, diphenylene iodonium; DTT, dithiothreitol; EDTA, ethylenediaminetetra-acetic acid; EPR, electron paramagnetic resonance; HX/XO, hypoxanthine/xanthine oxidase; SOD, superoxide dismutase; DMPO, 5,5-dimethyl-1-pyrroline N-oxide.
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References |
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