TY - JOUR
AU - Noctor, Graham
AB - Abstract Leaf metabolism produces H2O2 at high rates, but current concepts suggest that the potent signalling effects of this oxidant require that concentrations be controlled by a battery of antioxidative enzymes. The extent to which H2O2 is allowed to accumulate remains unclear. There is little consensus on leaf H2O2 values in the literature and measured concentrations in unstressed conditions range from 50–5000 nmol g−1 fresh weight, a difference that probably reflects technical inaccuracies as much as biological variability. This article uses new experimental and literature data to examine some of the difficulties in accurately measuring H2O2 in leaf extracts. Potential problems relate to sensitivity, interference from other redox-active compounds, and H2O2 stability during sample preparation. Particular attention is drawn to the influence of tissue mass/extraction volume in the quantitative estimation of H2O2 contents, and the possibility that this factor could contribute to the variability of literature data. Ascorbate, chemiluminescence, oxidative stress, redox signalling, xylenol orange H2O2 signalling and chemistry Intense interest has focused on H2O2 as a potent signalling molecule whose production can occur though basal energy metabolism as well as through specific enzyme systems. Production of H2O2 through both routes can be affected by a variety of environmental factors such as light, cold, and biotic stress (Lamb and Dixon, 1997; Apel and Hirt, 2004). Changes in H2O2 concentrations are also important in developmental and hormone signalling (Kwak et al., 2003; Gapper and Dolan, 2006). Unlike other reactive oxygen species, which have been characterized in the twentieth century, H2O2 was first discovered in the early nineteenth century. The molecule is not a radical and is relatively unreactive with only weak oxidizing power. For example, the rate constant for uncatalysed reaction of H2O2 with ascorbate is about 100 000 times slower than that of superoxide (Polle, 2001). However, in the presence of transition metals such as iron or metal complexes, H2O2 can be rapidly degraded through oxidation or reduction reactions. Such reactions are exploited in industrial processes where H2O2 is reductively cleaved to the highly oxidizing hydroxyl radical, the existence of which was first inferred by Fenton in 1876. The potential for conversion of H2O2 to more reactive species is controlled inside cells, first, by systems that regulate the availability of metals such as iron (Hell and Stephan, 2003) and, second, by the existence of numerous H2O2-metabolizing enzymes, many of which themselves depend on iron-dependent catalysis through haem groups (Mittler et al., 2004). That H2O2 can function as a potent signalling molecule in both stress and developmental processes implies that concentrations are likely to be under homeostatic regulation. Proof of the importance of H2O2 homeostasis has come from the study of plants deficient in ascorbate peroxidase and catalases. These plants mimic stress-induced redox perturbation, gene expression and/or phenotypes such as cell death (Smith et al., 1984; Willekens et al., 1997; Dat et al., 2001; Noctor et al., 2002; Rizhsky et al., 2002; Davletova et al., 2005; Vanderauwera et al., 2005; Queval et al., 2007). However, despite the well-characterized redox perturbation observed in these studies, marked increases in leaf H2O2 contents have generally not been described. It is therefore unclear to what extent localized changes in H2O2 accumulation affect whole leaf contents, and whether measurement of these contents provides useful information on tissue redox state. H2O2 production and accumulation in leaves: inferences from modelling, enzyme inactivation, and the kinetic properties of H2O2-processing enzymes Model calculations Photosynthetic tissues have a high capacity for H2O2 production through electron transport chain activity, and in C3 plants, through the photorespiratory enzyme, glycolate oxidase. Although it is very difficult to establish precise rates of H2O2 production linked to photosynthesis, approximate calculations can easily be performed by simple modelling (Noctor et al., 2002). Thus, it is possible to estimate likely rates of H2O2 generation by electron leakage to O2 followed by dismutation or reduction of superoxide. A typical net rate of photosynthesis under moderate light is 50 μmol O2 evolved mg−1 chlorophyll h−1, equivalent to more than 200 μmol electrons mg−1chlorophyll h−1. Assuming that 1% of the total electrons from water reduce O2 (a fairly conservative estimate), that all superoxide is converted to H2O2, and that the chloroplast volume is 25–70 μl mg−1chlorophyll (Heldt, 1980; Winter et al., 1993), the photosynthetic electron transport chain is theoretically capable of generating a stromal H2O2 concentration of 0.2–0.5 M within 12 h photosynthesis. In C3 plants, equally or more rapid H2O2 production can simultaneously occur in the peroxisomes through photorespiration, and the mitochondrial electron transport chain is a further source of H2O2 production via superoxide (Foyer and Noctor, 2003). Enzyme inactivation It is evident that the overwhelming majority of the H2O2 that is produced must be metabolized since it has long been recognized that micromolar concentrations of H2O2 can inactivate thiol-regulated enzymes such as fructose-1,6-bisphosphatase and modulate the activities of metabolic pathways in isolated chloroplasts (Kaiser, 1979; Charles and Halliwell, 1980). It is unclear whether the H2O2 sensitivity of isolated enzymes or organelles is an accurate reflection of the sensitivity of photosynthesis in vivo, where reductant availability could be greater and H2O2 metabolism more powerful. H2O2-metabolizing enzymes Among the primary enzymes in H2O2 metabolism are ascorbate peroxidases (APX), whose existence was demonstrated in isolated thylakoids and chloroplasts by Groden and Beck (1979) and Kelly and Latzko (1979). Subsequently, the notion that catalase is an important enzyme in H2O2 metabolism inside chloroplasts became discounted and it is now accepted that APX functions alongside peroxiredoxins as a major H2O2-metabolizing enzymes in this organelle (Asada, 1999; Dietz, 2003). Unlike catalase, APX has high affinity for H2O2 (Km around 20–50 μM: Chen and Asada, 1989; Yoshimura et al., 1998). The Km H2O2 of a chloroplastic peroxiredoxin has been estimated at about 2 μM (König et al., 2002). Although the affinity of an enzyme for its substrate is not proof of the substrate's concentration range in vivo, Km values that are close to or higher than typical substrate concentrations allow catalytic activity to accelerate rapidly in response to increasing availability of substrate. Such properties act both to drive metabolic flux and to damp increases in substrate pools in response to increases in substrate production. They may be important in controlling the accumulation of molecules such as H2O2, whose rate of production likely shows rapid changes in response to factors such as fluctuating irradiance. If substrate concentrations are higher than enzyme Km values, the enzyme activity has more limited sensitivity to increases in substrate. For chloroplast APX, this may well be the case for ascorbate, since Km values are within the range 0.2–0.4 mM (Chen and Asada, 1989; Yoshimura et al., 1998), probably significantly below ascorbate concentrations in this compartment. Evidence that ascorbate concentration has little influence over chloroplastic APX activity has come from analysis of ascorbate-deficient mutants (Veljovic-Jovanovic et al., 2001; Müller-Moulé et al., 2002). If the catalytic rate of APX is similarly insensitive to changes in H2O2 concentrations then other mechanisms such as increased enzyme abundance or post-translational modification would be necessary to allow flexibility in enzyme activity in response to fluctuations in H2O2 production. Indeed, increases in extra-chloroplastic APX capacities in ascorbate-deficient lines (Veljovic-Jovanovic et al., 2001) could reflect a compensatory response to limitation by cytosolic ascorbate concentration. Mitochondria contain an APX isoform encoded by the same gene as the stromal APX (Chew et al., 2003). In the cytosol, APX isoforms so far characterized have Km H2O2 values around 20 μM (Chen and Asada, 1989; Mittler and Zilinskas, 1991; Yoshimura et al., 1998). Given that the primary physiological ‘function’ of these enzymes is considered to be maintenance of H2O2 homeostasis, their properties could be taken as circumstantial evidence for H2O2 concentrations in the micromolar range. Organelle concentrations, whole tissue contents, and compartmentation Chloroplasts and mitochondria are often included among the major sites of intracellular H2O2 production. The densities of many leaves are close to 1 g ml−1 (Vile et al., 2005), and so if H2O2 sensitivity of metabolic enzymes and APX affinities are indicators of typical H2O2 concentrations, and these are uniform within the leaf (see below), this would convert to about 20–50 nmol g−1 fresh weight (FW). Such values are at the low end of those reported in the literature (Table 1). It should be emphasized that even concentrations of 20–50 μM in chloroplasts, mitochondria, and the cytosol could be considered somewhat high. In rat liver cells, the global intracellular H2O2 concentration has been estimated at 1–100 nM with a calculated steady-state mitochondrial concentration of 40 nM, and a threshold for H2O2-triggered apoptosis of 700 nM (Stone and Yang, 2006). Similar concentrations have been reported for E. coli cells, where the intracellular concentration was estimated at 200 nM with less than 2-fold variation over the culture cycle (González-Flecha and Demple, 1997). Either plant cells are more tolerant to H2O2 than other types of cell or most measured leaf contents provide information of little relevance to H2O2 concentrations in compartments such as the chloroplast and mitochondria. Table 1. Diversity of literature values for leaf H2O2 contents Leaf H2O2 content Plant Technique Reference ≤100 nmol g−1 FW Barley Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002 Arabidopsis Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002; Bouché et al., 2003 100–500 nmol g−1 FW Tobacco Peroxidase catalysed-MBTH-DMAB absorbance Dutilleul et al., 2003 Tobacco Peroxidase-catalysed homovanillic acid fluorescence Creissen et al., 1999 Mustard Luminol chemiluminescence Dat et al., 1998 500–2000 nmol g−1 FW Mangrove Ferrous oxidation/xylenol orange Cheeseman, 2006 Tobacco Peroxidase-catalysed resorufin fluorescence Keetman et al., 2002 Maize Titanium-peroxide complex assay Zhang et al., 2006 Arabidopsis Luminol chemiluminescence Rao and Davis, 1999 ≥2000 nmol g−1 FW Soybean Ferrous oxidation/xylenol orange Cheeseman, 2006 Arabidopsis Luminol chemiluminescence Karpinski et al., 1997, 1999 Maize Titanium-peroxide complex assay Anderson et al., 1995 Tobacco Titanium-peroxide complex assay Mur et al., 2005 Leaf H2O2 content Plant Technique Reference ≤100 nmol g−1 FW Barley Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002 Arabidopsis Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002; Bouché et al., 2003 100–500 nmol g−1 FW Tobacco Peroxidase catalysed-MBTH-DMAB absorbance Dutilleul et al., 2003 Tobacco Peroxidase-catalysed homovanillic acid fluorescence Creissen et al., 1999 Mustard Luminol chemiluminescence Dat et al., 1998 500–2000 nmol g−1 FW Mangrove Ferrous oxidation/xylenol orange Cheeseman, 2006 Tobacco Peroxidase-catalysed resorufin fluorescence Keetman et al., 2002 Maize Titanium-peroxide complex assay Zhang et al., 2006 Arabidopsis Luminol chemiluminescence Rao and Davis, 1999 ≥2000 nmol g−1 FW Soybean Ferrous oxidation/xylenol orange Cheeseman, 2006 Arabidopsis Luminol chemiluminescence Karpinski et al., 1997, 1999 Maize Titanium-peroxide complex assay Anderson et al., 1995 Tobacco Titanium-peroxide complex assay Mur et al., 2005 Values listed are a selection of data for wild-type plants grown under ‘unstressed’ conditions. Neither the techniques nor the references cited are intended to represent an exhaustive inventory of the relevant literature. Open in new tab Table 1. Diversity of literature values for leaf H2O2 contents Leaf H2O2 content Plant Technique Reference ≤100 nmol g−1 FW Barley Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002 Arabidopsis Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002; Bouché et al., 2003 100–500 nmol g−1 FW Tobacco Peroxidase catalysed-MBTH-DMAB absorbance Dutilleul et al., 2003 Tobacco Peroxidase-catalysed homovanillic acid fluorescence Creissen et al., 1999 Mustard Luminol chemiluminescence Dat et al., 1998 500–2000 nmol g−1 FW Mangrove Ferrous oxidation/xylenol orange Cheeseman, 2006 Tobacco Peroxidase-catalysed resorufin fluorescence Keetman et al., 2002 Maize Titanium-peroxide complex assay Zhang et al., 2006 Arabidopsis Luminol chemiluminescence Rao and Davis, 1999 ≥2000 nmol g−1 FW Soybean Ferrous oxidation/xylenol orange Cheeseman, 2006 Arabidopsis Luminol chemiluminescence Karpinski et al., 1997, 1999 Maize Titanium-peroxide complex assay Anderson et al., 1995 Tobacco Titanium-peroxide complex assay Mur et al., 2005 Leaf H2O2 content Plant Technique Reference ≤100 nmol g−1 FW Barley Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002 Arabidopsis Peroxidase catalysed-MBTH-DMAB absorbance Veljovic-Jovanovic et al., 2002; Bouché et al., 2003 100–500 nmol g−1 FW Tobacco Peroxidase catalysed-MBTH-DMAB absorbance Dutilleul et al., 2003 Tobacco Peroxidase-catalysed homovanillic acid fluorescence Creissen et al., 1999 Mustard Luminol chemiluminescence Dat et al., 1998 500–2000 nmol g−1 FW Mangrove Ferrous oxidation/xylenol orange Cheeseman, 2006 Tobacco Peroxidase-catalysed resorufin fluorescence Keetman et al., 2002 Maize Titanium-peroxide complex assay Zhang et al., 2006 Arabidopsis Luminol chemiluminescence Rao and Davis, 1999 ≥2000 nmol g−1 FW Soybean Ferrous oxidation/xylenol orange Cheeseman, 2006 Arabidopsis Luminol chemiluminescence Karpinski et al., 1997, 1999 Maize Titanium-peroxide complex assay Anderson et al., 1995 Tobacco Titanium-peroxide complex assay Mur et al., 2005 Values listed are a selection of data for wild-type plants grown under ‘unstressed’ conditions. Neither the techniques nor the references cited are intended to represent an exhaustive inventory of the relevant literature. Open in new tab In addition to pathways that generate H2O2 in bacteria and animal cells, plants produce H2O2 through specific metabolic pathways such as photosynthesis and photorespiration. However, contents higher than 2 μmol g−1 FW have been reported for dark-grown maize seedlings (Prasad et al., 1994), in which these pathways should not be operating. Leaf contents in the range of 1 μmol g−1 FW or greater could reflect preferential accumulation of H2O2 concentrations in specific compartments. With a pKa value of 11.6 for deprotonation to HO2− ⁠, H2O2 is the overwhelmingly predominant form at physiological pH and its movement across membranes is facilitated by specific aquaporins (Henzler and Steudle, 2000; Bienert et al., 2007). On the basis of data obtained with knockout mutants for a cytosolic APX form, it has been suggested that some part of the H2O2 produced within the chloroplast could diffuse into the cytosol to be metabolized (Davletova et al., 2005). H2O2 and free radicals are important in cell wall metabolism (Halliwell, 1978; Bolwell et al., 2002), and the apoplast compartments could be enriched in H2O2. Other compartments in which H2O2 could accumulate include the intrathylakoid space, mitochondrial intermembrane space, endomembrane systems, and vacuole. H2O2 has been reported to be preferentially concentrated in endosomes that are targeted to the vacuole during the response to salt stress (Leshem et al., 2006). It has previously been noted that current concepts suggest that redox buffering in most of these compartments could be low compared with the chloroplast stroma and mitochondrial matrix (Foyer and Noctor, 2005). The extent to which intercompartmental gradients in H2O2 concentration are established will depend on restriction of movement by membranes compared with the sink effect of the antioxidant system. The relative importance of these two factors has been proposed to vary with H2O2 concentration (Henzler and Steudle, 2000). Stress and H2O2 accumulation It is commonly considered that numerous stress conditions promote enhanced production of H2O2, and that this leads to increases in concentrations. A model of reactive oxygen production in the chloroplast showed that H2O2 concentration increased linearly with increasing superoxide production by the photosynthetic electron transport chain (Polle, 2001). These changes could occur in vivo as irradiance increases or as stress conditions limit the reoxidation of the photosynthetic electron transport chain by metabolism, potentially favouring auto-oxidation by O2. The model of Polle (2001) also showed that a significant factor that could cause very marked increases in chloroplast H2O2 was the amount of APX enzyme. When APX decreased to around 15% of typical chloroplast concentrations, H2O2 began to increase from the theoretical starting-point of about 1 μM, and concentrations reached 10 mM when APX was present at less than 1% of the typical concentration (Polle, 2001). Whether such effects are physiologically relevant is unclear, and other enzymes such as peroxiredoxins could also make some contribution to H2O2 homeostasis in vivo (Dietz, 2003). Nevertheless, it has been shown that stress conditions such as combined drought and high light cause marked inactivation of APX, together with inhibition of extractable phosphoribulokinase, a thiol-regulated stromal enzyme (Shikanai et al., 1998). Similar, though less dramatic, effects on APX can be observed when H2O2 is added directly to tobacco BY-2 cells or generated in the extracellular medium (De Pinto et al., 2006). Assuming that chloroplasts contribute about 25% of the volume of an expanded photosynthetic leaf cell (Winter et al., 1993), a hypothetical increase in H2O2 from 1 μM to 10 mM in this compartment alone would contribute an increase of about 2.5 μmol g−1 FW. Much attention has focused on the plasmalemma and apoplast as compartments in which increases in H2O2 contents occur, particularly in response to biotic stress. Such increases could result from the activities of NADPH oxidases or peroxidases (Lamb and Dixon, 1997; Frahry and Schopfer, 1998; Bolwell et al., 2002; Sagi and Fluhr, 2006; Bindschedler et al., 2006). H2O2 was estimated to accumulate to 1.2 mM in packed soybean cell cultures in which the oxidative burst is induced, and similar concentrations of added H2O2 are required for optimal defence expression (Lamb and Dixon, 1997). Higher concentrations have been used to study transcriptomic effects of H2O2, though most of the added H2O2 is metabolized by cells within a few minutes (Desikan et al., 2001). This indicates that longer-term maintenance of H2O2 concentrations in the millimolar range would require continuous generation and perhaps also decreased activity of the antioxidative system. A concentration as high as 10 mM H2O2, distributed uniformly in the apoplast, would contribute around 1 μmol g−1 FW, assuming that the apoplastic free space is about 10% of total volume (Winter et al., 1993; Fleischer and Ehwald, 1995). H2O2 detection in situ and quantification in extracts In situ staining of H2O2 can be achieved by using compounds such as 2,2′-diaminobenzidine (DAB). This provides an indicator of H2O2 production rather than concentration and is semi-quantitative, notably because DAB oxidation relies on in vivo peroxidases whose activity could be different between compartments or conditions (Thordal-Christensen et al., 1997). Another semi-quantitative staining method uses cerium chloride to generate cerium perhydroxides in vivo (Bestwick et al., 1997). Various techniques have been used to quantify H2O2 contents in plant tissue extracts. Most are based on oxidation of substrates to form either complexes or oxidized products with altered spectral characteristics. In general, light emission as fluorescence or luminescence offers greater sensitivity than absorbance measurements. Potential problems of H2O2 specificity can be overcome by using enzymes. Specificity can be conferred by using peroxidase in the assay itself, for example, linked to changes in the fluorescence yields of scopoletin, homovanillin, or resorufin (Guilbault et al., 1967; Mohanty et al., 1997). Other assays rely on chemical specificity conferred through metal catalysts of the H2O2 (or peroxide)-dependent reaction. The ferrous xylenol orange (FOX) assay is linked to the oxidation of iron, which can be achieved by various peroxides (Wolff, 1994). In the chemiluminescence (CL) reaction, ferricyanide can be used as the catalyst that allows oxidation of luminol (Warm and Laties, 1982). When ferricyanide is the catalyst, there is a linear response of luminescence yield to H2O2 concentration over a certain range, whereas a more complex relationship is evident when other catalysts, including peroxidase, are employed (Navaz Díaz et al., 1996). When assays are themselves non-enzymatic, specificity for H2O2 can be verified by assaying parallel aliquots pre-treated or not with commercial catalase. Most of these techniques have been available for many years and, as Table 1 shows, there is considerable divergence in literature data for leaf H2O2 contents, even in wild-type plants growing under ‘non stress’ conditions. The range of contents is at least 100-fold and even higher values have been reported in differentiating cotton fibres using the titanium complex technique (up to 5 mmol g−1 dry weight (DW): Potikha et al., 1999). Divergent values have been reported even for the same technique (e.g. for the luminol assay). Likewise, there is no strong correlation between H2O2 contents and species. Though some of the variation could possibly be due to differences in developmental stage or culture conditions, it would perhaps be surprizing if an important signalling molecule that is produced at high rates during primary metabolism showed great elasticity in concentration. One example of a species that shows highly divergent values is Arabidopsis. Using peroxidase-coupled oxidation, values of approximately 50 nmol g−1 FW were obtained (Veljovic-Jovanovic et al., 2002; Bouché et al., 2003) whereas luminol chemiluminescence with catalase pre-treatment yielded values of around 5 μmol g−1 FW (Karpinski et al., 1997, 1999). This 100-fold difference is much greater than reported condition-dependent changes in measured contents. For example, differences in H2O2 between light and dark or between low and excess irradiance are less than 2-fold (Karpinski et al., 1999; Veljovic-Jovanovic et al., 2002; Bouché et al., 2003; Dutilleul et al., 2003). These small changes in H2O2 contents relative to what are probably much greater changes in the rate of H2O2 production are consistent with the idea that global leaf H2O2 homeostasis is promoted through concomitant changes in engagement of the antioxidative system. Chilling induced a transient 4-fold increases in the H2O2 contents of dark-grown maize seedlings, with a peak value of 8 μmol g−1 FW after about 4 h (Prasad et al., 1994), but this 4-fold change is still much smaller than the variation in literature values (Table 1). It therefore remains unclear what factors contribute to the large variability in absolute leaf H2O2 contents that are reported under non-stress conditions. One possibility is that they are partly caused by differences in experimental procedures, and some of the factors that could be responsible for such effects are discussed below with reference to two different H2O2 assays. The ferrous xylenol orange (FOX) assay: complex interactions involving ascorbate and O2 The FOX technique measures the absorbance of the xylenol orange complex that forms with oxidized metals such as ferric iron. In an assay mix containing xylenol orange and ferrous ion, the formation of a ferric ion–xylenol orange complex is dependent on hydroperoxides, including H2O2 (Wolff, 1994). A modified FOX assay was recently used to measure leaf H2O2 values for several species sampled under natural conditions (Cheeseman, 2006). A number of improvements to the assay were reported, including the use of differential absorbance changes (A550–A800), deoxygenation of assay solutions with nitrogen, and the use of sensitizer compounds such as ethanol (Cheeseman, 2006). Deoxygenation is important to prevent tissue-dependent artefactual changes: when the FOX assay is performed in oxygenated solutions, continuous increases in absorbance are observed in the presence of leaf extracts (Fig. 1A). These continuous changes are not observed in blanks or standard H2O2 solutions, for which a plateau value is reached within about 5 min (Fig. 1B). The slow extract-dependent changes can be prevented by pre-treatment with ascorbate oxidase (AO; Fig. 1A) and can be mimicked in standards and blanks by inclusion of ascorbate (Fig. 1A, B). This suggests that continuous changes occurring during the assay of extracts are due to ascorbate-dependent interactions with iron present in the assay, as previously described for FOX assays in animal tissues (Bleau et al., 1998). Interference from ascorbate has also been described for two other H2O2 assays (Veljovic-Jovanovic et al., 2002). Fig. 1. Open in new tabDownload slide Interaction between ascorbate and O2 in the ferrous xylenol orange (FOX) assay. (A) Slow increases in A550 in leaf extracts are abolished by pre-treatment with ascorbate oxidase (AO). (B) Ascorbate (asc) causes a slow gradual increase in A550 in both blank (0 H2O2) and standard (5 μM H2O2) cuvettes. (C) Standard curves for H2O2 are not affected by the presence of extracts assayed in air. (Circles) H2O2 alone. (Triangles) H2O2+leaf extract. (D) When assay solutions are deoxygenated, the response factor of the FOX assay to H2O2 is much lower, and is affected by the presence of extract. (Circles) H2O2 alone. (Triangles) H2O2+leaf extract. Representative data are shown; similar effects were observed in two other experiments. Standard solutions or neutralized acidic extracts of Arabidopsis leaves were assayed for H2O2 in 0.25 mM ferrous ammonium sulphate, 0.1 mM xylenol orange, 0.1 mM sorbitol, 1% ethanol (v/v), 25 mM H2SO4. Leaf extracts (100 mg FW) were ground in liquid nitrogen then in 1 ml 0.2 N HCl. After centrifugation, supernatant aliquots were adjusted to pH 5 with 0.2 M NaOH in the presence of phosphate buffer and assayed as such or after incubation of aliquots for 5 min with 2 U ml−1 AO. Absorbance changes were monitored at 550 and 800 nm either without or following deoxygenation by bubbling of argon through reagent solutions and extract followed by 10 min degassing of the assay mix. No changes in A800 were observed during the assay. Fig. 1. Open in new tabDownload slide Interaction between ascorbate and O2 in the ferrous xylenol orange (FOX) assay. (A) Slow increases in A550 in leaf extracts are abolished by pre-treatment with ascorbate oxidase (AO). (B) Ascorbate (asc) causes a slow gradual increase in A550 in both blank (0 H2O2) and standard (5 μM H2O2) cuvettes. (C) Standard curves for H2O2 are not affected by the presence of extracts assayed in air. (Circles) H2O2 alone. (Triangles) H2O2+leaf extract. (D) When assay solutions are deoxygenated, the response factor of the FOX assay to H2O2 is much lower, and is affected by the presence of extract. (Circles) H2O2 alone. (Triangles) H2O2+leaf extract. Representative data are shown; similar effects were observed in two other experiments. Standard solutions or neutralized acidic extracts of Arabidopsis leaves were assayed for H2O2 in 0.25 mM ferrous ammonium sulphate, 0.1 mM xylenol orange, 0.1 mM sorbitol, 1% ethanol (v/v), 25 mM H2SO4. Leaf extracts (100 mg FW) were ground in liquid nitrogen then in 1 ml 0.2 N HCl. After centrifugation, supernatant aliquots were adjusted to pH 5 with 0.2 M NaOH in the presence of phosphate buffer and assayed as such or after incubation of aliquots for 5 min with 2 U ml−1 AO. Absorbance changes were monitored at 550 and 800 nm either without or following deoxygenation by bubbling of argon through reagent solutions and extract followed by 10 min degassing of the assay mix. No changes in A800 were observed during the assay. Deoxygenation, employed in the study of Cheeseman (2006), prevents these slow, continuous increases in absorbance (data not shown), but also decreases the response to standard H2O2 solutions from around 0.08 to about 0.01 μM−1 (Fig. 1, circles in C and D), corresponding to a decrease in apparent extinction coefficient from 8 to 1×104 M−1 cm−1. Moreover, leaf extracts increase the assay response to H2O2 about 2–2.5-fold when assay solutions are deoxygenated (Fig. 1D). Such interference between extracts and standards is not observed when assays are conducted in the presence of oxygen (Fig. 1C). The experiments shown in Fig. 1 suggest that removal of ascorbate is an alternative approach to removing oxygen in optimizing the FOX assay for H2O2, with the advantage that it enables relatively high sensitivity without interference from extracts. The data of Fig. 1 were obtained using leaf material extracted into HCl and neutralized with NaOH. Small but significant effects of salt concentrations were observed on the assay. Other experiments revealed that a more reliable response was obtained when extracts were performed into HClO4 then neutralized with K2CO3 to allow removal of KClO4 by centrifugation. However, similar interactions between extract, ascorbate, and O2 to those shown in Fig. 1 were also observed using this second extraction protocol. As in HCl extracts, these could be prevented in HClO4 extracts either by deoxygenation or by pre-treatment with ascorbate oxidase (data not shown). After calibration with reference to standard curves, the FOX assay detects total peroxides that increase linearly with extract volume (Fig. 2). In the deoxygenated assay, two to three times more peroxide was found when calculated relative to external standards (Fig. 2). This discrepancy reflects the stronger internal standard response in the deoxygenated assay (Fig. 1D), an effect that is absent when assays are performed in the presence of oxygen (Fig. 1C). Despite the greater quantity of peroxides detected in deoxygenated extracts, the absolute absorbance change was smaller, reflecting the lower apparent extinction coefficient under these conditions, as discussed above. Fig. 2. Open in new tabDownload slide Response of the FOX assay to extract volume. Arabidopsis leaf samples (100 mg FW) were extracted into 1 ml 1 N HClO4 and after centrifugation aliquots were neutralized with 1 M K2CO3 in the presence of phosphate buffer. Extracts were clarified by centrifugation and treated with AO as in the legend to Fig. 1. Peroxides were assayed as A550–A800 under anaerobic (black circles) or aerobic (white circles) conditions and quantified by reference to external standard curves using H2O2 treated with ascorbate oxidase and deoxygenated or not. Data are means ±SE of two or three data points obtained with different extracts. Fig. 2. Open in new tabDownload slide Response of the FOX assay to extract volume. Arabidopsis leaf samples (100 mg FW) were extracted into 1 ml 1 N HClO4 and after centrifugation aliquots were neutralized with 1 M K2CO3 in the presence of phosphate buffer. Extracts were clarified by centrifugation and treated with AO as in the legend to Fig. 1. Peroxides were assayed as A550–A800 under anaerobic (black circles) or aerobic (white circles) conditions and quantified by reference to external standard curves using H2O2 treated with ascorbate oxidase and deoxygenated or not. Data are means ±SE of two or three data points obtained with different extracts. Tissue concentration is a crucial determinant of H2O2 contents An obvious response to overcoming limitations of sensitivity is to increase the amount of tissue extracted. When this was done for the FOX assay, absorbance changes did not increase proportionally, even though, as shown in Fig. 2, the assay response was more or less linearly related to extract volume. A detailed examination of the effects of tissue mass on calculated leaf contents was performed. In this analysis, all extracts were performed into HClO4, neutralized, pre-treated with AO, then H2O2 was estimated specifically by pre-treatment or not of parallel aliquots with catalase. The FOX assay was performed in both aerobic and deoxygenated conditions, and was compared with the luminol chemiluminescence (CL) assay. When peroxides and H2O2 contents were calculated relative to fresh weight, contents decreased markedly as increasing amounts of tissue were extracted (Fig. 3). This effect was observed for all three types of assay (Fig. 3A–C). When 5 mg leaf material was extracted per ml, leaf H2O2 contents were about 500 nmol g−1 FW, but values were less than 50 nmol g−1 FW when 200 mg was extracted (Fig. 3A, B). In the case of the CL assay, no peroxide or H2O2 was detected when extracts were more concentrated than 100 mg ml−1, suggesting quenching reactions that completely mask the peroxide response. This effect was even more severe when extracts were not pre-treated to remove ascorbate, with chemiluminescence being completely quenched at even lower tissue mass extracted (data not shown). Because such quenching effects can also result from the action of phenolics (Navaz Díaz et al., 1995), ion-exchange columns were used to remove anions prior to assay. However, this did not prevent the decrease in calculated peroxide contents as more tissue mass was extracted, though it did generally increase the values obtained, particularly in more dilute extracts (Fig. 3D–F). Fig. 3. Open in new tabDownload slide Tissue mass/extraction volume affects peroxide detection by both the FOX assay and the chemiluminescence (CL) in Arabidopsis leaves. (A) FOX assay in the presence of air (no deoxygenation). (B) FOX assay after deoxygenation of assay mixes with argon. (C) Chemiluminescence assay. (D–F) As (A–C) except that extracts were pre-treated on a DOWEX ion-exchange resin (strongly basic Cl- form, 50–100 mesh : Sigma, Saint Quentin-Fallavier, France). Black circles show total peroxides. White circles show H2O2 estimated after pre-treatment with commercially obtained catalase. Leaf extracts were performed and pre-treated with AO as described in the legend to Fig. 2, and FOX assays as in the legend to Fig. 1 using 100 μl neutralized extract. For catalase treatments, aliquots of neutralized extract were incubated with 25 U ml−1 enzyme for 5 min. Control experiments with standard solutions of H2O2 showed that significant amounts of peroxide could not be detected after this treatment. For chemiluminescence assays, 50 μl neutralized extract was added to 35 μM luminol in 0.2 M ammonia (pH 9.5). After mixing, the reaction was started by addition of 70 μM potassium ferricyanide (final assay volume 0.7 ml) and luminescence was integrated over 2 s. Leaf contents were calculated with reference to H2O2 standard solutions. Fig. 3. Open in new tabDownload slide Tissue mass/extraction volume affects peroxide detection by both the FOX assay and the chemiluminescence (CL) in Arabidopsis leaves. (A) FOX assay in the presence of air (no deoxygenation). (B) FOX assay after deoxygenation of assay mixes with argon. (C) Chemiluminescence assay. (D–F) As (A–C) except that extracts were pre-treated on a DOWEX ion-exchange resin (strongly basic Cl- form, 50–100 mesh : Sigma, Saint Quentin-Fallavier, France). Black circles show total peroxides. White circles show H2O2 estimated after pre-treatment with commercially obtained catalase. Leaf extracts were performed and pre-treated with AO as described in the legend to Fig. 2, and FOX assays as in the legend to Fig. 1 using 100 μl neutralized extract. For catalase treatments, aliquots of neutralized extract were incubated with 25 U ml−1 enzyme for 5 min. Control experiments with standard solutions of H2O2 showed that significant amounts of peroxide could not be detected after this treatment. For chemiluminescence assays, 50 μl neutralized extract was added to 35 μM luminol in 0.2 M ammonia (pH 9.5). After mixing, the reaction was started by addition of 70 μM potassium ferricyanide (final assay volume 0.7 ml) and luminescence was integrated over 2 s. Leaf contents were calculated with reference to H2O2 standard solutions. Decreases in peroxide contents with increasing fresh weight reflect the fact that the amount of peroxides detected in the cuvette or luminescence tube remained relatively constant over a wide range of extracted fresh weights. This also occurred for specific analysis of H2O2 after catalase treatment, suggesting that the response was not simply due to an artefactual background. A possible explanation of these effects is that the assay response declines with extract concentration. However, no effect of extract concentration on response of standards is observed in the FOX assay in the presence of O2 (Fig. 4A, white bars). In the FOX assay under argon, the response of the standards actually increases in concentrated extracts (Fig. 4A, black bars). For the CL assay, the response of the standards does indeed decline with increasing tissue mass, but even after correction using internal standards, tissue contents were still higher in dilute extracts (Fig. 4B). Thus, the three types of assay show different effects of extract on the assay response, when this is estimated with internal standards. Despite this, all assays yielded much lower leaf H2O2 contents for concentrated extracts than for dilute ones. Fig. 4. Open in new tabDownload slide Effect of tissue mass/volume ratio on internal standards and glutathione and ascorbate extraction efficiency. (A) Internal standard response as a percentage of external standard response in the FOX assay under different conditions. Arabidopsis leaves were extracted as in Fig. 2 at 5 or 100 mg tissue ml−1 acid and pre-treated for 5 min with 2 U ml−1 AO. Assays were performed without (white bars) or after deoxygenation of assay mixes with argon (black bars). In each condition, parallel extracts were assayed with or without addition of 2 nmol H2O2 and the difference was related to external standard responses. The dotted line indicates parity between internal and external standards. (B) Effect of correction for internal standards on leaf peroxides detected by the chemiluminescence (CL) technique. Leaf extracts were extracted as in (A), pre-treated for 5 min with 2 U ml−1 AO, and the CL assay was performed as described for Fig. 3. For each extract mass, a second aliquot was assayed after addition of 2 nmol H2O2, and tissue contents were calculated relative to the internal standard. Black circles, total peroxides. White circles, peroxides after pre-treatment with catalase as described for Fig. 3. (C, D) Leaf glutathione and ascorbate contents measured at two different tissue fresh weight/extraction volume ratios. Total leaf contents of both compounds were extracted as for peroxides and assayed spectrophotometrically as described in Queval and Noctor (2007). Fig. 4. Open in new tabDownload slide Effect of tissue mass/volume ratio on internal standards and glutathione and ascorbate extraction efficiency. (A) Internal standard response as a percentage of external standard response in the FOX assay under different conditions. Arabidopsis leaves were extracted as in Fig. 2 at 5 or 100 mg tissue ml−1 acid and pre-treated for 5 min with 2 U ml−1 AO. Assays were performed without (white bars) or after deoxygenation of assay mixes with argon (black bars). In each condition, parallel extracts were assayed with or without addition of 2 nmol H2O2 and the difference was related to external standard responses. The dotted line indicates parity between internal and external standards. (B) Effect of correction for internal standards on leaf peroxides detected by the chemiluminescence (CL) technique. Leaf extracts were extracted as in (A), pre-treated for 5 min with 2 U ml−1 AO, and the CL assay was performed as described for Fig. 3. For each extract mass, a second aliquot was assayed after addition of 2 nmol H2O2, and tissue contents were calculated relative to the internal standard. Black circles, total peroxides. White circles, peroxides after pre-treatment with catalase as described for Fig. 3. (C, D) Leaf glutathione and ascorbate contents measured at two different tissue fresh weight/extraction volume ratios. Total leaf contents of both compounds were extracted as for peroxides and assayed spectrophotometrically as described in Queval and Noctor (2007). Extraction efficiency and H2O2 stability during sample preparation One explanation of the decreasing contents with fresh weight is that extraction efficiency decreases as fresh weight increases. To test this possibility, ascorbate and glutathione were assayed in the same extracts as H2O2. Similar leaf values for ascorbate and glutathione were obtained whether the extract concentration was 5 or 100 mg ml−1 (Fig. 4C, D), suggesting that tissue extraction efficiency is not markedly affected by the tissue concentration within the range examined in this study. A second explanation of the effects shown in Fig. 3 is that endogenous peroxides become progressively lost with increasing tissue mass. When the FOX assay was used to estimate recoveries of known amounts of H2O2 through the sample preparation procedure relative to external standards, the values depended on the extraction method and the assay procedure. First, recoveries were tested after extraction into different media at 100 mg FW ml−1, a ratio that gave low values for leaf H2O2 contents. When samples were extracted into water, without acidification, the recovery of added H2O2 was negligible (Fig. 5B), suggesting that enzymes such as catalase and peroxidases remain sufficiently active to destroy virtually all the added H2O2 under these conditions. Recoveries were much improved by extraction into acid. Under these conditions, apparent recoveries were close to 100% when assayed under argon and around 50% when assayed in the presence of O2 (Fig. 5A, B). Since assay of high amounts of tissue under argon causes an approximately 2-fold amplification of the assay response for both HCl and HClO4 extracts (Figs 1D, 4A), the values in air more accurately represent the true recovery. This suggests that approximately half of the added H2O2 was lost during sample preparation, even when powder ground freshly in liquid nitrogen is immediately acidified. Fig. 5. Open in new tabDownload slide Recoveries of H2O2 through the extraction procedure in different conditions of extraction and FOX assay. Material from approximately ten Arabidopsis leaves was cut into small pieces, mixed, and weighed in order to produce multiple samples that were immediately frozen in liquid nitrogen. Four to six samples were then finely ground in liquid nitrogen. Half were extracted into medium without added H2O2 (sample group 1) and half into medium supplemented with H2O2 sufficient to give 2 nmol in the assay cuvette after sample preparation (sample group 2). All aliquots were pre-treated for 5 min with 2 U ml−1 AO before assay. Recoveries were estimated after subtraction of endogenous leaf values found in sample group 1 from measured values in sample group 2. Data are means ±SE of two to three independent extracts. The dotted lines indicate 100% recovery. Black bars, total peroxides recovered. White bars, H2O2 estimated after treatment of parallel neutralized extracts with catalase. (A) Samples extracted into HClO4 as in Fig. 2 at 100 mg FW ml−1. Assays were performed without (right) or after deoxygenation of assay mixes with argon (left). (B) Samples extracted into HCl and neutralized as described in Fig. 1 (left) or into H2O (right). Assays were performed under argon. (C) Effect of tissue mass/HClO4 volume ratio on recoveries of H2O2 after HClO4 extraction and assay under argon. (D) Effect of tissue mass/HClO4 volume ratio on recoveries of H2O2 after HClO4 extraction and assay in air. Fig. 5. Open in new tabDownload slide Recoveries of H2O2 through the extraction procedure in different conditions of extraction and FOX assay. Material from approximately ten Arabidopsis leaves was cut into small pieces, mixed, and weighed in order to produce multiple samples that were immediately frozen in liquid nitrogen. Four to six samples were then finely ground in liquid nitrogen. Half were extracted into medium without added H2O2 (sample group 1) and half into medium supplemented with H2O2 sufficient to give 2 nmol in the assay cuvette after sample preparation (sample group 2). All aliquots were pre-treated for 5 min with 2 U ml−1 AO before assay. Recoveries were estimated after subtraction of endogenous leaf values found in sample group 1 from measured values in sample group 2. Data are means ±SE of two to three independent extracts. The dotted lines indicate 100% recovery. Black bars, total peroxides recovered. White bars, H2O2 estimated after treatment of parallel neutralized extracts with catalase. (A) Samples extracted into HClO4 as in Fig. 2 at 100 mg FW ml−1. Assays were performed without (right) or after deoxygenation of assay mixes with argon (left). (B) Samples extracted into HCl and neutralized as described in Fig. 1 (left) or into H2O (right). Assays were performed under argon. (C) Effect of tissue mass/HClO4 volume ratio on recoveries of H2O2 after HClO4 extraction and assay under argon. (D) Effect of tissue mass/HClO4 volume ratio on recoveries of H2O2 after HClO4 extraction and assay in air. A direct comparison of recoveries at different tissue mass/extract volume showed that these were around 50% in all conditions except when concentrated extracts were assayed under argon (Fig. 5C, D), a condition in which, as discussed above, the H2O2 response is amplified compared with external standards. This agrees with the data of Fig. 5A, and confirms that the recovery of added H2O2 was approximately 50% and that it was independent of the tissue mass:extract volume. What causes the tissue concentration-dependent differences in extractable H2O2? It is not clear how the smaller values obtained with high tissue weights can be explained, although they do raise question marks about the accuracy of at least some H2O2 assays. The data of Fig. 5 show that differences in leaf contents as a function of extracted tissue mass (Fig. 3) are not due to preferential loss of H2O2 during preparation of more concentrated samples, at least when this H2O2 is present in the extraction medium added to the tissue. The effect does not seems to be due to events occurring in the assay, since the response increases proportionally to extract volume (Fig. 2). Such linearity with extract volume was observed for extracts prepared at divergent tissue mass/volume ratios (data not shown). Despite this, when large volumes of dilute extracts and small volumes of concentrated extracts were compared, there was no correspondence of calculated values for the same equivalent fresh weight in the assay (data not shown). This suggests that the problem is linked to events occurring during the extraction. One possibility is that the dependence on tissue weight reflects different rates of reactions that occur at the very beginning of the extraction procedure and that cannot be inhibited by either low temperature or acid. This could involve metal-catalysed reactions that generate H2O2 in dilute extracts. More likely, perhaps, is that endogenous H2O2 is preferentially destroyed in more concentrated extracts. If this is so, it would presumably reflect reactions that show a non-linear dependence on reactant concentrations. If the leaf contents estimated in concentrated extracts are artefactually low, data obtained for low tissue mass/extraction medium ratios will deliver the most reliable data on tissue H2O2. In this case, H2O2 contents in leaves would be far superior to those that are reported for other organisms. Moreover, if the recovery of 50% added H2O2 through the extraction procedure is accurate, this would suggest that contents could be underestimated by about 2-fold. Thus, H2O2/peroxide contents of around 3 μmol g−1 FW, obtained in dilute extracts (Fig. 3E), would be adjustable to 6 μmol g−1 FW, which is within the range of values obtained by CL assay at the lowest tissue weight/extraction volume (Fig. 3F). Previous data in Arabidopsis, obtained with HClO4 extraction (tissue mass 67 mg FW ml−1), and using internal standards and catalase for accurate quantification (Karpinski et al., 1997), reported values of about 4 μmol g−1 FW. Sensitivity of H2O2 assays One caveat to the conclusion that H2O2 contents estimated at low tissue mass:extraction volume are an accurate reflection of leaf contents is that these are derived from small changes in the assay. This applies particularly to the FOX method. Typical absorbance changes for 100 μl of extract (100 mg FW ml−1) were 0.02–0.06 (assayed in air) and 0.013–0.016 (deoxygenated assay). Leaf contents in the μmol g−1 FW range, obtained in this study by assaying 100 μl of an extract prepared at 5 mg FW ml−1 HClO4, are calculated from an absorbance change (A550–A800) of about 0.005 in the deoxygenated procedure previously used by Cheeseman (2006). While changes are greater when sensitivity is increased by the presence of oxygen, the absorbance changes remain small. As well as ethanol and oxygen, high concentrations of sorbitol can be used to enhance the assay response and, in these conditions, the apparent extinction coefficient can reach 22×104 M−1 cm−1 (Bellincampi et al., 2000). However, the enhancing effect of concentrated sorbitol is linked to free-radical chain reactions (Gay and Gebicki, 2000) that could complicate comparisons of tissue H2O2 in biological samples if the abundance of other potentially interfering compounds differs. Concluding remarks The present discussion has drawn attention to problems in the quantitative assay of H2O2 in leaf extracts, and the implications of measured leaf contents for in vivo concentrations. Potential problems include interference in the assay, which have previously been noted for other techniques (Veljovic-Jovanovic et al., 2002), and, particularly, events occurring during the extraction. It has been considered that most H2O2 assays, while they may be of questionable specificity, provide an indicator of ‘generalized oxidative stress’ (Shulaev and Oliver, 2006). However, even this concept could be debatable if the data reflect complex interactions between compounds such as metals and ascorbate that can occur during the extraction and/or assay, but that are regulated or prohibited in the intact tissue. Alhough some of these problems might be intractable, Table 2 lists a number of procedures which could help to minimize or at least identify them. Table 2. Some procedures that could favour accurate quantification of H2O2 in tissue extracts Extraction     Acid with subsequent removal of added salt during neutralization     Low temperature, rapid Post-extraction sample preparation     Pre-treatment to remove ascorbate (particularly for ascorbate-rich tissues)     Removal of phenolics     Negative control with parallel catalase-treated extract fractions (especially if the assay is not enzyme-based) Assay sensitivity and quantification     Fluorescence or luminescence     Use of internal standard to correct for possible influence of extract on assay response Control experiments     Assay linearity with extract volume     Recovery of H2O2 through extraction and sample preparation procedures     Verification that detected leaf contents are not markedly dependent on small changes in tissue mass/extraction volume ratio Extraction     Acid with subsequent removal of added salt during neutralization     Low temperature, rapid Post-extraction sample preparation     Pre-treatment to remove ascorbate (particularly for ascorbate-rich tissues)     Removal of phenolics     Negative control with parallel catalase-treated extract fractions (especially if the assay is not enzyme-based) Assay sensitivity and quantification     Fluorescence or luminescence     Use of internal standard to correct for possible influence of extract on assay response Control experiments     Assay linearity with extract volume     Recovery of H2O2 through extraction and sample preparation procedures     Verification that detected leaf contents are not markedly dependent on small changes in tissue mass/extraction volume ratio Open in new tab Table 2. Some procedures that could favour accurate quantification of H2O2 in tissue extracts Extraction     Acid with subsequent removal of added salt during neutralization     Low temperature, rapid Post-extraction sample preparation     Pre-treatment to remove ascorbate (particularly for ascorbate-rich tissues)     Removal of phenolics     Negative control with parallel catalase-treated extract fractions (especially if the assay is not enzyme-based) Assay sensitivity and quantification     Fluorescence or luminescence     Use of internal standard to correct for possible influence of extract on assay response Control experiments     Assay linearity with extract volume     Recovery of H2O2 through extraction and sample preparation procedures     Verification that detected leaf contents are not markedly dependent on small changes in tissue mass/extraction volume ratio Extraction     Acid with subsequent removal of added salt during neutralization     Low temperature, rapid Post-extraction sample preparation     Pre-treatment to remove ascorbate (particularly for ascorbate-rich tissues)     Removal of phenolics     Negative control with parallel catalase-treated extract fractions (especially if the assay is not enzyme-based) Assay sensitivity and quantification     Fluorescence or luminescence     Use of internal standard to correct for possible influence of extract on assay response Control experiments     Assay linearity with extract volume     Recovery of H2O2 through extraction and sample preparation procedures     Verification that detected leaf contents are not markedly dependent on small changes in tissue mass/extraction volume ratio Open in new tab While the sensitivity of some enzymes to inactivation or the affinity of peroxidases suggests that H2O2 concentrations in certain compartments are unlikely to exceed 100 μM, many literature data point to considerably higher concentrations at other sites. Some of the potential artefacts associated with H2O2 assays might explain the variability of literature data. The present analysis suggest that some variability could be related to differences in the fresh weight:extraction volume ratio, as well as possible complex interactions during H2O2 assay. If significant underestimation occurs when relatively high tissue mass is extracted, the most accurate values would be obtained at very low amounts of extracted tissue. A recent careful study by Cheeseman (2006) reported values for several species within the range 0.5–5 μmol g−1 FW. This study used relatively dilute extracts, and the lowest leaf values were obtained for mangroves, for which extracts were more concentrated than other species such as soybean (Cheeseman, 2006). The implications of leaf H2O2 contents as high as 1 μmol g−1 FW are worthy of consideration. They imply that plants are able to tolerate much higher intracellular H2O2 concentrations than other organisms or that marked concentration gradients exist between intracellular and extracellular compartments. One possibility is that accumulation preferentially occurs in the apoplast or vascular tissues and that intracellular concentrations are much lower, possibly contributing to a redox gradient across the plasmalemma. If this is the case, leaf contents above 1 μmol g−1 FW would imply apoplastic values as high as 10 mM. Whether H2O2 accumulation in the vacuole contributes appreciably to leaf contents is unclear. Further analysis of these issues might have to await development of H2O2 sensors that can be calibrated to deliver reliable quantitative data on in vivo concentrations in specific compartments. High background H2O2 concentrations suggest that specialized mechanisms must operate to allow signalling that occurs, for example, through NADPH oxidases or peroxidases. One possibility is that production and/or perception of H2O2 is localized through protein–protein associations or through vesicle formation (Leshem et al., 2006). Alternatively, signalling specificity could be related to superoxide (rather than H2O2) or through the production of hydroxyl radicals generated from H2O2 by changes in metal availability (Foreman et al., 2003). A further possibility is that some of the functions of NADPH oxidases are linked to changes in cytosolic NADP redox state rather than ROS production. Finally, if background levels really are as high as some data suggest, extractable tissue H2O2 contents, even if they can be measured accurately, will be of limited value as indicators for oxidative stress or signalling. 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TI - Why are literature data for H2O2 contents so variable? A discussion of potential difficulties in the quantitative assay of leaf extracts
JO - Journal of Experimental Botany
DO - 10.1093/jxb/erm193
DA - 2008-02-01
UR - https://www.deepdyve.com/lp/oxford-university-press/why-are-literature-data-for-h2o2-contents-so-variable-a-discussion-of-IaWPw2MFLS
SP - 135
EP - 146
VL - 59
IS - 2
DP - DeepDyve
ER -