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* This work was supported, in part, by National Science Foundation Grants MCB-9310371 and MCB-9514239. The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Salicylic acid (SA) plays a key role in the establishment of resistance to microbial pathogens in many plants. The discovery that SA inhibits catalase from tobacco led us to suggest that H2O2 acts as second messenger to activate plant defenses. Detailed analyses of SA's interaction with tobacco and mammalian catalases indicate that SA acts as an electron donor for the peroxidative cycle of catalase. When H2O2 fluxes were relatively low (1 μM/min or less), SA inhibited catalase, consistent with its suggested signaling function via H2O2. However, significant inhibition was only observed at 100 μM SA or more, a level reached in infected, but not in uninfected, leaves. This inhibition was probably due to siphoning catalase into the slow peroxidative reaction. Surprisingly, SA was also able to protect catalase from inactivation by damaging levels of H2O2 (lower millimolar range), which is generally assumed to reflect accumulation of inactive ferro-oxy intermediates. SA did so by supporting or substituting for the protective function of catalase-bound NADPH. These results add new features to SA's interaction with heme enzymes and its in vivo redox properties. Thus, SA, in addition to its proposed signaling function, may also have an important antioxidant role in containing oxidative processes associated with plant defense responses.
Vertebrate animals possess a novel and highly specific immune system that acts as a defense against disease. Plants react to pathogen attack by activating elaborate defense mechanisms, which are much more poorly characterized than the vertebrate immune system. These defense mechanisms are activated not only at the sites of infection, which are manifested, in part, as necrotic lesions (hypersensitive response; HR),
but also in neighboring and even distal uninfected parts of the plant, leading to systemic acquired resistance (SAR). Both HR and SAR are associated with induction of a large number of defense-related genes. The products of these genes may play important roles in the restriction of pathogen growth and spread either indirectly, by participating in strengthening host cellular structures, or directly, by providing antimicrobial activities (for review see Ryals et al. (
)). Additionally, defense responses in surrounding cells become activated, which include synthesis of phytoalexins, pathogenesis-related (PR) proteins, and cell wall polymers such as lignin (Dempsey and Klessig,
). The detailed sequence of molecular events required for the initiation and regulation of HR and SAR is unknown, but progress has been made in identifying several components of the signal transduction pathways leading to disease resistance, among them salicylic acid (SA; for review see Staskawicz et al. (
SA is present in many plants. While the healing benefits of plants containing high levels of SA have been known since antiquity, the first insights regarding SA's role in plants have emerged only during the past decade. A mounting body of evidence has accumulated that indicates that SA plays an important role in plant defense responses (for review see Ryals et al. (
) was the first to demonstrate that application of exogenous SA or acetylsalicylic acid (aspirin) to tobacco induces PR gene expression and partial resistance to pathogens such as tobacco mosaic virus. Endogenous levels of SA increase dramatically after tobacco mosaic virus inoculation of resistant, but not susceptible, tobacco cultivars and parallel the induction of PR genes (Malamy et al.,
). Finally, tobacco mosaic virus-infected transgenic tobacco plants, which express the nahG gene that encodes the SA-metabolizing enzyme salicylate hydroxylase from Pseudomonas putida, accumulate little or no SA, fail to establish SAR, and develop viral lesions that are larger than those produced on wild type plants (Gaffney et al.,
To investigate how SA might function in plant defense responses, our laboratory has focused on the identification of cellular factors with which SA directly interacts. We have suggested that one mechanism of SA action is to inhibit catalase, thereby elevating endogenous levels of H2O2, which result either from the oxidative burst associated with the HR or from metabolic processes such as photorespiration, photosynthesis, and oxidative phosphorylation (Chen et al.,
). According to this working hypothesis, the elevated H2O2 or other reactive oxygen species derived from it would activate plant defense-related genes such as the PR-1 genes. This mode of activation of plant defenses has been compared with the induction of genes associated with mammalian immune, inflammatory, and acute phase responses that are mediated through H2O2 activation of the transcription factor NF-κB (Schmidt et al.,
). In support of this model, we have found that (i) 2,6-dichloroisonicotinic acid (INA; a synthetic inducer of PR genes and enhanced resistance) and its biologically active, but not inactive, analogues also inhibit tobacco catalase in vivo (Conrath et al.,
However, currently there is considerable controversy about the involvement of catalase inhibition by SA and the subsequent increase of H2O2 in plant defense responses. Several recent reports have indicated that H2O2 is unlikely to be involved in PR gene induction or SAR. Inhibition of catalase in leaf extracts requires concentrations of SA far above those observed in uninfected tissues (Bi et al.,
). In addition, while H2O2 and H2O2-inducing chemicals activate PR-1 genes in wild type tobacco, there is little or no gene induction in NahG plants. Therefore, it has been concluded that H2O2 induction of SAR genes requires SA rather than the reverse (Neuenschwander et al.,
). Taken together, these results suggest that H2O2 does not function downstream of SA (i.e. by inhibition of catalase) in the regulation of PR genes.
However, catalase is still the subject of many mechanistic investigations. There is increasing evidence that catalase is a major factor in a variety of pathological states such as cancer, diabetes, aging, and oxidative stress (see DeLuca et al. (
)). Inactivation and reactivation of catalase in vivo and in vitro are far from being fully understood. Numerous recent publications suggest new approaches regarding in vitro assays and inhibition studies on catalase (Feuers et al.,
). In the present report, we provide new insights into SA's effects on catalase. SA acts as an electron donor for the peroxidative cycle of both plant and animal catalases. As such, it can protect as well as inhibit catalase activity, depending on the concentration of H2O2. It is hypothesized that, in healthy tissue of infected leaves where H2O2 levels are low, SA inhibits catalase, which could lead to activation of defense-related genes. In contrast, in infected cells and in tissue immediately adjacent to necrotizing cells, where high levels of H2O2 and other reactive oxygen species are produced, SA protects catalase from inactivation. This property of SA might serve to contain the oxidative damage associated with spread of the lesion and resembles closely some antioxidative properties of SA in activated HeLa cells or inflamed mammalian tissues, which are unrelated to inhibition of prostaglandin H synthase.
Tobacco plants (Nicotiana tabacum cv. Xanthi nc) were grown at 22°C in growth chambers programmed for a 14-h light and 10-h dark cycle. 6-8-week-old plants were used for experimentation.
Chemicals and Enzymes
SA, SA analogues, simple phenolic compounds, phlorizin (phloretin-2′-β-D-glucoside), and bumetanide (3-n-butylamino-4-phenoxy-5-sulfomoyl-benzoic acid) were purchased from Sigma or Aldrich. INA was kindly provided by Helmut Kessmann of CIBA-Geigy Ltd. (Basel, Switzerland). Catalase (H2O2:H2O2 oxidoreductase; EC 184.108.40.206) from Aspergillus niger, bovine liver, and human erythrocytes were from Sigma. The heme content of the various preparations varied only slightly (A405/A280 was between 0.78 and 0.90).
Catalase activity was measured in 50 mM potassium phosphate, pH 6.6, at 24°C with a commercial oxygen electrode probe (model 5739; Yellow Springs Instruments, Yellow Springs, OH). If not otherwise indicated, the reaction was started by the addition of H2O2 to 10 mM (assuming the absence of any remaining H2O2 after the pretreatment). Enzyme activity (i.e. oxygen production) was followed for 2-4 min. Catalase activity is expressed in units of mmol of H2O2 decomposed/min, based on the catalatic cycle in which two molecules of H2O2 are degraded per molecule of O2 generated.
Alternatively, catalatic activity was measured spectrophotometrically by monitoring the consumption of H2O2 at 240 nm. The molar extinction of H2O2 at 240 nm was taken as 39.4 M−1 cm−1. The amount of enzyme added for the controls (without inhibitor/activator) was such that the decrease in A240 at 24°C occurred at a rate of 0.1 per min.
Commercially available catalase enzymes were treated as described previously for horseradish peroxidase (Durner and Klessig,
). 1 mg of enzyme was dissolved in 1 ml of 200 mM potassium phosphate, pH 6.2, containing 150 mM NaCl. Following a 15-min incubation at room temperature, the enzyme solution was centrifuged (10,000 × g, 10 min). After buffer exchange (NAP-10, Pharmacia Biotech Inc., equilibrated with 25 mM potassium phosphate, pH 6.6), the protein concentration was adjusted to 0.5 mg/ml.
Since peroxidase substrates such as guaiacol or pyrogallol are poor electron donors for catalase, its peroxidase reaction was measured by the oxidation of ethanol to acetaldehyde, which was determined by the 3-methyl-2-benzothiazolone hydrazone test (Zamocky et al.,
Protein concentration was determined with the Bio-Rad microassay.
Purification of Catalase from Tobacco
Tissue homogenization and all precipitation and chromatography steps were performed at 4°C. 1.3 kg of tobacco leaves were homogenized in a blender in 2.6 liters of extraction buffer consisting of 100 mM potassium phosphate (pH 6.6) containing 10% glycerol, 10 mM dithiothreitol, 1 μM leupeptin, 10 μM antipain, and 1 mM phenylmethylsulfonyl fluoride. The homogenate was filtered through four layers of cheesecloth and centrifuged at 9,000 × g for 15 min. Ammonium sulfate was added to 22% saturation, and the resulting suspension was stirred for an additional 1 h. After centrifugation at 14,000 × g for 25 min, the ammonium sulfate concentration of the supernatant was brought to 65% saturation. The pellet resulting from centrifugation (14,000 × g, 25 min) was resuspended in extraction buffer containing 25% glycerol and 1 mM dithiothreitol. At this point, the extract could be stored at −80°C until further use without significant loss of catalase activity (4820 mg of protein with a specific catalase activity of 0.098 units/mg, resulting in a total of 481 units).
The thawed extract was fractionated with 0.8 volumes of 0°C ethanol/chloroform (3:1, v/v) containing 1 mM phenylmethylsulfonyl fluoride. The upper aqueous layer, which contained the catalase activity, was centrifuged at 47,000 × g for 25 min. The supernatant (121 mg of protein, 1.6 units/mg) was diluted 4-fold with 20 mM potassium phosphate, pH 6.6, and applied to a phenyl-Sepharose column (5 × 12 cm, Pharmacia), equilibrated with the same buffer. After washing with several bed volumes, catalase activity was eluted by a ethylene glycol step (50%, v/v) in 20 mM potassium phosphate, pH 6.6 (11 mg of protein with 16.4 units/mg). Catalase activity was precipitated by the addition of 3 volumes of 50 mM potassium phosphate, pH 6.6, containing 90% (w/v) ammonium sulfate, and centrifugation at 47,000 × g for 25 min. After resuspension and buffer exchange (PD-10, Pharmacia) against 10 mM potassium phosphate, pH 6.6, the sample was applied to a hydroxyapatite column (1 × 5 cm, Bio-Rad) equilibrated with the same buffer. After washing (40 mM potassium phosphate), catalase was eluted with 200 mM potassium phosphate, pH 6.6. After the addition of glycerol to a final concentration of 25%, the purified catalase (1.9 mg, 64.3 units/mg) could be stored at −80°C.
A Mono P HR 5/20 column (Pharmacia) was connected to a fast protein liquid chromatography system and equilibrated with 25 mM triethanolamine, adjusted to pH 8.2 with iminodiacetic acid (start buffer). Prior to application, the sample was exchanged in start buffer using a NAP-5 column (Pharmacia). The composition of the Mono P eluent was Polybuffer 96/Polybuffer 74 (Pharmacia) in a ratio of 3.5/6.5, resulting in a pH of 5.7. The flow rate was 0.5 ml/min, and the fraction size was 0.5 ml. In order to obtain pools of catalase isozymes consisting of either subunit, preparative chromatofocusing with Polybuffer exchanger 94 (1 × 35 cm) was carried out under the conditions described above for the Mono P column.
To analyze the redox states of catalase, spectra in the near UV region (Soret region, 360-450 nm) were scanned with a Beckman DU-7 spectrophotometer, using 1-ml semimicro black sidewall quartz cuvettes. The binding of NADPH by tobacco and mammalian catalase was assayed by fluorescence spectroscopy using a LS-3B/R 100A system (Perkin-Elmer) and 0.5-ml fluorescence cells. After excitation at 340 nm, emission spectra from 360-560 nm were recorded. Immediately before the measurements, commercially available enzymes were treated as described under “Enzyme Assays.”
SDS-PAGE was performed with 10% (2.7% cross-linker) gels. Gels were stained with Coomassie Blue R-250 or with silver nitrate using the Bio-Rad silver stain kit. For immunoblotting, proteins were transferred to a nitrocellulose membrane, and catalase was detected with a mixture of monoclonal antibodies (MAb3B6 and MAb1F5) made against tobacco catalase and with the ECL detection kit from DuPont (Chen et al.,
Horizontal isoelectric focusing was carried out on Ampholine gels, pH 5.5-8.5 (Pharmacia), at 50 mA for 1 h (270-1100 V), followed by 1650 V for 1.5 h. The samples were prepared by grinding tissue under liquid nitrogen (0.2 g/ml of extraction buffer as described for the large scale extraction of catalase). After centrifugation for 10 min, the homogenate was desalted (NAP-5) against 10 mM potassium phosphate, pH 6.6. After focusing, gels were negatively stained for catalase activity using horseradish peroxidase and 3,3′-diaminobenzidine as described by Mullen and Gifford (
). The active enzyme is a tetramer made up of four identical or similar subunits encoded by the same or different family members, respectively. Therefore, plants contain multiple isoforms of this enzyme. In tobacco, Zelitch et al. (
) detected 6-12 isoforms. We have extended this isoform analysis using purified tobacco leaf catalase. When purified tobacco catalase was subjected to analytical chromatofocusing on a Mono P column, multiple peaks were obtained (Fig. 1A). It should be noted that the use of purified catalase allowed for direct detection of the isoforms (absorbance at 280 nm), in contrast to an indirect approach based on an activity profile. At least 10 catalase species eluted between pH 7.6 and 6.0. SDS-PAGE analysis indicated that the most basic and most acidic isoforms consisted exclusively of larger (57-kDa) and smaller (55-kDa) subunits, respectively, whereas some intermediate isoforms in fractions 15-17 appeared to be heterotetramers (visible in the inset of Fig. 1B). Interestingly, the specific activity of homotetramers consisting only of 57-kDa subunits was much higher (124 units mg−1 for the pooled fractions 8-11, hereafter referred to as pool 1) than that of isoforms containing only the 55-kDa subunits (18.4 units mg−1 for the pooled fractions 19-21, hereafter referred to as pool 2). As a consequence, the elution profile shown in Fig. 1A does not reflect the distribution of catalase activity among the various isoforms (Fig. 1C).
The relative catalase activity of selected fractions, as well as their sensitivity to SA is shown in Fig. 1C. Isoforms consisting only of large subunits accounted for 75-80% of the overall catalase activity. Catalase activity throughout the tested fractions was inhibited 36-51% by 1 mM SA. Thus, there appears to be little difference in SA sensitivity between isoforms, regardless of their subunit composition.
In addition to degradation of H2O2 to H2O and O2, catalase has a peroxidative activity (see Fig. 3); the ratios of these two activities can differ substantially among isoforms (Havir and McHale,
). For tobacco leaf catalase, the ratios of the peroxidative activity to the catalatic activity were 0.42 × 10−4 for pool 1 and 4.5 × 10−4 for pool 2. Note that the peroxidative-like activity was measured with ethanol rather than with true peroxidase substrates such as guaiacol or pyrogallol, which are poor electron donors for catalase.
When crude extracts from tobacco leaves were subjected to isoelectric focusing on nondenaturing gels followed by activity staining for catalase, 8-10 isoforms were detected (Fig. 2B). Thus, the number of isoforms in the purified catalase preparation closely matched that detected in crude extracts. Furthermore, we investigated the isozyme pattern of transgenic tobacco plants expressing the catalase 1 (cat1) or catalase 2 (cat2) gene in an antisense (AS) orientation (construction of the transgenic plants will be described elsewhere).
H. Takahashi, Z. Chen, Y. Liu, and D. F. Klessig, unpublished results.
ASCAT2 plants were devoid of the smaller (55-kDa) subunit (Fig. 2A) and did not contain the acidic isozymes (Fig. 2B). In ASCAT1 plants, the larger (57-kDa) subunit was absent. As a consequence, the majority of basic and neutral isoforms was missing. In agreement with the isoform pattern shown in Fig. 1, the larger subunit encoded by cat1 assembles into at least five different isoforms. The activity profile of the chromatofocused, purified catalase (Fig. 1C) mimics the catalase activity staining pattern of isoelectric focused crude extracts (Fig. 2B), with the majority of the activity represented by isoforms with neutral or basic isoelectric points and consisting of 57-kDa subunits. Therefore, the majority of the isoforms and their relative abundance appear to have been retained during purification.
Mode of Action of SA on Catalase
The first step in the catalase cycle (Fig. 3) involves a two-electron (e−) equivalent reduction of H2O2 to H2O and the corresponding oxidation of ferric enzyme (ferricatalase) to compound I, a spectroscopically distinct and enzymatically active form of catalase (for review see Deisseroth and Dounce (
)). Compound I is converted back to ferricatalase by a 2e− equivalent reduction and the corresponding oxidation of a second molecule of H2O2 to O2, thus completing the extraordinarily rapid catalatic cycle (Fig. 3, steps 1 and 2). It is this “α activity” that makes catalase one of the fastest enzymes known. However, compound I can be siphoned from the catalatic cycle by its conversion into the inactive (with respect to the catalatic cycle) ferryl intermediate compound II through a 1e− equivalent reduction (Fig. 3, step 3). By a second 1e− equivalent reduction, compound II can be converted back to the ferricatalase (Fig. 3, step 4). Because of the slow turnover number of this peroxidative cycle (β activity), the activity of catalase depends on the relative frequency of conversion of compound I back to ferricatalase versus its conversion to inactive compound II (and the subsequent conversion of compound II to ferricatalase). Thus, any enhancement of the β activity, as described previously for phenolic compounds (Schonbaum and Chance,
), would result in inhibition of catalase activity.
The effects of H2O2 and SA on catalase activity were analyzed (Fig. 4). In order to more closely mimic the in vivo situation where H2O2 is almost continuously being produced, low to moderate levels of H2O2 were generated at a prescribed rate during the catalase reaction using various concentrations of glucose and glucose oxidase. The effects of continuous H2O2 fluxes on catalase (inactivation) have been studied in detail by several groups (Kirkman et al.,
). This H2O2-generating system, however, could not be used to attain high levels of H2O2, since all commercial glucose oxidase preparations contain low levels of other contaminating enzymes that affect O2 and H2O2 metabolism. Therefore, reaction mixes were adjusted to millimolar levels of H2O2 with 30% H2O2 stocks at the start of the reaction as described by Deisseroth and Dounce (
)) to study inactivation of catalase in vitro. Under these conditions, high H2O2 levels are maintained for only short periods after the addition of catalase; however, this method is commonly used to assay the susceptibility of catalase to H2O2 (Feuers et al.,
). This is, in part, due to H2O2-mediated accumulation of compound II (at low H2O2 concentrations) and conversion of compound II to compound III (at high H2O2 concentrations), an inactive form of catalase that cannot be easily converted back to active enzyme (Fig. 3; see Schonbaum and Chance (
). At a relatively low rate of H2O2 production (0.1 nmol ml−1 min−1), the addition of SA (0.2 mM) inhibited catalase about 50% compared with the control, which exhibited only very limited activity loss during the 200-min incubation. When the rate of H2O2 generation was increased 10-fold, SA accelerated the inactivation of catalase by H2O2. However, eventually (at ∼200 min) the catalase activity dropped to approximately 50% of the initial level, regardless of the presence of SA. At high levels of H2O2 (i.e. an initial concentration of 10 mM), SA again initially accelerated the rate of catalase inactivation, but as the reaction time increased, SA protected the enzyme against further inactivation. Because of the different kinetics of catalase inactivation by H2O2 and SA, dramatically different results were obtained with respect to the effects of SA on catalase activity depending on the concentration of H2O2 present. This is illustrated in Fig. 5, A and B, in which the effect of SA (0.2 mM) on purified tobacco catalase was measured after a 3-h incubation at different levels of H2O2. Qualitatively similar results were obtained after a 1-h incubation (data not shown). Note that the values are given in percentage of catalase activity of a control without SA. In the absence of added or generated H2O2, SA had relatively little effect on activity, probably because most of the enzyme was in the ferricatalase form (see Fig. 3; see below). At low levels of H2O2, inhibition by SA approached 50% or more. As H2O2 levels were increased, the amount of inhibition by SA decreased (Fig. 5A). At high H2O2 levels, catalase activity was up to 2.5-fold higher in the presence of SA, consistent with SA protecting the enzyme from inactivation by H2O2 (Fig. 5B). Lower concentrations of SA (0.05 mM and 0.1 mM) also provided significant protection (1.8- and 2-fold of the activity in the absence of SA, respectively) against inactivation at high levels of H2O2 (10 mM). It should be noted that desferrioxamine (a strong chelating agent and an inhibitor of the Fenton reaction) did not show any protection of catalase (data not shown).
To help elucidate how SA can both inhibit and activate catalase, we examined the effects of SA on formation of the various redox states or reaction intermediates of catalase (Fig. 3) that can be distinguished spectroscopically by their absorption spectra in the Soret (near UV) region. The absorption spectrum in the Soret region of the purified enzyme is shown in Fig. 6A (curve 1). The broad peak centered around 405 nm is consistent with the enzyme existing primarily in the ferricatalase (Fe(III)) state. After incubation of the enzyme with H2O2 (generated at 1 nmol ml−1 min−1) for 1 h, the curve flattened slightly (curve 2), consistent with the conversion of some of the ferricatalase to compound I (Fe(V)), which has a lower extinction coefficient than ferricatalase. In contrast, when the enzyme was incubated with H2O2 plus 0.2 mM SA for 1 h, there was increased absorbance at 420-440 nm, which is characteristic of compound II, and decreased absorbance at 405 nm, consistent with a reduction in the amount of ferricatalase and compound I (Schonbaum and Chance,
). However, this spectral change induced by SA suggests that it is serving as an electron donor for conversion of compound I to compound II (Fig. 3, step 3).
The ability of SA to serve as an electron donor for conversion of compound II back to ferricatalase (Fig. 3, step 4), thus completing the peroxidative cycle, was then analyzed. To address this question, we chose conditions in which some of the enzyme was trapped in the compound II state and then determined whether the addition of SA would convert the trapped compound II to ferricatalase. A similar approach has been carried out in order to show the effect of NADPH on bovine catalase (Jouve et al.,
; see below). During the catalatic cycle (Fig. 3, steps 1 and 2), a small amount of compound I can be spontaneously converted into compound II (step 3), even within a short period and even in the absence of an electron donor (Schonbaum and Chance,
; Deluca et al., 1995). Since this spontaneous generation of compound II is low compared with that seen when an electron donor like SA is present, it is difficult to follow compound II formation as an increase in absorbance at 420-440 nm. However, its formation can be surmised from a modest reduction in absorbance at 405 nm and a small shift to longer wavelengths. To facilitate spontaneous formation of compound II, purified tobacco enzyme was allowed to react with modest levels of H2O2 (250 μM) three times. Thus, H2O2 was exogenously provided as described recently (DeLuca et al.,
). The reduction in absorbance at 405 nm of the reacted catalase mixture is evident when curve 1 in Fig. 6B is compared to curve 1 in Fig. 6A, which represents the absorbance of purified catalase before reacting with H2O2. It should be noted, however, that in the presence of small amounts of H2O2, catalase will exist as a mixture of all intermediates. The addition of 0.2 mM SA to the reacted catalase mixture without the addition of more H2O2 led to increased absorbance at 405 nm (Fig. 6B, curves 2-4). This increase is consistent with reformation of ferricatalase (Jouve et al.,
). Thus, SA must act as an electron donor for compound II, as well as for compound I.
Interestingly, two of the three commercial preparations of mammalian catalase (one of two bovine liver preparations and one from human erythrocytes), showed similar increases in absorbance at 405 nm with the addition of SA, even without pretreatment with H2O2 (data not shown). This is reminiscent of the observation of others that many catalase preparations consist of a mixture of intermediates (Deisseroth and Dounce,
). We conclude from these results that SA can serve as electron donor for the peroxidative or β activity of plant and animal catalases.
Many catalase inhibitors, representing different classes of chemicals, have been reported, and the modes of action of several of these, such as 3-aminotriazole, resorcinol, ascorbate, and dithiothreitol have been described (for review see Schonbaum and Chance (
)). The efficacy of inhibition of tobacco catalase by SA and related chemicals is compared with that of various traditional catalase inhibitors in Table I. This data also allows for a pharmacological comparison of tobacco catalase with published results on catalase from other species. In general, chemicals that are biologically active for induction of PR-1 gene expression and enhanced disease resistance in plants, which include SA, aspirin, 2,6-dihydroxybenzoic acid, 4-chloro-SA, 3-chloro-SA, 3,5-chloro-SA, and benzoic acid (Chen et al.,
), were good inhibitors, while biologically inactive chemicals, which include 3-hydroxybenzoic acid and 4-hydroxybenzoic acid, were not. There are two noted exceptions. Catechol is a good inhibitor of catalase, as reported previously (Bi et al.,
), but it is biologically inactive. These results suggest that SA and its active analogues may have additional effects besides inhibition of catalase that contribute to their biological activity. In this regard it is noteworthy that SA and its active analogues induce lipid peroxidation. Lipid peroxides can induce PR-1 gene expression.
). In contrast to catechol, INA, which is a potent inducer of PR-1 genes and enhanced resistance, failed to inhibit purified catalase. While INA might also act through a mechanism other than, or in addition to, inhibition of catalase, an alternative explanation is that INA needs to be metabolized to an active form that both inhibits catalase and induces defense responses. This view is supported by the observation that INA is a very effective inhibitor of tobacco catalase in vivo, while in crude tobacco extracts its inhibition is less pronounced (Conrath et al.,
SA, aspirin, 2,6-DHBA, 4Cl-SA, 3Cl-SA, 3,5-Cl-SA, benzoic acid, and INA are biologically active for induction of PR gene expression and disease resistance, while 3-HBA, 4-HBA, and catechol are not (Chen et al., 1993a, 1993b; Gaffney et al., 1993; Conrath et al., 1995).
0.08 μM catalase (pool 1 from Fig. 1) was incubated for 1 h in presence of 0.4 mM inhibitor, except were indicated differently. H2O2 production by the glucose/glucose oxidase system was adjusted to 1 nmol ml−1 min−1. Catalase activity was measured as described. Means ± S.E. are shown, with n = 3. Similar results were obtained for SA, catechol, and INA with pool 2, which consisted of isoforms made up of subunits encoded by the cat2 gene.
a SA, aspirin, 2,6-DHBA, 4Cl-SA, 3Cl-SA, 3,5-Cl-SA, benzoic acid, and INA are biologically active for induction of PR gene expression and disease resistance, while 3-HBA, 4-HBA, and catechol are not (Chen et al.,
b 0.08 μM catalase (pool 1 from Fig. 1) was incubated for 1 h in presence of 0.4 mM inhibitor, except were indicated differently. H2O2 production by the glucose/glucose oxidase system was adjusted to 1 nmol ml−1 min−1. Catalase activity was measured as described. Means ± S.E. are shown, with n = 3. Similar results were obtained for SA, catechol, and INA with pool 2, which consisted of isoforms made up of subunits encoded by the cat2 gene.
In addition to traditional catalase inhibitors, SA, and related chemicals, two drugs, bumetanide and phlorizin, were analyzed for their effect on catalase (Table I). Both compounds have recently been reported to interact with mammalian catalase. The diuretic drug, bumetanide, is an inhibitor of the mammalian Na+/K+/Cl− cotransporter and has recently been shown to bind to membrane-associated catalase from liver (Ottallah-Kolac, 1995). Bumetanide very effectively inhibited tobacco catalase (activity was inhibited by more than 50% with as little as 50 μM). Phlorizin is an inhibitor of another cotransporter, the Na+/glucose cotransporter from kidneys, and it binds at the NADPH-binding site of mammalian catalase (Kitlar et al.,
), to date NADPH-binding catalases have not been detected in plants. However, the inhibition of tobacco catalase by phlorizin and the affinity of the enzyme for the NAD(P)H analogue Cibacron blue (Chen et al.,
), were used as a positive control. After excitation, bovine catalase exhibited a peak of fluorescence centered at 430 nm (Fig. 7, curve 1b). Pretreatment of this catalase with NADPH to saturate the binding sites increased the fluorescence, as expected (curve 1a). Purified tobacco catalase gave a similar but lower peak of fluorescence (curve 2b), which was again elevated by pretreatment with NADPH (curve 2a). In contrast, catalase from A. niger, which does not have NADPH-binding sites (Hillar et al.,
) have shown that NADPH protects mammalian catalases from H2O2 inactivation, which can occur even at low concentrations of H2O2. Since tobacco catalase also contains NADPH-binding sites, it was of interest to know if this cofactor could also protect the plant catalase from inactivation by H2O2. In Fig. 8A, incubation of tobacco or bovine catalase in the presence of a relatively low rate of H2O2 production (1 nmol ml−1 min−1) for 1 h resulted in a slight loss of activity, whereas in the presence of 4 μM NADPH (supplied initially as NADPH and maintained by a regenerating system) catalase activity was enhanced. Since catalase is purified in the absence of exogenous NADPH, the enhancement of initial activity by NADPH may be due to saturation of the NADPH-binding sites. With 0.2 mM SA, catalase (both bovine and tobacco) was inhibited 35-45%, regardless of the presence of NADPH, indicating that SA and NADPH are affecting catalase differently. At a high concentration of H2O2 (10 μmol ml−1, initial concentration), catalase activity of the controls (without SA and NADPH) decreased dramatically (Fig. 8B; also see Fig. 4C), again likely due to formation of compounds II and III. Both SA and NADPH protected catalase to some extent, and when applied together, the protective effect was additive. Thus, at low H2O2 levels, SA inhibits both enzymes, while at high levels of H2O2, it protects them from almost complete inactivation. In contrast, NADPH protects both enzymes regardless of the H2O2 concentration. These data are consistent with the proposed role of SA as an electron donor for the β activity of catalase and with the protective effect afforded by the binding of NADPH to catalase (Kirkman et al.,
). However, despite extensive biophysical, biochemical, and genetic analyses, there is an ongoing discussion as to whether the only, or even major, role of this very abundant protein is to convert H2O2 to H2O and O2 (its catalatic or α activity). This may, in part, reflect the complexity of catalase's redox chemistry. In recent years, catalase has gained renewed attention. There is increasing interest in the involvement of oxidative stress in environmental pollution, aging, diabetes, cancer, and other human diseases and in catalase's role as one of the main antioxidative enzymes. In particular, this has led to renewed interest in the mechanism of catalase inhibition and inactivation (Feuers et al. (
). Given the multiplicity of isozymes in tobacco, we wanted to know, first, whether heterotetramers were primarily responsible for this diversity and, second, whether the various isozymes had different sensitivity to SA. We found that the majority of the 10 or more isoforms in tobacco leaves were homotetramers rather than a mixed population of heterotetramers (Fig. 1, Fig. 2). The nature of the probable posttranslational modification(s) responsible for the differences in charge among isoforms is not known. Alternatively, some of the forms may result from in vitro modifications such as oxidation of sulfhydryl groups, which may have occurred during purification, handling, and storage (Mörikofer-Zwez et al.,
). However, the similar number of forms seen in the crude extracts (Fig. 2) suggests that most of these represent true isoforms rather than in vitro artefacts. The different forms present in purified tobacco catalase exhibited similar sensitivity to SA (36-51% inhibition by 1 mM SA, Fig. 1C). This level of inhibition, while slightly lower, is similar to that reported previously for crude tobacco leaf extracts (45-70%; Chen et al.,
Mechanism of SA Action on Tobacco and Mammalian Catalases
The biphasic kinetics of catalase inhibition by SA (Fig. 4) have been reported for other phenolic compounds such as hydroquinone and pyrogallol and have been interpreted as the transition from the fast catalatic or α activity to the slow peroxidative or β activity (Goldacre and Galston (
). Further evidence that SA acts as a typical phenolic by stimulating the peroxidative activity of tobacco catalase at the expense of its catalatic activity was provided by spectral analysis of catalase and its reaction intermediates (Fig. 6). Together, these analyses indicate that SA acts as an electron donor for the enzyme intermediates compound I and compound II. This is consistent with previous studies, which demonstrated that phenolics can reduce compound I to compound II and compound II to the ferric enzyme (reviewed by Deisseroth and Dounce (
). In other words, SA inhibits catalase by acting as a one-electron donor that siphons compound I from the extremely fast catalatic cycle (see Fig. 3) into the relatively slow peroxidative cycle (∼1000 times slower) (Havir and McHale,
), as part of their studies on the effects of carboxylic acids on catalase, were the first to demonstrate that SA (at extremely high levels of ≥10 mM) inhibited mammalian catalases and to speculate that the inhibition probably resulted from promotion of the peroxidative reaction rather than from chelation of the heme iron of catalase as has been suggested by others (e.g. Rüffer et al.,
) proposed that catalase contains a novel binding site on its surface based on structural similarities to the calycin superfamily. They suggested that SA inhibits catalase by binding to this site and causing a conformational change (allosteric inhibition). While such a site is not inconsistent with data presented here, our results argue that SA inhibits catalase by acting as an electron donor rather than by inducing a conformational change. SA could bind to this surface site and still act as an electron donor to the deeply buried heme of catalase, since Bonagura et al. (
). It also illustrates the difficulties that can be encountered when determining the effects of potential inhibitors like SA on this complex enzyme, whose reaction chemistry is still debated. H2O2 itself can dramatically alter the effects seen with SA as illustrated in Figs. 4, 5, and 8. This likely is responsible for some of the discrepancy in results recently reported (Sánchez-Casas and Klessig,
) is adequate for determining relative catalase activity in different tissues. However, it is poorly suited to analyze the effects of potential inhibitors on catalase activity. Inhibition by phenolics is time-dependent and requires H2O2 (Fig. 4), just as has been previously described (Ogura et al.,
), even modest effects on the activity of these two major H2O2-scavenging enzymes could feed back to cause further inactivation of catalase by the slow and time-dependent accumulation of H2O2.
The role of catalase and SA in uninfected parts of an infected plant is considerably less clear. While SA also accumulates in these tissues, the level appears to be far below the concentration required to effectively inhibit catalase and ascorbate peroxidase (Malamy et al.,
). SA's role in SAR development is unlikely to involve elevated levels of H2O2 resulting from its inhibition of catalase, as originally proposed, unless SA is highly concentrated in certain subcellular compartments (Chen et al.,
). Nonetheless, SA induction of SAR may be mechanistically coupled to its interaction with catalase and peroxidases. SA serves as a one-electron donor for catalase (Fig. 6) and peroxidases (Durner and Klessig,
) and in so doing is converted to a free radical. Free radicals of phenolics (e.g. Savenkova, et al. (1994)) can initiate formation of lipid peroxides. Our preliminary studies indicate that SA induces lipid peroxidation, while several naturally occurring lipid peroxides activate PR-1 genes in tobacco cells.3 A SA free radical could result in the formation of an effective lipid peroxide signal without readily discernible inhibition of catalase. However, the biological significance of a SA radical generated by catalase remains to be proven.
In addition to its ability to inhibit catalase, SA could also protect plant and mammalian catalases against inactivation by H2O2in vitro (Figs. 4C, 5, and 8). This is functionally similar to the protective effects of NADPH on mammalian catalases in the presence of small fluxes of H2O2 as described by Kirkman et al. (
). Indeed, we found that tobacco catalase, like mammalian catalases, contains bound NADPH (Fig. 7). Therefore, it appears that, under some conditions, SA can support or substitute for NADPH's protective role. Might it serve a similar function in vivo? In animal systems accumulation of compound II (and thus catalase inhibition) has been associated with “abnormal” stress conditions such as found in tumors or during prolonged hypoxia or cell necrosis (Oshino et al.,
). In the case of plants, similar stress conditions may occur during necrotic lesion formation in the HR. A strong oxidative burst (probably produced by a NADPH oxidase) is associated with the HR (Doke and Ohashi,
). Catalase inactivation during the HR would be enhanced by the proposed depletion of NADPH by NADPH oxidases and antioxidative enzymes of the ascorbate/glutathione cycle (Mehdy et al., 1994). One might speculate that under these conditions, SA may protect or reactivate a basal catalase activity. This notion is consistent with the observation that SA appears to act as an antioxidant at sites of inflammation in animals (Halliwell et al.,
); one property of SA may be to maintain a basal level of catalase activity by acting as an electron donor that converts inactive compound II to the active ferricatalase (Fig. 6). In fact, it has been suggested that SA protects various heme proteins such as leghemoglobin and metmyoglobin from H2O2-induced inactivation by maintaining the peroxidative cycle of these O2-binding proteins (Galaris et al.,
). SA is a direct scavenger of OH· (in vitro and in vivo), and it is a iron-chelating compound, thereby inhibiting the direct impact of OH· as well as its generation via the Fenton reaction (Halliwell et al.,
). However, since desferrioxamine (a strong chelating agent) did not protect catalase from inactivation, we hypothesize that SA maintains a basal catalase activity through its ability to serve as an electron donor.
In sum, whether SA positively or negatively modulates catalase activity will depend on the redox status of the cell. In the healthy tissue surrounding, but not immediately adjacent to, the infection site H2O2 concentrations will be relatively low to moderate, and the elevated SA levels probably inhibit catalase by promoting the slow peroxidative cycle (note that in normal healthy leaf tissue H2O2 has been estimated at ∼100 nM; Scandalios (
) could lead to substantial inactivation of catalase by accumulation of inactive enzyme intermediates. Under conditions of such oxidative stress, SA might help to maintain and/or reestablish a basal level of catalase activity. These protective, antioxidative properties of SA might serve to limit the impact of the oxidative processes associated with development and spread of the lesion.
We thank members of the laboratory, particularly D'Maris Dempsey and Marc D. Anderson for critical reading of the manuscript. Transgenic plants expressing the cat2 gene and the cat1 gene in an antisense orientation, respectively, were kindly provided by Hideki Takahashi (this laboratory). Helmut Kessmann, Theo Staub, and John Ryals are acknowledged for generously providing INA.