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J. Biol. Chem., Vol. 282, Issue 2, 929-937, January 12, 2007
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From the Department of Microbiology, University of Illinois at Urbana-Champaign, Urbana, Illinois 61801
Received for publication, August 10, 2006 , and in revised form, November 9, 2006.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Exogenous H2O2 rapidly diffuses across cell membranes (2) and can impose a much more acute stress on cells; accordingly, it is often used as a biological weapon. For example, H2O2 is formed by phagocytes and may accumulate to 10–4 M inside phagosomes that have engulfed invading bacteria. Lactic acid bacteria suppress the growth of competitors by releasing H2O2 as a primary metabolic product, achieving millimolar concentrations in laboratory cultures. And redox-cycling antibiotics, which are produced as microbicides by both plants and bacteria, suffuse target organisms with a continuous stream of H2O2.
If we wish to understand the severity and nature of the stress that H2O2 imposes upon cells, we must identify the biomolecules with which it primarily reacts. This problem has not been easy to solve. In vitro studies have shown that H2O2 can oxidize methionine (3) and cysteine (4) residues, but the rates at which it does so suggest that these types of damage will be scant at physiological doses of H2O2, unless the surrounding polypeptide context somehow strongly activates the residues. Reactions between H2O2 and loosely bound iron generate hydroxyl radicals (Reaction 1) and are suspected of being involved in protein carbonylation, lipid peroxidation, and DNA oxidation (5).
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An alternative approach to pinpointing the important targets of H2O2 is to expose cells to increasing doses in a way that identifies the first cell processes to fail. Escherichia coli and other organisms have calibrated their defensive systems to detect submicromolar levels of H2O2 (2, 9), so we anticipate that these low concentrations are sufficient to threaten the most sensitive biomolecules. Unfortunately, if scavenging enzymes are active, it is difficult to impose such a low dose of H2O2 over an extended period of time because the enzymes will degrade the H2O2 and end the stress. Therefore, these experiments are most easily conducted with scavenger-deficient mutants. We have constructed E. coli strains that lack peroxidase and catalase activities (10). These mutants grow at wild-type rates in anaerobic environments, but when they are exposed to oxygen, they grow at reduced rates in complex medium and fail to grow at all in minimal medium. The former defect is due, at least in part, to Fenton-mediated DNA damage (8). The second defect stems from problems with biosynthetic pathways. In this study, we identified the mechanism by which micromolar H2O2 blocks leucine biosynthesis, and we found that this class of injury affects multiple pathways in the cell.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-ketoisovalerate (TCI America) and -ketoisocaproate (Sigma), were present at 0.5 mM.
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Aerobic Cell Growth—To ensure that cells were growing exponentially before they were exposed to oxygen, anaerobic overnight cultures of hydroperoxidase-deficient (Hpx–)2 cells were diluted to A600 = 0.005 in fresh anaerobic glucose minimal medium. Cells were then grown anaerobically to A600 
0.1 prior to dilution into aerobic medium.
Enzyme Assays—Cell extracts were prepared by suspending and sonicating cells in anaerobic buffers inside an anaerobic chamber. Isopropylmalate isomerase (IPMI) activity was measured by monitoring the decrease in the absorbance (235 nm) of citraconate (Sigma) (13), an analog of isopropylmalate; reactions contained 100 mM Tris-Cl (pH 7.6) and 0.4 mM citraconate. Fumarase activity was determined from the appearance of fumarate (250 nm) in a reaction containing 50 mM sodium phosphate (pH 7.4) and 50 mM malate (Sigma) (14). To assay 6-phosphogluconate dehydratase, lysates were prepared from cultures grown in minimal medium containing 0.2% gluconate. Turnover of 6-phosphogluconate dehydratase produces pyruvate; its formation (in 50 mM Tris-Cl (pH 7.65)) was determined in a second reaction catalyzed by lactate dehydrogenase (Sigma) (15). To assay NADH dehydrogenase I, inverted membrane vesicles were isolated from extracts after sonication in 50 mM MES (pH 6.0) (16). NADH dehydrogenase I oxidizes deamino-NADH (340 nm) in 50 mM potassium phosphate buffer (pH 7.8), whereas NADH dehydrogenase II does not. Aconitase was assayed by the conversion of isocitrate to aconitate (17).
Plasmid Construction—The leuCD open reading frame was PCR-amplified from E. coli MG1655 using the forward primer 5'-ATATCGAATTCTTAACGATATCATTGCCCGCTATATAGCAG-3' and the reverse primer 5'-CTGGATCTAGATTAATTCATAAACGCAGGTTG-3'. To construct the pLEUCD2 plasmid, the PCR products were digested with XbaI and EcoRI and cloned into the pWKS30 vector (18) behind the lac promoter. The plasmid was confirmed by restriction analysis and IPMI assay. Hpx– cells containing pLEUCD2 or pWKS30 were cultured in lactose minimal medium with histidine, aromatic amino acids, and 50 µg/ml ampicillin. This plasmid was used in complementation experiments; the leuCD genes were induced 3-fold above wild-type levels by 1 mM isopropyl 
-D-thiogalactopyranoside, which was added to both anaerobic precultures and final aerobic cultures.
For EPR experiments and purification purposes, IPMI and fumarase A were strongly overproduced by expression from a tac promoter. The leuCD genes were excised from pLEUCD2 with XbaI and EcoRI and cloned into the pCKR101 vector to construct pLEUCD3. The fumA open reading frame was PCR-amplified from E. coli MG1655 using the forward primer 5'-ATATCGAATTCTTAACATAACCAAACCAGGCAGTAAGTG-3' and the reverse primer 5'-CATGGATCTAGATTATTTCACACAGCGGGTGCATTG-3'. PCR products were digested with XbaI and EcoRI and cloned into the pCRK101 vector to construct pFUMA. Cells containing the plasmids were cultured in glucose minimal medium with histidine, aromatic amino acids, and 50 µg/ml ampicillin. These plasmids overproduced IPMI and fumarase A >60-fold above wild-type levels.
Inactivation of Enzymes—Hpx– cells were grown anaerobically to A600 = 0.2. H2O2 was added to the cultures when they were aerated. At designated time points, aliquots were removed; catalase was added to 200 units/ml; and cells were returned to the anaerobic chamber for lysis and assay.
In vivo inactivation of IPMI by endogenous H2O2 was initiated by aerating heretofore anaerobic cultures without any addition of exogenous H2O2. Under these conditions, cells steadily generate H2O2. The H2O2 equilibrates so quickly across membranes that, in Hpx– cultures, the intracellular H2O2 concentration is essentially equivalent to the extracellular concentration (2). Extracellular H2O2 was measured directly by the Amplex Red/horseradish peroxidase method (2). At the same time, extracts were prepared and assayed in the anaerobic chamber.
Inactivation of enzymes in vitro was accomplished by the addition of H2O2 to lysates or to purified enzyme in anaerobic buffer. The H2O2 was subsequently removed by catalase prior to anaerobic assay. In some cases, damaged iron-sulfur clusters were chemically rebuilt by incubation with 50 µM Fe(NH4)2(SO4)2 (Sigma) and 2.5 mM dithiothreitol (Sigma) (17) at room temperature.
EPR Analysis—In vivo EPR samples were prepared with Hpx– cells overproducing IPMI or fumarase A. To overexpress the structural genes, 1 mM isopropyl -D-thiogalactopyranoside was added when the cells reached A600 = 0.2. After another 2 h of incubation, the cells were harvested by centrifugation, and the cell pellets were resuspended at 
of the original culture volume in 10% glycerol. The resuspended cells were incubated with H2O2 for 1 min at 37 °C. The cell suspension (250 µl) was then transferred into an EPR tube and frozen in dry ice. EPR spectra of [3Fe-4S]+ clusters (19) were obtained at the following settings: microwave power, 1 milliwatt; microwave frequency, 9.05 GHz; modulation amplitude, 8 G at 100 KHz; and time constant, 0.032.
Purification of Enzymes—Four-liter cultures of the Hpx– mutant overproducing fumarase A were grown anaerobically to A600 = 0.2 at 37 °C. Expression of fumA was induced by incubation with 1 mM isopropyl -D-thiogalactopyranoside for 2 h at 37 °C. The cultures were harvested by centrifugation; the cell pellets were resuspended in 15 ml of anaerobic 50 mM Tris-Cl and 10 mM Mg2+ (pH 8) and disrupted by sonication; and cell debris was removed by centrifugation. The purification procedure was derived from the protocol described by Flint et al. (20). All steps of the purification were conducted in an anaerobic chamber at room temperature, and all buffers were anaerobic. Protamine sulfate (1%) was added to the supernatant to remove nucleic acids. The supernatant was then loaded onto a DEAE-Sepharose column (16 x 140 mm) and eluted by a 0–1 M gradient of KCl in 50 mM Tris-Cl and 10 mM Mg2+. The fractions containing the highest activity were pooled. Ammonium sulfate (1.5 M) was added; the precipitate was removed by centrifugation; and the supernatant was loaded onto a phenyl-Sepharose column (16 x 110 mm). The fractions were eluted by a 1.5 to 0 M gradient of ammonium sulfate in 50 mM Tris-Cl and 10 mM Mg2+. The fractions containing the highest activity were pooled and concentrated (Amicon). The enzyme was loaded onto a Superdex column (16 x 50 mm). The fractions were eluted with 500 mM KCl in 50 mM Tris-Cl and 10 mM Mg2+. The three fractions containing the highest activity were frozen in dry ice containing ethanol. Purified fumarase A was >90% pure based on SDS-PAGE. The enzyme was quantified by dye binding assay using bovine serum albumin as a standard and the conversion factor determined by Flint et al. (20). Aconitase A was overproduced and purified as described (17).
Detection of Iron Released upon Cluster Oxidation—EPR spectroscopy was used to quantitatively correlate the creation of [3Fe-4S]+ clusters with the release of iron when fumarase A was damaged in vitro. Desferrioxamine (1 mM; Sigma) was added to purified fumarase A (18 µM) immediately prior to the addition of H2O2. The sample was then transferred into an EPR tube and frozen. The [3Fe-4S]+ signal was detected as described above, whereas EPR spectra of free iron were obtained at the following settings: microwave power, 10 milliwatt; microwave frequency, 9.05 GHz; modulation amplitude, 12.5 G at 100 KHz; and time constant, 0.032. Iron concentrations were quantified using standard solutions of FeCl3 (Sigma) in 50 mM Tris-Cl containing 10 mM Mg2+ and 1 mM desferrioxamine. Desferrioxamine binds both ferric and ferrous iron and triggers the oxidation of the latter species. Thus, the EPR method did not distinguish whether the released iron was in the ferric or ferrous form.
Ferene is commonly used to detect ferrous iron released by metalloenzymes, but we found that ferene itself directly inactivated fumarase A. Dipyridyl did not. Dipyridyl binds ferrous iron in a complex that exhibits an absorbance maximum at 522 nm (21); we determined that this complex cannot be oxidized by H2O2. To detect Fe2+ released during cluster decomposition, 1 mM dipyridyl (Sigma) was added to 18 µM fumarase A prior to H2O2 addition. Control experiments confirmed that this concentration of dipyridyl captured ferrous iron before H2O2 could oxidize it: when Fe(NH4)2(SO4)2 was added to the H2O2-containing reaction mixture, we were able to quantitatively recover it as a dipyridyl chelate, and we could accurately detect as little as 2 µM.
Protein Mass Spectroscopy—Fumarase A (3 µM) was treated with 5 µM H2O2 for 2 min at room temperature. The reaction was terminated by the addition of 0.1 unit/µl catalase. Assays showed that >98% of the enzyme had been inactivated. For mass spectroscopic analysis, 2 µg of sample was desalted using the Genotech Perfect-FOCUS two-dimensional sample cleanup kit. The desalted sample was suspended in 25 mM ammonium bicarbonate containing 12.5 µg/ml trypsin and incubated for 12 h at 37 °C. The sample was then dried, suspended in 5% acetonitrile and 0.1% formic acid, and injected into a Waters Q-Tof quadrupole time-of-flight mass spectrometer via a high pressure liquid chromatography interface. Peptide masses were detected using a data-dependent method and were subjected to tandem mass spectrometry for de novo sequencing and characterization of post-translational modifications. Analysis was done using ProteinLynx Global SERVER Version 2.1 (Waters), Mascot (Matrix Science), and PEAKS (Bioinformatics Solutions Inc.).
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Hpx– growth in the presence of H2O2 was improved by supplementation with exogenous -ketoisocaproate, the final intermediate in the leucine pathway, but not by -ketoisovalerate, the first one (data not shown). There are five reactions between the two intermediates catalyzed by four enzymes. We anticipated that H2O2 might inhibit or inactivate one of them. Indeed, the growth defect was partially relieved by a plasmid that overproduces IPMI 3-fold (Fig. 1B), suggesting that this is the rate-limiting enzyme in the pathway during H2O2 stress.
Enzyme assays confirmed that IPMI lost activity rapidly when 8 µM H2O2 was added to cultures (Fig. 2A). Furthermore, significant enzyme inactivation occurred upon aeration even without the addition of exogenous H2O2 (Fig. 2B), indicating that <0.4 µM intracellular H2O2 was sufficient to poison a substantial fraction of IPMI. Thus, this enzyme is exquisitely sensitive, and scavenging enzymes are needed to protect it from endogenously formed H2O2.
The Nature of IPMI Damage—IPMI activity was lost when H2O2 was added to anaerobically prepared Hpx– extracts (Fig. 3A), indicating that inactivation occurred by direct action of H2O2 upon the enzyme. Catalase protected completely (data not shown).
IPMI is a dehydratase, and its protein sequence suggests that it belongs to a family of enzymes that employ [4Fe-4S] clusters as active-site catalysts (22). The solvent-exposed clusters of these enzymes both coordinate substrate and act as Lewis acids in abstracting the hydroxide anion from the bound substrate (23). These enzymes are typified by aconitase, and they are notoriously sensitive to inactivation by univalent oxidants such as superoxide and ferricyanide (15, 24–27). These agents abstract a single electron from the cluster, converting it to a [4Fe-4S]3+ form. The cluster is unstable at that valence and releases the substrate-binding iron atom as Fe2+ so that the residual cluster is left in a [3Fe-4S]+ form that lacks the key iron atom and is catalytically inactive.
Consistent with this model, the addition of citraconate, a pseudosubstrate of IPMI, protected the enzyme from H2O2 in vitro, suggesting that H2O2 must directly contact the cluster to inactivate the enzyme (data not shown). Damaged clusters can often be reassembled chemically by treatment with dithiothreitol and ferrous iron; when damaged IPMI was subjected to this protocol, 60% of the activity was regained within 3 min (Fig. 3B).
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85% of the IPMI activity was recovered when the extracts from H2O2-exposed cells were treated with dithiothreitol and ferrous iron. Other [4Fe-4S] Dehydratases Are Similarly Sensitive to H2O2— Other cluster-containing dehydratases (6-phosphogluconate dehydratase and fumarases A and B) also lost activity when they were exposed to low concentrations of H2O2 in vitro (Fig. 4, A–C). In each case, full activity could be restored by subsequent treatment with dithiothreitol and ferrous iron (data not shown). Furthermore, a strong [3Fe-4S]+ signal was detected when H2O2 was added to Hpx– cells overproducing fumarase A (Fig. 5). Thus, it appears that H2O2 efficiently damages the clusters of all members of this enzyme family.
Glucose medium, which we employed for our initial growth experiments, does not demand that E. coli process substrate through its tricarboxylic acid cycle to generate ATP; therefore, growth would not have been affected by the inactivation of aconitase or fumarase. However, the Hpx– strain exhibited a severe growth defect when it was cultured in malate medium (Fig. 6). This result indicates that the submicromolar H2O2 that is generated by endogenous processes is sufficient to debilitate this pathway.
Iron-sulfur clusters are also used by respiratory enzymes to transfer electrons between active sites. The NADH dehydrogenase I complex is characteristic of these enzymes, as it contains at least nine iron-sulfur clusters (28). However, even 5 mM H2O2 was unable to diminish its activity (data not shown). Because these clusters are buried within polypeptide, the implication is that H2O2 can oxidize only those clusters that it can contact directly.
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Strikingly, the fumarase rate constants were orders of magnitude higher than the apparent constant that we calculated in vivo using Hpx– mutants. We suspected that substrates protect the enzyme inside the cell. Indeed, both malate and fumarate fully protected fumarase when they were added at saturating concentrations (Fig. 7). The doses needed for half-maximal protection (0.4 mM for malate and 0.3 mM for fumarate, measured at 0 °C) were in reasonable agreement with the Km values of the enzyme for those substrates (0.7 and 0.6 mM, respectively, at 37 °C) (29). Thus, it is likely that the inactivation of the enzyme in vivo is tempered by the consequent accumulation of substrate. These data also support the suspicion that H2O2 must intimately contact the cluster to inactivate it.
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Previous experiments that were conducted with millimolar doses of oxidant indicated that, with time, the cluster disintegration might continue past the [3Fe-4S]+ state (19). However, the ease with which we were able to chemically reactivate damaged dehydratases was inconsistent with the formation of apoprotein, which was only slowly reactivated. In fact, EPR analysis showed that the [3Fe-4S]+ fumarase cluster was unaffected by a 10-min exposure to 100 µM H2O2 (Fig. 8A). We conclude that, over this period, physiological doses of H2O2 do not directly degrade clusters beyond the easily repaired [3Fe-4S]+ state.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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The sensitivity of iron-sulfur clusters to univalent oxidants has been appreciated for some time; in fact, much of the toxicity of superoxide is due to inactivation of the same enzymes that were identified in this study. Damage by millimolar doses of H2O2 has been reported (17, 30). However, although the chemistry is not unexpected, the surprise is that these reactions occur so rapidly. The rate constant of the Fenton reaction was measured to be 76 M–1 s–1 at pH 3, and this number has been cited widely (6). It carried with it the implication that reactions between iron and H2O2 are too slow to occur at an important rate in most biological scenarios, where H2O2 concentrations are micromolar or lower. However, more recent measurements (8) have shown that, at physiological pH, the rate constant for hexa-aqueous iron is actually orders of magnitude higher, presumably because, at neutral pH, hydroxide anion coordinates iron and lowers its reduction potential. Extrapolating from measurements made at lower temperatures (8), we estimate that the rate may be 20,000–30,000 M–1 s–1 at 37 °C. The same enhancement evidently pertains to the substrate-binding iron atom within a dehydratase iron-sulfur cluster.
The biological consequence of this high reactivity is that far less H2O2 is needed to create toxicity than had been suspected. The data directly show that E. coli must synthesize scavenging enzymes to avoid being poisoned by the high nanomolar H2O2 that it makes through the adventitious oxidation of its own redox enzymes. Basal levels of the NADH peroxidase solve that problem. However, this basic vulnerability is still exploited by phagocytes and plants, both of which suppress the growth of invading bacteria by generating far higher levels of H2O2 either directly or through the synthesis of redox-active antibiotics (31–34).
These deliberate oxidative assaults, as well as the H2O2 that accumulates in the environment through chemical oxidation processes, are a true threat to bacteria. Because H2O2 influx across membranes can outstrip the action of scavenging enzymes, extracellular concentrations of 5 µM H2O2 are sufficient to raise the intracellular H2O2 in wild-type (scavenger-proficient) cells to 1 µM (2). Therefore, as little as 5 µM environmental H2O2 is sufficient to cause the enzyme inactivity and pathway defects that were reported in this study.
Interestingly, lactic acid bacteria employ a similar stratagem to inhibit competing organisms, utilizing lactate and pyruvate oxidases, which can drive H2O2 levels in their immediate environment up to millimolar levels (35, 36). How do the lactic acid bacteria themselves tolerate the stress that they are creating? Lactic acid bacteria do not rely upon pathways that contain iron-sulfur dehydratases. Instead, they obtain from their environment the amino acids that dehydratase-dependent biosynthetic pathways would normally produce, and they ferment sugars to lactic acid rather than using the tricarboxylic acid cycle.
To defend themselves against such H2O2-mediated assaults, microbes throughout the biotic world have evolved H2O2-inducible defenses, most of which are either homologs or analogs of the OxyR system of E. coli. The frontline defense of these systems is the strong induction of scavenging enzymes, but the repair of damaged clusters is also an important feature. Ongoing cluster repair may contribute to the disparity between the rates of enzyme damage that we measured in vitro and the apparent rates that we determined in vivo. In a previous study (19), we observed that cells quantitatively repaired H2O2-damaged dehydratases, but we were perplexed as to why the hydroxyl radical, which presumably had been formed upon cluster oxidation, did not irreversibly damage active-site amino acid residues. This puzzle is resolved by the observation that ferric (rather than ferrous) iron is released during cluster oxidation. Therefore, a ferryl radical (or a non-diffusing hydroxyl radical) evidently pulls a second electron from the residual cluster rather than allowing an unexpended hydroxyl radical to be released into the active-site bulk solution. Interestingly, an analogous phenomenon was noted for the PerR protein of Bacillus subtilis in that a Fenton reaction between its prosthetic ferrous iron atom and H2O2 caused immediate oxidation of the iron ligands rather than the more widely distributed oxidation of the polypeptide, as might have been expected from a diffusible hydroxyl radical (37).
In the dehydratase case, we do not stipulate whether the second electron transfer would generate an intermediate all-ferric [4Fe-4S]4+ cluster or whether it would create a disulfide bond concomitant with ferric iron release. However, in either case, polypeptide oxidation would be avoided. The physiological significance is that the enzyme can be restored to its native state by cell processes that reduce and remetallate the [3Fe-4S]+ cluster. The components that catalyze that repair process have not yet been identified. However, the OxyR system responds to H2O2 by inducing the Suf proteins (38), which compose a backup system for cluster assembly.
A secondary consequence of cluster damage is that iron is released in an uncontrolled manner into the cytosol. Loose iron can bind to DNA and catalyze the formation of damaging hydroxyl radicals; therefore, this iron leak has the effect of accelerating the rate at which H2O2 produces DNA damage (39–41). The OxyR system addresses this threat by inducing the synthesis of Dps, a ferritin-like storage protein that scavenges loose iron and sequesters it into an unreactive ferric hydroxide core (8, 42–44). Thus, it has become clear that several aspects of the inducible defense against H2O2 are well matched to the threat that H2O2 poses.
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FOOTNOTES |
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1 To whom correspondence should be addressed: Dept. of Microbiology, University of Illinois at Urbana-Champaign, 601 South Goodwin Ave., Urbana, IL 61801. Tel.: 217-333-5812; Fax: 217-244-6697; E-mail: jimlay{at}uiuc.edu.
2 The abbreviations used are: Hpx–, hydroperoxidase-deficient; IPMI, isopropylmalate isomerase; MES, 4-morpholineethanesulfonic acid.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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) and H2O2 concentrations (
) were monitored.
) and Hpx– (
) cells). All media included histidine and aromatic amino acids.
