From the Department of Microbiology, University of Illinois, Urbana, Illinois 61801
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ABSTRACT |
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The fitness of organisms depends upon the rate at
which they generate superoxide (O The discovery of superoxide dismutase
(SOD)1 in 1969 (1) was the
first indication that aerobic organisms are threatened by superoxide
(O2) and hydrogen peroxide
(H2O2) as toxic by-products of aerobic
metabolism. In Escherichia coli these oxidants arise
primarily from the autoxidation of components of its respiratory chain.
Inverted vesicles that were incubated with NADH generated O
2
and H2O2 at accelerated rates either when
treated with cyanide or when devoid of quinones, implicating an NADH
dehydrogenase as their source. Null mutations in the gene encoding NADH
dehydrogenase II averted autoxidation of vesicles, and its
overproduction accelerated it. Thus NADH dehydrogenase II but not NADH
dehydrogenase I, respiratory quinones, or cytochrome oxidases formed
substantial O
2 and H2O2. NADH
dehydrogenase II that was purified from both wild-type and quinone-deficient cells generated ~130 H2O2
and 15 O
2 min
1 by autoxidation of its reduced
FAD cofactor. Sulfite reductase is a second autoxidizable electron
transport chain of E. coli, containing FAD, FMN,
[4Fe-4S], and siroheme moieties. Purified flavoprotein that contained
only the FAD and FMN cofactors had about the same oxidation turnover
number as did the holoenzyme, 7 min
1 FAD
1.
Oxidase activity was largely lost upon FMN removal. Thus the autoxidation of sulfite reductase, like that of the respiratory chain,
occurs primarily by autoxidation of an exposed flavin cofactor. Great
variability in the oxidation turnover numbers of these and other
flavoproteins suggests that endogenous oxidants will be predominantly
formed by only a few oxidizable enzymes. Thus the degree of oxidative
stress in a cell may depend upon the titer of such enzymes and
accordingly may vary with growth conditions and among different cell
types. Furthermore, the chemical nature of these reactions was
manifested by their acceleration at high temperatures and oxygen
concentrations. Thus these environmental parameters may also directly
affect the O
2 and H2O2 loads that organisms must bear.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
2). SOD catalyzes the dismutation of superoxide (Reaction 1)
and, in combination with catalase (Reaction 2), helps clear the cell of
reactive oxygen species.
SOD was found to be virtually ubiquitous among aerobic organisms,
suggesting that O
2 might be an unavoidable by-product of
metabolism in air. This idea was extended to develop the
hypothesis that oxygen toxicity might be generally mediated by
intracellular O
2 (2). The molecular details that underpin this
idea (the intracellular sources and targets of O
2) were unresolved.
Within the past 10 years many details of oxygen toxicity have been
revealed. In 1986 Carlioz and Touati (3) reported the properties of a
mutant strain of Escherichia coli that lacked both of its
cytosolic isozymes of SOD. The mutant grew normally in the absence of
oxygen; however, in aerobic medium it exhibited requirements for
branched chain, aromatic, and sulfurous amino acids, an inability to
grow on non-fermentable carbon sources, and a high rate of spontaneous
mutagenesis (4). These same traits were elicited when SOD-proficient
wild-type strains were exposed to hyperbaric oxygen (5), suggesting
that in these conditions O2 formation must be accelerated
enough to overwhelm the cellular defenses. Therefore these observations
supported the original model of oxygen toxicity.
The root cause of the branched chain auxotrophy was tracked to the
ability of O2 to inactivate dihydroxy-acid dehydratase, an
enzyme midway in this pathway (6). O
2 does so by oxidizing and
destabilizing the [4Fe-4S] cluster that acts as a Lewis acid during
catalysis (7, 8). Iron dissociates from the oxidized cluster, causing a
complete loss of activity. The requirement for fermentable carbon
sources apparently stems from similar damage to aconitase and fumarase,
which also belong to the [4Fe-4S] dehydratase class (9, 10). The high
rate of mutagenesis is linked to the same damage: the iron that is lost
from destabilized clusters floats freely into the cytosol where it
catalyzes DNA oxidation by H2O2 (11-13). Thus
to date all the well understood deficits of SOD mutants arise from this
single type of lesion. In higher organisms O
2 toxicity is
linked to similar damage to mitochondrial aconitase (14).
Less clear is the mechanism by which O2 is generated in cells
in the first place. Molecular oxygen is actually a poor chemical oxidant, because its triplet state constrains it to accept one electron
at a time from potential donors. Because biological electron carriers
such as NAD(P)H resist the loss of a single electron, and the oxidizing
potential of the O2/O
2 couple is low (
0.18 V
using 1 M O2 as standard state), most electron
traffic is unaffected by the presence of oxygen. However, complex
electron transport chains include redox moieties such as flavins,
quinones, and metal centers that excel at univalent electron transfers
and are therefore plausible electron donors for oxygen. In the seminal
experiments that led to the discovery of SOD, xanthine oxidase served
as an enzymic source of O
2. Xanthine oxidase is actually a
damaged form of xanthine dehydrogenase and has lost the ability to bind NAD+; as a consequence, electrons accumulate on its flavin,
iron-sulfur cluster, and molybdopterin cofactors. In the absence of the
native substrate, the electrons are transferred at a moderate rate from the flavin to dissolved oxygen, and an admixture of
H2O2 and O
2 is produced (15-17). This
example provides evidence that adventitious O
2 (and
H2O2) production can occur quite rapidly but
that the physical and electronic structures of proteins have evolved to suppress it. Although a number of native flavoproteins were examined by
Massey's group (18), none generated O
2 at rates close to that
of xanthine oxidase.
More recently we undertook a systematic effort to identify the primary
sources of endogenous O2 in E. coli. This organism does not have any enzymes that deliberately generate either
O
2 or H2O2; as a facultative
anaerobe, E. coli must maintain fluxes through its pathways
even in anaerobic habitats. In this aspect E. coli is unlike
committed aerobes, which sometimes employ
H2O2-generating enzymes in biosynthetic
pathways. In earlier work Imlay and Fridovich (19) demonstrated that
the respiratory chain is the apparent source of most endogenous
O
2. Gonzalez-Flecha and Demple (20) arrived at the same
conclusion with respect to H2O2. Subsequently fumarate reductase was determined to be the primary source of O
2 in the particular situation of transit from anaerobic to
aerobic environments (21). Although fumarate reductase contains three iron-sulfur clusters, its autoxidation, like that of xanthine oxidase,
occurred exclusively from the flavin. One purpose of the present study
was to determine whether that is a general rule.
Sulfite reductase comprises a second, albeit self-contained, electron
transport chain in E. coli. Like the respiratory chain, sulfite reductase transfers electrons through FAD, FMN, [4Fe-4S], and
siroheme, which are all good univalent redox enzymes and could plausibly transfer an electron to oxygen. In fact its reactivity with
oxygen has been well documented (22-24). By comparing the sites,
rates, and valencies of electron transfers from these carriers to
oxygen, we wished to identify characteristics that may predispose redox
enzymes to generate O2 and/or H2O2. If
these oxidants are preponderantly generated by only a few enzymic
sources, then variations in the titer of such enzymes (from one growth
condition to another or one organism to another) might substantially
change the degree of endogenous oxidative stress.
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MATERIALS AND METHODS |
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Chemicals and Enzymes--
Bovine erythrocyte copper, zinc
superoxide dismutase, horseradish peroxidase, E. coli
manganese superoxide dismutase, NADH, NADPH, deamino-NADH,
acetylpyridine adenine dinucleotide (AcPyNAD+),
acetylpyridine adenine dinucleotide phosphate (AcPyNADP+),
plumbagin, potassium ferricyanide, ubiquinone-1, FAD, FMN, chloromercuriphenyl sulfonic acid, cytochrome c, ADP,
H2O2, ferene, o-dianisidine,
scopoletin, dichlorofluorescein, hydroxylamine, -glycerol phosphate,
fumaric acid, 4-hydroxybenzoate, djenkolic acid, chloramphenicol,
kanamycin, and tetracycline were purchased from Sigma. Catalase was
from Boehringer Mannheim, ferric chloride was from G. Frederich Smith
Chemical, and potassium cyanide and ferrous chloride were from Aldrich.
Deionized house water was further purified with a Labconco Water Pro PS system.
Strains-- Strains used in this study are listed in Table I. Mutant strains were constructed by P1 transduction. All comparisons in this work were between congenic strains. Mutant fnr alleles were co-transduced with a linked tetracycline-resistant Tn10; inheritance of the fnr allele was demonstrated by the inability of mutants to grow anaerobically on glycerol/nitrate medium (25). In a ubi men (quinoneless) background this screen cannot work, so the allele from putative transductants was transduced back out into a quinone-proficient background and screened. The frd deletion was co-transduced with a Tn10 and screened for growth on glycerol/fumarate. ndh::cam alleles were selected by chloramphenicol. The absence of NADH dehydrogenase II was screened by enzymatic assay for NADH:plumbagin oxidoreductase activity after vesicles were incubated at pH 7.8 to inactivate the NADH dehydrogenase I complex (26). The nuo mutant allele was selected by the linked Tn10. Potential NADH dehydrogenase I-deficient transductants were screened for deamino-NADH:plumbagin oxidoreductase activity (27).
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Growth and Media-- Vesicles were prepared from cells grown in either LB medium supplemented with 0.2% glucose, minimal A glucose medium (28), or minimal A supplemented with 1% casamino acids and 0.5 mM tryptophan. Standard antibiotic concentrations were used for plasmid maintenance and P1 transductions (29). Media for anaerobic cultures were degassed by autoclaving and equilibrated in an anaerobic chamber (10% H2, 5% CO2, 85% N2) for at least 24 h before inoculation. For purification of sulfite reductase, the MgSO4·7H2O and (NH4)2SO4 present in minimal A salts were replaced with (per liter) 0.08 g of MgCl2 and 0.8 g of NH4Cl, and cultures were supplemented with 0.5 mM each amino acid except methionine and cystine. For preparation of holoenzyme 0.25 g of djenkolic acid was provided as sole sulfur source (30); for preparation of flavoprotein, starter cultures grown with 0.8 mM MgSO4·7H2O were diluted 1:100 into sulfur-free medium (31). The difference in protocols reflects our evolving strategies to induce maximal expression of the sulfite-reductase operon.
Enzyme Purification-- Respiratory vesicles were customarily prepared as described previously (19). When we sought to avoid the inactivation of NADH dehydrogenase I, cells were lysed, and vesicles were prepared in 50 mM MES buffer, pH 6.0, containing 10% glycerol (26). These conditions were not necessary for activity, however, so reactions with respiratory vesicles and enzymes were conducted in 50 mM KPi, pH 7.8.
NADH dehydrogenase II was purified from both quinone-proficient and -deficient strains. The wild-type enzyme was purified from KM50, an ndh-overexpressing strain which is quinone-proficient but lacks cytochrome oxidases o and d. Preliminary preparations had revealed that trace amounts of cytochrome oxidase activity interfered slightly with assays of NADH oxidase activity during purification. Because cytochrome oxidase mutants grow poorly in air, KM50 was grown in 10 liters of anaerobic LB, 0.2% glucose to an A600 of 0.3 and then shifted into air and grown to a final A600 of 0.45. The anaerobic culture permitted a high enzyme titer to be achieved, particularly because of the loss of suppression (32) by Fnr protein. The final growth period in air ensured ubiquinone sufficiency. 6.7 g of cell pellet was suspended in 20 ml of 50 mM KPi, pH 7.8, and lysed by French press. Debris was removed by centrifugation at 15,000 rpm × 20 min, and vesicles were pelleted at 100,000 × g for 3 h. NADH dehydrogenase was extracted from resuspended membranes with deoxycholate and purified over hydroxyapatite as described (33). Active fractions were verified by both NADH:ferricyanide and NADH:ubiquinone-1 oxidoreductase assays. Purified fractions produced a single band on SDS gels (not shown). Samples were dialyzed against 90 volumes of 5 mM KPi, pH 7.5 and 0.1% cholate with two changes of buffer, in order to remove exogenous FAD.
Ubiquinone-deficient NADH dehydrogenase II was purified from an anaerobic culture of KM44, which lacks both menaquinone and ubiquinone. The deletion of fnr and presence of a multicopy ndh-expressing plasmid again ensured abundant synthesis of the enzyme even during anaerobiosis. Six liters of 0.2 A600 cultures were harvested. The lack of respiratory activity in the vesicles verified that quinones were absent. NADH dehydrogenase II was extracted and purified as above.
Sulfite reductase holoenzyme was purified from JI132/pJRS102. This plasmid (34) includes cysG, which encodes an enzyme needed to generate mature siroheme for sulfite reductase and nitrite reductase, and causes abundant overproduction of sulfite reductase in sulfur-limited cells. A 2.6-g cell pellet was recovered from 4.5 liters of 0.4 A600 culture. Cells were lysed by French press. The purification scheme of Ref. 24 was followed, including streptomycin sulfate and ammonium sulfate precipitations, Superose sizing column, and hydroxyapatite column. The enzyme activity was monitored by assay for sulfite reductase activity. Purified enzyme migrated as 56- and 61-kDa bands. The spectra of pure fractions matched that observed by other investigators (22).
Sulfite reductase flavoprotein (subunit B) was purified from
DH5/pcysJ. Six liters of culture were harvested at 0.25 A600, and the purification scheme described
above was followed. Activity was followed by NADPH:cytochrome
c reductase activity. Active fractions were bright yellow
throughout the purification, and the spectrum of the pure enzyme
reproduced that reported earlier for the flavoprotein (30).
The FAD and FMN content of purified proteins was measured by extraction and fluorescence detection (22), with authentic FMN and FAD as standards. Specific removal of FMN from the sulfite reductase flavoprotein subunit was achieved by 18 h incubation with 1 mM chloromercuriphenylsulfonic acid at 4 °C (31); the FMN content of treated enzyme was below the detection limit, and ~90% of FAD was retained.
Unless noted otherwise, sulfite reductase reactions were conducted in 50 mM Tris, pH 7.5, containing 50 µM EDTA.
Assay of O2 Formation--
The production of
O
2 by the aerobic respiratory chain in the absence of active
NADH dehydrogenase I was measured by the standard cytochrome
c method (1). This method did not work when NADH dehydrogenase I was active, because that enzyme can directly reduce cytochrome c. Acetylation of cytochrome c (35)
diminished the rate of its direct reduction. Five ml of 0.62 mM cytochrome c was treated with 125 mM acetic anhydride for 30 min on ice, dialyzed against two
changes of 1 liter of 50 mM KPi, pH 7.0, and
frozen at
70 °C. 1.5 µM acetylated cytochrome
c then replaced 10 µM native cytochrome
c in the standard assay of O
2. O
2 produced by xanthine oxidase and NADH dehydrogenase II was quantitively detected
by the acetylated cytochrome c.
Acetylated cytochrome c was still reduced too rapidly by
sulfite reductase to serve as an assay of O2 production by
that enzyme. Epinephrine, tetranitromethane, luciferase, and nitro blue
tetrazolium assays (reviewed in Ref. 35) also failed because of direct
interactions between sulfite reductase and the detector molecules.
Therefore, O
2 was detected by its ability to reduce Fe3+. Reactions contained 5.2 mM ADP, 210 µM FeCl3, 800 units of catalase, enzyme, and
enzyme substrate in 3.5 mM Tris, pH 7.8. All stock solutions were prepared in 3.5 mM buffer, with the
exception of enzymes, which were in 50 mM Tris, pH 7.8. The
ADP stock solution was adjusted to pH 7.8. Control reactions included
30 units of SOD. Reactions were initiated by the addition of enzyme
substrate and aliquoted to cuvettes that were each continuously
monitored for absorbance at 562. At defined 1-min intervals, ferene was added to consecutive cuvettes to a 900 µM final
concentration, and the jump in absorbance was recorded. This increase
was used to calculate the amount of Fe2+ that had
accumulated by that time, using
mM = 27.5 at 562 nm for
the Fe2+-ferene complex. In control experiments this system
efficiently detected the O
2 that was generated by xanthine
oxidase, fumarate reductase, and NADH dehydrogenase II. Catalase was
included to prevent re-oxidation of Fe2+ by
H2O2. The rate of NADPH oxidation by sulfite
reductase was also measured in the same reaction mixture (without
ferene addition) so that O
2 yields could be compared with
enzyme turnover. The oxidation rates of some enzymes were affected by
the presence of ferric-ADP. Some of this effect was due to direct
reduction of the ferric chelate, and this flux was subtracted in
calculations of the oxidase activity of the enzymes.
Assay of H2O2 Formation-- H2O2 production was measured using horseradish peroxidase. H2O2 was generated in high yield by quinoneless respiratory chains and could be conveniently assayed using o-dianisidine as the dye (36). Because NADH is not consumed by quinoneless membranes and interferes with horseradish peroxidase-based assays by competing with the dye, respiration-proficient vesicles were added to scavenge the residual NADH before detection by the horseradish peroxidase system. The H2O2 yielded by the oxidizing vesicles was scant compared with that from the quinoneless membranes, which were present in higher concentration. The initial reaction contained in a total volume of 4.2 ml: 30 µM NADH, quinoneless vesicles containing 0.2 units of NADH dehydrogenase, and 30 units of SOD. At time points over 8 min, 800-ml aliquots were removed, and 0.02 units of NADH oxidase activity were added as UM1 membranes to scavenge rapidly the residual NADH. Then to the sample was added a 100 µM mixture consisting of 150 µM o-dianisidine, 0.06 mg/ml horseradish peroxidase, 25 µM EDTA, and 50 mM KPi, pH 7.8. Absorbance was determined within 5 min at 460 nm. This protocol provided a time course of H2O2 production by quinoneless vesicles; since the scant H2O2 that was generated by UM1 vesicles was common to all time points, it was subtracted. We noted that when vesicles were formed by French pressing, some catalase was typically trapped inside the vesicles. Because this catalase interfered with H2O2 detection, scavenging vesicles were prepared from the catalase mutant UM1, and the measurements of H2O2 formation by respiratory chains were typically conducted on vesicles derived from catalase-free mutants.
The yield of H2O2 from actively respiring vesicles was too small to be quantitated accurately by the dianisidine assay. Therefore the fluorescent dyes scopoletin, diacetyldichlorodihydrofluorescein, and amplex red were used as horseradish peroxidase substrates. Diacetyldichlorodihydrofluorescein was synthesized by standard methods (37). Vesicles were incubated with different amounts of NADH up to 300 µM, in each case until the NADH was exhausted, and the H2O2 that was generated was quantitated by published methods (38-40). By so doing we determined the H2O2 yield as a function of the amount of NADH that was oxidized. With all three fluorescent substrates, we observed a high yield of H2O2 when as little as 10 µM NADH was added; above 25 µM NADH, however, the ratio of H2O2 produced per NADH oxidized was consistent, and that is the value cited in this work. Whereas it is formally possible that H2O2 formation is unusually rapid at low NADH concentrations, we suspect an horseradish peroxidase-based artifact, since the H2O2 yield at these doses was dependent upon the period of dye development. The basis of the artifact remains unclear.
Enzyme Assays-- NAD(P)H oxidase activities of vesicles and of purified enzymes were determined spectrophotometrically at 340 nm. NADH dehydrogenase activity was quantitated by monitoring NADH oxidation in the presence of 100 µM plumbagin and 3 mM cyanide (21). Transhydrogenase, ferricyanide reductase, hydroxylamine reductase, and sulfite reductase activities were measured by standard methods (23). Cytochrome c reduction was assayed with 0.1 mM cytochrome c rather than 1 mM.
Unless otherwise indicated, all assays were conducted in air-saturated
buffers at room temperature. For studies of enzyme oxidation at
different oxygen concentrations, reaction mixtures were assembled in an
anaerobic chamber. Anaerobic buffer was added to cuvettes, and the
cuvettes were then sealed with rubber septa. The desired amount of
aerobic buffer was then added by injection through the septa from an
air-tight syringe. The cuvettes were filled to the top throughout this
process to avoid any head space and prevent oxygen exchange with a gas
phase. The reactions in the sealed cuvettes were then monitored at 340 nm. The temperature dependence of reactions was determined by using a
thermostatted spectrophotometer and equilibrating the buffer
temperature before initiating the reaction.
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RESULTS |
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NADH dehydrogenase II Is the Primary Site of O2 and
H2O2 Formation in the Aerobic Respiratory
Chain--
The first goal of this study was to identify the
autoxidizable members of the aerobic electron transport chain. In a
previous investigation fumarate reductase was identified to be an
autoxidizable component of the anaerobic chain, but that enzyme is not
normally synthesized in air and was shown not to be responsible for the O
2 produced by most aerobically grown cells (21).
Membrane vesicles were prepared from cells that had been grown both
aerobically and anaerobically on minimal glucose medium. These vesicles
contained intact respiratory chains but were prepared at neutral pH,
which inactivates NADH dehydrogenase I, one of two respiratory NADH
dehydrogenases in E. coli. That was useful, because NADH
dehydrogenase I has a cytochrome c reductase activity that
interferes with the standard O2 assay (see below). In these vesicles NADH dehydrogenase II was intact, and the chain oxidized NADH
efficiently. As was observed previously, substantial O
2 was
formed (0.9 nmol min
1 O
2/unit of NADH
dehydrogenase; Table II, line 1). The
precise yield per electron flux varied slightly among preparations but was consistently in the range of 3-5 O
2 per 10,000 electrons, also in agreement with a previous report (19). O
2 was not
formed from vesicles that lacked both of the NADH dehydrogenases,
confirming that the O
2 evolved from the respiratory chain
(Table II, line 1 versus 7).
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Experiments verified that fumarate reductase was not a significant
source of O2 in aerobic, NADH-reduced chains. First, fumarate, a potent inhibitor of Frd autoxidation (21), failed to abate the
NADH-driven O
2 production (data not shown). Second, the
absence of neither menaquinone, which delivers electrons to Frd, nor of Frd itself slowed O
2 formation by NADH-reduced cells (Table
II, lines 1-3). In contrast, both of these mutations blocked the
formation of O
2 during the respiration of
-glycerol
phosphate, a substrate that efficiently directs electrons to Frd (lines
4-6). The vesicles used in the experiments of Table II were derived
from cells grown on aerobic casamino acids medium, which is known to
permit substantial synthesis of Frd; vesicles from glucose medium
showed no dependence at all of O
2 production upon Frd (not
shown). Thus the O
2 that was generated during the oxidation of
NADH evolved from some site on the respiratory chain other than
fumarate reductase.
Superoxide is a formal intermediate during the reduction of oxygen at
cytochrome oxidase. However, free O2 is not formed in
significant quantity, since oxygen remains tightly bound during its
four-electron reduction to water (42). To test whether the substoichiometric O
2 that we detected above escaped from
cytochrome oxidase, cyanide was provided to block the binding of oxygen
to that enzyme. NADH oxidation was 98% inhibited, but O
2
production actually increased (Table II, line 1). The increase
presumably occurred because electrons were backed up onto an
autoxidizable component of the chain upstream of the cytochrome oxidase.
Rapid O2 formation was also observed in vesicles from mutants
that lacked both ubiquinone and menaquinone (Fig.
1A). In these vesicles the
electrons from NADH could proceed no further than NADH
dehydrogenase II itself. The high rate of O
2 production exceeded that which was measured with quinone-proficient vesicles, presumably because NADH dehydrogenase II remained highly reduced, as if
cyanide were present (compare Table II, lines 1 and 8). Using a
ubiA high Km mutant that generates some
ubiquinone if its substrate, 4-hydroxybenzoate, is provided exogenously
(43), we demonstrated that O
2 production and respiratory
capacity were inversely related (Fig.
2).
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E. coli contains two abundant NADH dehydrogenases: NADH
dehydrogenase I, a large proton-translocating complex analogous to the
mammalian enzyme, and NADH dehydrogenase II, a non-proton-translocating single subunit enzyme important in maintaining a redox-balanced dinucleotide pool. Whereas the previous experiments pinpointed NADH
dehydrogenase II as a primary O2 source, they did not test the
autoxidizability of NADH dehydrogenase I, which had been deliberately inactivated by the neutral pH used during vesicle preparation. NADH
dehydrogenase I interferes with standard O
2 detection because it directly reduces exogenous cytochrome c. In order to
determine whether NADH dehydrogenase I generates O
2, we
chemically acetylated cytochrome c and provided it in lower
concentrations in the O
2 assays (see "Materials and
Methods"). Vesicles were then prepared from cells that lacked either
NADH dehydrogenase I (nuo) or NADH dehydrogenase II
(ndh). The NADH dehydrogenase I-deficient membranes generated O
2 during respiration as abundantly as did wild-type membranes, but the NADH dehydrogenase II-deficient membranes generated very little (Table III). The
ndh and nuo mutations were also transduced into
the quinoneless mutants, and once again substantial amounts of
O
2 were made only when NADH dehydrogenase II was present (not shown). Thus NADH dehydrogenase II, but not NADH dehydrogenase I,
appeared to be a significant source of O
2.
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H2O2 is the other toxic product of enzyme
autoxidation, and its production was measured in respiring membranes
that contained either NADH dehydrogenase I or II. An artifact that
affects H2O2 detection restricted us to observe
H2O2 production between 25 and 200 µM NADH (see "Materials and Methods"), but this is
the range of internal NADH concentration inside exponentially growing, glucose-fed cells (19). The H2O2 yield was much
higher when electrons passed through NADH dehydrogenase II than through
NADH dehydrogenase I (Table III). Since the remainder of the
respiratory chain is shared between these two dehydrogenases, the
implication was that NADH dehydrogenase II is the preponderant
respiratory source of H2O2 as well as
O2. In fact, substantial H2O2 was
formed when quinoneless vesicles containing only NADH dehydrogenase II were incubated with NADH (Fig. 1B).
Side-by-side measurements suggested that NADH dehydrogenase II formed
about 4 O2 per 10 molecules of H2O2
both during respiration and in the quinoneless membranes. Since 1 molecule of H2O2 was generated by dismutation
per each 2 O
2, it appeared that NADH dehydrogenase can
transfer electrons to oxygen either singly or in pairs. Xanthine
oxidase shows a similar behavior: approximately 15% of the
autoxidations occur by two consecutive electron transfers to oxygen,
forming two molecules of O
2, whereas the other 85% involve
the direct divalent reduction of oxygen to H2O2
(15).
Purified NADH Dehydrogenase Generates O2 by Chemical
Oxidation of Its Flavin--
The autoxidation of NADH dehydrogenase II
was more closely examined after purification. The enzyme contains an
FAD cofactor, which receives electrons from NADH, and a tightly bound
ubiquinone cofactor that co-purifies with the enzyme and apparently
mediates electron transfer from the FAD to the diffusible quinone pool (33). No metal centers exist in the enzyme.
NADH dehydrogenase II was overexpressed from a plasmid in both
wild-type and quinone-deficient backgrounds. Membrane vesicles derived
from overexpressing cells generated proportionately more O2
than did wild-type cells (Table II, lines 9 and 10). The enzyme was
purified to homogeneity from both the wild-type and quinone-deficient membranes using an established protocol (33). The turnover numbers for
O
2 and H2O2 production were similar
for both forms of the enzyme (Table IV).
Thus it is the flavin moiety that directly transfers electrons to
molecular oxygen.
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Electron transfer to oxygen was much slower than electron transfer to
good univalent oxidants such as ferricyanide or plumbagin, indicating
that the rate-limiting step in O2 formation is the transfer of
electrons from the reduced enzyme to dissolved molecular oxygen. This
transfer was not mediated by adventitious metals, since metal chelators
such as EDTA and DETAPAC had a slight (~40% maximum) accelerating
effect on autoxidation. Similar stimulations have been described for
xanthine oxidase and fumarate reductase, which also autoxidize at
reduced flavins (21), although the basis of the effect is not known.
The reactivity of NADH dehydrogenase II with oxygen is presumably an
accidental consequence of the exposure of its reduced flavin to
dissolved oxygen. Accordingly, the reaction followed chemical kinetics
and slowed proportionately at low concentrations of oxygen (Fig.
3A). The second-order rate
constant for enzyme oxidation was 1.9 × 104
M1 s
1 at 37 °C (with
saturating NADH). An apparent energy of activation of 26 kJ was
calculated from data above 15 °C. At lower temperatures autoxidation
was slower than expected from Arrhenius behavior, possibly
because of changes in the enzyme structure (Fig.
3B).
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Sulfite Reductase Is an Autoxidizable Soluble Protein-- The conclusion that isolated respiratory proteins autoxidize primarily or exclusively from flavin moieties raised the question of whether there is any abundant protein in E. coli that spuriously reacts with oxygen at a non-flavin site. This question cannot be answered comprehensively without examination of every redox protein. However, most of the known non-respiratory redox enzymes contain only flavins as redox cofactors. Among the exceptions, sulfite reductase may be the most abundant. It contains FAD, FMN, [4Fe-4S] clusters, and siroheme, and electrons flow in that order from the NAD(P)H donor to sulfite. Sulfite reductase was known to be capable of transferring electrons to a broad range of chemical oxidants, including oxygen (23). We therefore sought to determine which of its moieties were involved in autoxidation.
When aerobic cells were grown on the poor sulfur source djenkolate, which induced sulfite reductase synthesis about 100-fold above that of cystine-supplemented cells, no reliable difference in the total NADPH oxidase activity of the soluble extract could be determined. However, the overproduction of sulfite reductase from a plasmid caused a substantial increase in the NADPH oxidase activity of the whole-cell extract (not shown). Superoxide formation was difficult to measure by established assays, since sulfite reductase directly reduces both naive and acetylated cytochrome c (see below).
Holoenzyme was purified by a standard protocol, from SOD-deficient
cells in which sulfite reductase had been overproduced. The enzyme
spectrum matched that of previous reports. Turnover numbers were 1450 min1 (2160 min
1), 4420 min
1
(6800 min
1), and 18,700 min
1 (31100 min
1) for sulfite, hydoxylamine, and cytochrome
c reduction, respectively. The parenthetical rates were
obtained with added FMN and ranged from 93 to 118% of published values
(23). The purified enzyme was somewhat FMN-deficient, containing 0.51 FMN/FAD.
Oxidase activity was indeed present; the turnover number was 63 NADPH
min1 per holoenzyme, similar to the value 75 min
1 measured by Siegel et al. (23). The
turnover number was increased somewhat by iron chelators, as seen for
other enzymes. Because FMN slowly dissociates from sulfite reductase
(44), it was important to verify that the turnover to oxygen was not
mediated by the enzymic reduction of free flavins. The sulfite
reductase transfers electrons from its FAD moiety to free flavins (31),
and the subsequent oxidation of those flavins by molecular oxygen could provide a spurious oxidase activity. To test whether the apparent oxidase activity arose from such a reaction, the NADPH oxidase activity
was measured as a function of FMN concentration when free FMN was
provided. In fact, the provision of FMN in high concentrations stimulated electron transfer both to oxygen and to other acceptors. However, the stimulation of the oxidase activity was biphasic. Extrapolation of the second phase to low FMN concentrations indicated that the initial phase approximately doubled the activity (Fig. 4). This was presumably achieved by
filling the empty FMN sites. The residual effect of higher FMN was less
pronounced, exceeded the capacity of the enzyme to stably bind
additional FMN at unoccupied sites, and was not saturated at 1 µM. We concluded that the excess FMN stimulated the
oxidase activity by acting as an artificial electron acceptor rather
than by occupying authentic FMN binding sites in the slightly
de-flavinated enzyme. However, substantial basal oxidase activity
existed even at very dilute enzymes concentrations (2 nM).
In that circumstance the possible amount of free FMN could not have
accounted for the oxidase activity, if one assumes a linear
relationship between free FMN and spurious oxidase turnover.
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Therefore the oxidase activity of the isolated, unsupplemented enzyme reflected bona fide electron transfer directly from the holoenzyme to oxygen, without a free-FMN intermediate. In subsequent oxidase experiments free flavin was not provided.
Electron Transfer Occurs from a Flavin-- As reported in earlier studies (23), cyanide blocked electron transfer to sulfite but did not inhibit the oxidase activity. That result suggested that the oxidase reaction did not occur at the siroheme site, although electron transfer from an alternative face of the heme was not formally ruled out. The latter possibility was refuted by the observation that oxidase activity persisted in enzyme that was synthesized by a cysG mutant, which generates a siroheme-deficient enzyme (data not shown).
Both the siroheme and the iron-sulfur cluster are contained in the subunit, whereas FAD and FMN are bound by the
subunit of the
holoenzyme. The
subunit (flavoprotein) can be purified in a stable
form in the absence of the
subunit, and this was achieved using an
established protocol with cells that overexpressed only
cysH, the flavoprotein structural gene. The purified
flavoprotein exhibited a turnover number per FAD cofactor in excellent
agreement with that of the holoenzyme (Table
V). Siegel and Davis (45) had observed
that flavoprotein that was dissociated from the iron protein by
denaturants retained 10-18% of the oxidase activity; their low yield
was matched by poor recoveries of other reductase activities and
presumably reflected enzyme damage during subunit dissociation. To
identify more precisely the autoxidizing moiety, we removed FMN by
treatment with p-chloromercuriphenylsulfonate. The treated
enzyme retained FAD but was effectively devoid of FMN (Table V).
Although the FMN-free flavoprotein retained transhydrogenase activity,
the oxidase activity was reduced by 75%. This activity loss is
quantitatively similar to the residual ferricyanide and menadione
reductase activities that others (30) have reported. These experiments
indicated either that FMN is the preponderant site of autoxidation or
that it electronically interacts with FAD to facilitate the reaction of
FAD with oxygen. FAD alone has slight reactivity with oxygen.
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Sulfite Reductase Generates H2O2, Not
O2, as Its Major Oxidation Product--
Greater than 97% of
the electrons that were transferred to oxygen by sulfite reductase were
recovered as H2O2 (data not shown); thus the
tetravalent reduction of oxygen to water did not occur. This yield
represented both the H2O2 and the O
2
that were generated as initial products, since any O
2
spontaneously dismutes to form H2O2. To
determine the initial yield of O
2, an iron-reduction assay was
developed (see "Materials and Methods"). We found that only about
10% of the oxidation of the holoenzyme and 20% of the oxidation of
the flavoprotein produced O
2 (Table V). In contrast, about
60% of the lesser oxidation of the FMN-depleted flavoprotein generated
O
2. It is not clear whether FAD oxidation occurs in the
FMN-containing flavoprotein; if so, virtually all of the O
2 from that protein could arise from autoxidation of the FAD. Thus FMN
appeared to be the predominant site of sulfite reductase oxidation, with H2O2 as the predominant direct product,
whereas a small minority of autoxidation occurred at the FAD site, with
approximately equal production of O
2 and
H2O2.
The kinetics of sulfite reductase oxidation reflected its binding
interactions with NADPH and chemical interactions with oxygen. The
oxidase activity exhibited an apparent Km for NADPH of 5 µM in air-saturated buffer, and high concentrations
of either NADPH or NADP+ did not suppress oxidation (data
not shown). The rate of autoxidation was proportionate to oxygen
concentration (Fig. 5A). The
rate constant for reaction with dissolved oxygen was 2.4 × 103 M1 s
1 per FMN
at 37 °C. The temperature dependence indicated an activation energy
of 49.9 kJ/mol (Fig. 5B).
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NADH-reduced Enzyme Reacts with Oxygen but Poorly with an NADP+ Analogue-- Eschenbrenner et al. (31) reported the interesting observation that NADH can reduce the FMN moiety of sulfite reductase but that the reduced enzyme then reacts poorly with cytochrome c. This is in stark contrast to NADPH-reduced enzyme. We repeated the observation, finding that NADH bleached the FMN absorbance peak. Interestingly, the NADH-reduced enzyme reacted well with oxygen, showing a turnover number only slightly lower than when NADPH was the reductant. The turnover number to cytochrome c was as low as that to oxygen, suggesting that the slow step might be the reduction of the enzyme. Surprisingly, acetylpyridine dinucleotide phosphate (AcPyNADP+), the NADP+ analogue that is reduced by reduced FAD, could only be reduced by the NADH-treated enzyme at a very slow rate, whereas acetylpyridine dinucleotide (AcPyNAD+), an NAD+ homologue, was as reducible as cytochrome c or oxygen (Table VI). This appears to confirm that the NADH- and NADPH-reduced enzymes are electronically dissimilar and that the electrons perhaps localize exclusively on the FMN in the NADH-reduced enzyme. Although the reason for this remains uncertain (perhaps a residual interaction with NAD+ affects the electron localization), it supports the conclusion that the preponderant site of electron transfer to oxygen is the FMN moiety rather than the FAD.
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DISCUSSION |
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Why Are Flavins the Predominant Sources of O2 and
H2O2?--
Oxygen is a triplet species, and
electron transfers to it from singlet donors must occur in
univalent steps. Consequently, O
2 is the immediate
product. The univalent reduction potential of oxygen is low enough that
it cannot pull electrons from unwilling donors; the donors must be
proficient at univalent redox reactions. For this reason reduced
dinucleotides and thiols are relatively stable in air. One anticipates
that oxygen will react only with those electron carriers whose
univalent oxidation states are stable: those carriers containing
transition metals or with sufficient conjugation to stabilize the
univalent oxidation products through resonance structures. In
biological systems the obvious candidates are iron-sulfur clusters,
hemes, quinones, and flavins. In this study we have identified flavins
as the primary sites of chemical oxidation of the respiratory chain and
of sulfite reductase. In both cases the hemes, iron-sulfur clusters,
and quinone pools were not substantial sources of either O
2 or
H2O2.
Why don't these other moieties react with oxygen? A number of answers are possible, but a likely one is that, unlike flavins, these moieties are sequestered in environments that are either sterically inaccessible to oxygen or are hydrophobic. The importance of burying clusters is underscored by those exceptional clusters whose catalytic function requires that they be solvent-exposed. These clusters (on nitrogenase, Fnr protein, and dehydratases) react rapidly with oxidants. In contrast, the clusters of sulfite reductase and NADH dehydrogenase I are buried in the protein, where they conduct internal electron transfers, and are stable in air.
The clusters of succinate dehydrogenase and fumarate reductase are
presumably near the enzyme surface, since their biochemical function is
to reduce directly diffusible quinones, but electron transfer to oxygen
may be blocked by the hydrophobicity of the membrane interior. That
environment would not tolerate the generation of a hard anion like
O2. The same issue may prevent the autoxidation of reduced
respiratory quinones. A contrasting example is the bc1 complex of other
bacteria and eukaryotes, which generates O
2 when its Qo site
semiquinone reacts with oxygen (46). It would be interesting to see
whether this site has a local tolerance for charge that has the side
effect of permitting O
2 formation.
Differential Rates of Flavin Autoxidation--
Free reduced
flavins react rapidly with oxygen (and may themselves be sources of
intracellular O2 (47)). The rates at which flavoproteins react
with oxygen are generally lower but range over orders of magnitude.
Massey (51) has noted that members of the dehydrogenase class typically
react slowly, in contrast with electron transferases, oxidases, and
monooxygenases. We have seen substantial variation within nominal
members of the dehydrogenase class, from undetectably low (NADH
dehydrogenase I), to moderate (succinate dehydrogenase, sulfite
reductase), to high (NADH dehydrogenase II, fumarate reductase). The
different energies of activation for the oxidation of sulfite reductase
(50 kJ/mol) and NADH dehydrogenase II (26 kJ/mol) suggest that
electronic effects may control these rates; certainly local polypeptide
context influences the relative stability of dihydroflavins and
flavosemiquinones. However, several other factors are likely to be
important, including the degree of flavin exposure and the local
electron density. The failure of succinate dehydrogenase to autoxidize
at the same efficiency as its homologue, fumarate reductase, may
reflect the tendency of these enzymes to distribute their electrons
differently; on reduced fumarate reductase electron density may be high
on the flavin, in preparation for fumarate reduction; on succinate
dehydrogenase the electron density is probably highest on its higher
potential iron-sulfur clusters, in anticipation of ubiquinone
reduction. It may be that NADH dehydrogenase I does not autoxidize
because its electrons are primarily sequestered away from the flavin, on its iron-sulfur clusters.
The rates at which flavoenzymes react with oxygen in vivo
will also depend upon the overall enzyme redox status, which in turn
depends upon the amount of oxidative substrate present. Thus the
elimination by mutation of oxidized ubiquinone as a competitor for
their electrons accelerated O2 formation both by NADH
dehydrogenase II and fumarate reductase. We presume that the presence
of sulfite would slow the rate at which sulfite reductase generates
H2O2, although interference by sulfide
prevented us from doing the H2O2 assays that
would have tested this idea directly.
O2 Versus H2O2 as the Direct
Oxidation Product--
The overall turnover number for flavin
oxidation reflects the rate-limiting step, which is the transfer of the
first electron from the reduced flavin to molecular oxygen. In
contrast, the valency of the electron transfer, that is whether
superoxide or hydrogen peroxide is the product that leaves the enzyme,
is determined by the relative rates at which the nascent O
2
either dissociates from the enzyme or, alternatively, reacts (after a
spin flip) with the flavosemiquinone radical to form a peroxy complex
(48). Whether flavoenzyme autoxidation generates O
2 or
H2O2 is critical to cell physiology, because
these two oxidants have qualitatively and quantitatively different
impacts upon fitness. O
2 damages dehydratases with high
reactivity and specificity, and 10
9 M is
toxic (49). H2O2 is less inhibitory to growth,
and levels up to 10
5 M can be tolerated. On
the other hand, H2O2 has a potentially more
lasting effect, since it can attack DNA; 10
4
M causes measurable mutagenesis and lethality (50). Overall it seems that cells would be more tolerant of
H2O2 than equimolar O
2 production, and
most committed oxidases generate H2O2 as an exclusive product.
NADH dehydrogenase II and sulfite reductase both followed the behavior
of typical flavin dehydrogenases (51), forming only a small amount of
O2. Steric factors may affect these yields as follows: if the
O
2 is formed within an enzyme cleft, the frequency of
collision between it and the flavin semiquinone rises before the
O
2 can escape, and that may lessen the yield of diffusible O
2. However, at least in the case of sulfite oxidase, the
valency of FMN oxidation is additionally determined by its interaction with the adjacent FAD. By preparing recombinant protein fragments that
contained only the FMN-binding site, Fontecave and colleagues (24) were
able to study the behavior of this flavin in isolation. The flavin
redox potentials of the protein fragment agreed with those determined
by Ostrowski et al. (30) for the entire flavoprotein as
follows:
0.322 and
0.382 V for the FADH2/FADH·
and FADH·/FAD couples, and
0.327 and
0.152 for the
FMNH2/FMNH· and FMNH·/FMN couples. These
potentials are low enough to favor oxygen reduction
(E'0 (O
2/O2) =
0.18 V).
The striking feature is that the univalent oxidation of reduced
FMNH2 will generate a semiquinone product that, with
respect to oxygen, could be fairly stable. In fact this was observed as
follows: while the reduced recombinant FMN fragment oxidized rapidly
upon exposure to air, the product was a stable neutral flavosemiquinone
whose own oxidation required hours. Since the enzyme lost only one
electron, O
2 must have been the exclusive product. That
observation contrasts with our finding that
H2O2 is almost the sole oxidation product of
the complete flavoprotein; the difference is probably the influence of
the adjacent FADH2. As the FMNH2 reacts with
oxygen, the FADH2 may be predisposed by its low potential
to push an electron toward nascent FMNH and, from there, onto the
nascent O
2, thus forming H2O2 rather
than O
2 as the free product. In fact, Siegel et al. (23) originally speculated that FMN cycles between reduced and semiquinone states in bridging electron flow between FAD and the [4Fe-4S] cluster; in our analysis, oxygen has displaced the cluster as the electron acceptor.
Strikingly analogous behavior was reported for xanthine oxidase; the
two-electron-reduced enzyme generated only O2 upon
autoxidation. However, if the metal sites next to the autoxidizing
flavin were also reduced, H2O2 was generated as
a near-stoichiometric product (16).
Variability in Oxidative Stress among Environments and
Organisms--
A priori it seemed possible to us that the
O2 production in E. coli might arise in relatively
similar proportions from many disparate redox enzymes. However, we have
found that turnover numbers (and titers) of autoxidizable enzymes vary
widely, and it appears that as a result most O
2 may arise from
only a few flavoenzymes. Earlier studies indicated that the great
majority of O
2 and H2O2 are formed by
the respiratory chain, which we found here is due to NADH dehydrogenase
II. In fact, extrapolation of the O
2 yield from this enzyme
provided an estimate of intracellular O
2 formation,
10
5 M s
1, that agreed well with
subsequent calculations of the O
2 load that E. coli
can bear (49). However, the technical difficulties of measuring the
yields of these species inside growing cells has not yet permitted us
to confirm this result directly in vivo.
If one or a few enzymes form most of the cellular O2 and
H2O2, then the degree of oxidative stress may
change with growth circumstance as those enzymes are induced or
repressed. Both NADH dehydrogenase II and sulfite reductase, for
example, are highly regulated enzymes, and their 10- to 100-fold
differences in expression in different environments would presumably
cause proportionate changes in the amounts of O
2 and
H2O2 that they form. One can also foresee that
organisms that feature a high-titer, autoxidizable enzyme in their
primary metabolic pathway are likely to suffer more O
2 and
H2O2 formation than will others. We are
investigating whether this is the circumstance that forces some
microaerophiles to avoid air-saturated environments. The first-order
effect of oxygen concentration on enzyme oxidation makes plain the
benefit of microaerobic habitats to such organisms.
Finally, because O2 formation accelerates at high
temperatures, oxidative stress is among those chemical problems to
which aerobic thermophiles must have adapted. Since flavin
autoxidizability depends upon polypeptide context, it will be
interesting to determine whether homologous enzymes from thermophiles
have evolved to be less reactive with oxygen than those of mesophiles.
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ACKNOWLEDGEMENTS |
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We thank Pierfrancesco De Propris for efforts in developing the superoxide detection assay used in this work and Sharon Currie for strain constructions. We also thank Jacques Coves, Marc Fontecave, Nick Kredich, and Bob Gennis for generously providing bacterial strains and plasmids.
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FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grant GM49640.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.
Supported in part by National Institutes of Health Grant T32
GM07283-20.
§ To whom correspondence should be addressed. Tel.: 217-333-5812; Fax: 217-244-6697; E-mail: jimlay{at}uiuc.edu.
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ABBREVIATIONS |
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The abbreviations used are: SOD, superoxide dismutase; Frd, fumarate reductase; AcPyNADP+, acetylpyridine dinucleotide phosphate; AcPyNAD+, acetylpyridine dinucleotide; MES, 4-morpholineethanesulfonic acid.
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REFERENCES |
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