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Originally published In Press as doi:10.1074/jbc.M306251200 on August 11, 2003

J. Biol. Chem., Vol. 278, Issue 43, 42495-42504, October 24, 2003
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Effects of Hydrogen Peroxide upon Nicotinamide Nucleotide Metabolism in Escherichia coli

CHANGES IN ENZYME LEVELS AND NICOTINAMIDE NUCLEOTIDE POOLS AND STUDIES OF THE OXIDATION OF NAD(P)H BY Fe(III)*

Julia L. Brumaghim{ddagger}, Ying Li§, Ernst Henle, and Stuart Linn||

From the Division of Biochemistry and Molecular Biology, University of California, Berkeley, California 94720-3202

Received for publication, June 13, 2003 , and in revised form, August 7, 2003.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
DNA is damaged in vivo by the Fenton reaction mediated by Fe2+ and cellular reductants such as NADH, which reduce Fe3+ to Fe2+ and allow the recycling of iron. To study the response of Escherichia coli to such cycling, the activities of several enzymes involved in nicotinamide nucleotide metabolism were measured following an H2O2 challenge. NADPH-dependent peroxidase, NADH/NADP+ transhydrogenase, and glucose-6-phosphate dehydrogenase were most strongly induced, increasing 2.5-3-fold. In addition, the cellular ratios of NADPH to NADH increased 6- or 92-fold 15 min after exposure to 0.5 or 5 mM H2O2, respectively. In vitro, NADH was oxidized by Fe3+ up to 16-fold faster than NADPH, despite their identical reduction potentials. To understand this rate difference, the interactions of Fe3+ and Ga3+ with NAD(P)H were examined by 1H, 13C, and 31P NMR spectroscopy. Association with NADH occurred primarily with adenine at N7 and the amino group, but for NADPH, strong metal interactions also occurred at the 2'-phosphate group. Interaction of M3+ (Fe3+ or Ga3+) with the adenine ring would bring it into close proximity to the redox-active nicotinamide ring in the folded form of NAD(P)H, but interaction of M3+ with the 2'-phosphate group would avoid this close contact. In addition, as determined by absorbance spectroscopy, the energy of the charge-transfer species was significantly higher for the Fe3+·NADPH complex than for the Fe3+·NADH complex. We therefore suggest that upon exposure to H2O2 the NADH pool is depleted, and NADPH, which is less reactive with Fe3+, functions as the major nicotinamide nucleotide reductant.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The utilization of oxygen as the terminal electron acceptor for aerobic respiration results in exposure to toxic reactive oxygen species that arise as incompletely reduced byproducts of respiratory electron transport (1) or as side products of enzymes such as xanthine oxidase, NADH/NADPH oxidase, monooxygenases, and cyclooxygenases (2, 3). In addition, environmental agents such as ionizing or near-UV radiation (4) and chemicals such as paraquat, plumbagin, and menadione can generate and H2O2 in the cell (5). Reactive oxygen species cause damage to DNA, proteins, and lipids and are implicated in a variety of human pathologies including Alzheimer's disease, cancer, arteriosclerosis, and aging (6-9).

Exposure of Escherichia coli or mammalian cells to hydrogen peroxide results in two modes of killing: Mode I cytotoxicity peaks at <3 mM H2O2, whereas Mode II cytotoxicity occurs between 3 and 25 mM and is independent of the dose. These complex kinetics are also observed for DNA damage in vitro. This toxicity is attributable to DNA damage by reactive oxygen species generated via the Fenton reaction (10).

The hydroxyl radical is an extremely powerful oxidant that reacts with most organic substrates at nearly diffusion-limited rates (11). Studies of oxidative DNA damage by the Fenton reaction have indicated that the species is not free hydroxyl radical, but is likely stabilized by coordination to the DNA-bound iron (12, 13).

To continue DNA damage, a reductant must be present in vivo to regenerate Fe2+ from Fe3+. Genetic studies suggested that NADH might act as this reductant, and biochemical studies showed that NADH could drive DNA damage by iron and peroxide in vivo (10, 14). The level of NAD(H) drops severely in glucose-starved E. coli cells, and these cells are remarkably resistant to H2O2 (14). Alternatively, raising the NADH level in vivo by eliminating or negatively regulating NADH dehydrogenase activity either genetically or chemically by inhibiting electron transport with KCN, dramatically sensitizes E. coli to killing by H2O2. KCN does not further enhance the sensitivity of an NADH dehydrogenase mutant to H2O2 (14). Acceleration of DNA damage in isolated nuclei also increases dramatically in the presence of NAD(P)H and Fe3+ bound to EDTA or diethylenetriaminepentaacetic acid (15).

NADPH is an essential cofactor for the catalytic activities of glutathione peroxidase (16, 17), catalase (18), and NADPH-dependent alkylhydroperoxidase (19). Moreover, the majority of NADPH in mammalian cells is bound to catalase, which it reactivates after the enzyme is inactivated by H2O2 (18, 20). It might be expected that, besides NAD(P)H, glutathione could facilitate redox cycling for the Fenton reaction. However, glutathione synthase and glutathione reductase mutants exhibit normal sensitivity to Mode I killing (21, 22). As a result of the induction of the soxRS regulon under oxidative stress, the expression of glucose-6-phosphate dehydrogenase (G6PD)1 increases, resulting in the conversion of NADP+ to NADPH and making these cells resistant to oxidants (23, 24). Although it was initially believed that reduced glutathione was responsible for the antioxidant effects of the soxRS response, it was shown that the redox state of NADPH, not glutathione, modulates oxidative sensitivity (25). This antioxidant role of G6PD was substantiated further by observations that disruption of the gene encoding G6PD in mouse embryonic stem cells resulted in greatly enhanced sensitivity to oxidative stresses (26) and that Saccharomyces cerevisiae G6PD null mutants also are sensitive and unable to adapt to hydrogen peroxide (27, 28). The introduction of an intracellular NADPH-generating system restored substantial oxidative stress resistance to the G6PD-deficient yeast (25). In sum, NADPH is clearly important for protection against oxidative stress (26).

To understand better the role of NADPH in resistance to oxidative stress, the effects of H2O2 on the activities of several enzymes involved in the synthesis and utilization of NAD(P)H and changes in the nicotinamide nucleotide pools have been monitored following exposure of E. coli to H2O2. In addition, the relative abilities of NADH and NADPH to reduce Fe3+ to Fe2+, as well as the nature of Fe3+ interactions with NAD(P)H, were studied.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Bacterial Strains, Buffers, and Reagents—E. coli strains AB1157 (F-thr-1 leuB proA2 his-4 thi-1 argE2 lacY1 galK2 rpsL supE) and UM1 (LacY rpsL thi-1 katE1 katG14) were grown with vigorous shaking in K medium (1% glucose, 1% casamino acids, 1 mM MgSO4, 0.1 mM CaCl2, and M9 salts) unless otherwise noted. Water was double distilled before use. H2O2 (30%) was from Fisher Scientific, NADH (disodium salt), NADPH (tetrasodium salt), and NMNH (disodium salt) were from Sigma. All other chemicals were from Sigma unless otherwise specified. Solutions of Fe3+ and Ga3+ salts were freshly prepared prior to use and stored at pH 2.5. Elemental analyses were performed by Desert Analytics (Tucson, AZ).

Spectroscopic Measurements—1H, 13C{1H}, and 31P{1H} NMR spectra were acquired on a Bruker DRX-500 spectrometer at 500, 125, and 202 MHz, respectively. Chemical shifts for 1H, 13C{1H}, and 31P{1H} NMR spectra are reported in ppm ({delta}) relative to SiMe4 and 40% H3PO4, respectively. Absorbance spectra for NAD(P)H were acquired on a Hewlett-Packard 8453 spectrophotometer with a diode array detector or a Cary 300 dual beam spectrophotometer. Kinetics measurements of NAD(P)H oxidation utilized the Cary spectrophotometer.

Chromatography—Nicotinamide nucleotides were resolved on a PerkinElmer 250 HPLC apparatus equipped with a guard column and a Jordi reverse phase C18-DVB column (250-mm length x 4.6-mm inner diameter). The mobile phase was 98% 25 mM tributylammonium bicarbonate, pH 10, 2% acetonitrile (eluent A) and 80% acetonitrile, 20% water (eluent B). A gradient program was initially set to 100% eluent A, increasing over 20 min to 65% eluent A and 35% eluent B, then finally increasing over 40 min to 50% each of eluents A and B. Absorbances of the nucleotides were monitored at 327 nm with a diode array detector.

Enzyme Assays—E. coli cells in K medium were challenged with 50 µM H2O2 at a density of 4 x 107 cells/ml. After 15 min at 37 °C, the bacteria were chilled, washed, and harvested by centrifugation. Cell pellets were resuspended in 5 ml of 50 mM KCl, 5 mM dithiothreitol, 100 mM Tris-HCl, pH 8.0, and the suspensions were sonicated four times for 15 s and then cleared by centrifugation. Protein concentration was determined by the method of Bradford (29).

NAD+ kinase activity was assayed by a procedure similar to that of Zercz et al. (30), but [32P]NAD+ was the substrate, and NADP+ was monitored by radioactivity after isolation with thin layer chromatography on silica plates with ethanol and 1 M ammonium acetate (1:1) as solvent. G6PD and isocitrate dehydrogenase activities were assayed according to the protocols described in the Worthington Manual (31). The assay of NADH transhydrogenase activity was based on the method of Zercz et al. (30). NADH dehydrogenase and NAD(P)H oxidases were assayed as described by Cavari et al. (32). NAD(P)H-dependent peroxidases were partially purified by ammonium sulfate fractionation and DEAE-cellulose chromatography and assayed following the method of Coves et al. (33). The reaction was carried out under aerobic conditions and in the presence of an NADH-generating system. The activity was determined from the appearance of absorbance of NADPH at 340 nm. NAD+ pyrophosphatase activity was determined as described previously (34). The assay of DNA ligase activity was based on the method of Olivera et al. (35), and NAD+ glycohydrolase assays were based on the method of Barbieri et al. (36).

Measurement of E. coli Nicotinamide Nucleotide Pools—Intracellular NAD(P)H and NAD(P)+ concentrations were determined by a modification of the method of Klaidman et al. (37). E. coli AB1157 was grown in 50 ml of K medium plus 1 µg/ml thiamine at 37 °C to an A600 of ~0.5, and then H2O2 was added to the culture medium, and incubation was continued for 15 min. Anaerobic cultures were grown with bubbling 95% N2 and 5% CO2 prior to and after the addition of H2O2. After H2O2 challenge, cell pellets were collected by centrifugation and resuspended in 1 M KCN, 0.3 N KOH, 5 mM EDTA and then mixed with an equal volume of 50 mM tributylammonium bicarbonate, pH 10, to give a final volume of ~2.5 ml. The lysate was cleared by centrifugation, filtered through a 0.2-µm filter, and analyzed by HPLC. Amounts of nicotinamide nucleotides were determined by measuring areas of the HPLC peak traces at 327 nm and comparing these with standard 3-nmol samples of each of the nicotinamide nucleotides run under identical conditions. Sample traces are given in the supplementary data (Fig. S1). Cellular concentrations were calculated assuming that an A600 = 1.0 corresponds to 109 cells/ml and that the volume of a cell is 10-12 ml (38).

Metal Titrations of NADH and Its Analogs Monitored by NMR Spectroscopy—NMR spectra of freshly prepared 0.5-ml solutions of NAD(P)H, NMNH, or ADP-ribose between 10 and 100 mM in D2O, pD ~ 7.5, were acquired after incremental addition of FeCl3·6 H2O or Ga(NO3)3 in D2O as indicated. Addition of DCl at pH 2.5 to a D2O solution of NADPH in the same amounts as those added for Fe3+ or Ga3+ resulted in no significant shifting of the NMR resonances. Spectra were acquired within 30 min of the addition of Fe3+. Percent broadening of NMR resonances upon addition of Fe3+ was calculated using Equation 1.

(Eq. 1)
A broadening of 100% indicates that no resonance was visible. Kd values and fractional occupancy of Ga3+ were determined as described previously (39). Percentages of NAD(P)H in linear and folded forms were calculated as described by Oppenheimer et al. (40).

Elemental Analyses of Ga3+/NAD(P)H Precipitates—20 mg of NAD(P)H was dissolved in 0.4 ml of H2O, and 1 eq of Ga(NO3)3 in 7.5 µl was added. A white precipitate formed immediately for the NADH sample, but an additional 1 eq of Ga(NO3)3 was added to the NADPH sample to obtain sufficient precipitate for analysis. Both samples were centrifuged at 14,000 rpm for 5 min, and the supernatant was discarded. The precipitates were washed three times by centrifugation with 1 ml of water and dried under vacuum.

For the NADH sample, the best fit formula was

For the NADH sample, the best fit formula was

Metal Titrations of NAD(P)H Monitored by Absorbance Spectroscopy—Absorbance spectra of freshly prepared solutions of NAD(P)H were acquired after incremental addition of Fe(NO3)3 to 50 or 500 µM solutions of NAD(P)H, or Ga(NO3)3 to 66 µM solutions of NADPH. All spectra were acquired within 30 min of the addition of M3+.

Kinetic Measurements of NAD(P)H Oxidation—Freshly prepared solutions of 16 µM NAD(P)H were brought to 100 mM ethanol, 1.25 µM H2O2, and/or 80 µM FeCl3 as indicated. A340 was measured at 25 °C every 0.2 s for 7 min and corrected with appropriate blanks. Best fit lines were calculated from the initial rates. For anaerobic measurements, the water used to prepare the samples was degassed with argon for 3 h prior to use, and the samples were prepared and sealed under an argon atmosphere. The initial rates were very dependent on concentration, so care was taken to use the same solutions and order of addition of reactants for both the NADH and NADPH trials.

Molecular Modeling—Molecular models for Fe3+ binding to NADPH were constructed using Spartan 02 (Wavefunction, Inc., Irvine, CA). The NADPH conformation was obtained from the most probable folded conformation as predicted by NMR spectroscopy (41): 4'-5'-gg and 5'-O-gg rotamers for both the adenyl and reduced nicotinamide ribose units, 2'-endo conformation for the nicotinamide ribose, and 3'-endo conformation for the adenine ribose. To ensure that the molecule remained in the folded conformation, the distance between the adenine and reduced nicotinamide rings was restrained to 3.6 Å. This NADPH conformation was energy minimized using MMFF94 force field calculations (42) before adding the Fe3+ ion at a restrained distance of 2.1 Å (43) to the adenine N7. Because of steric hindrance resulting from the close contacts between the adenine N7-bound Fe3+ and the adenine amino group, a bond between the Fe3+ and the amino group was included and restrained to a distance of 2.3 Å.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Nicotinamide Nucleotide Enzyme Levels after H2O2 Challenge of E. coli—Because of a possible involvement of NAD(P)H in mediating H2O2 toxicity, the activities of several enzymes involved in nicotinamide nucleotide metabolism (Table I) were measured before and after challenge with H2O2 (Table II). In consideration of the existence of the "early" H2O2 induction of proteins, the activities of which are induced during the first 10-18 min but return to normal within 30 min after the addition of hydrogen peroxide (44), the activities were measured 15 min after challenge of catalase-deficient cells with 50 µM H2O2. Catalase is the major enzyme for removing hydrogen peroxide, but catalase-deficient mutants are not more sensitive to killing by H2O2 than are wild type cells (22). Thus, catalase mutants were used in this study to elucidate other defense systems against hydrogen peroxide. E. coli G6PD, which specifically reduces NADP+, is one of the major sources of cellular NADPH. Upon H2O2 challenge, the activity of G6PD was induced by about 2.9-fold (Table II). Meanwhile, the activity of isocitrate dehydrogenase, which also reduces NADP+, was not induced.


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  		TABLE I
Enzymatic activities assayed

 

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  		TABLE II
Changes in enzymatic activities after H2O2 challenge

Cell cultures were treated with 50 µM H2O2 and then harvested after 15 min for assay. All measured values are the average of at least three trials. An enzymatic activity of 100% indicates no change from unchallenged cells. Numbers in parentheses are the activities before challenge.

 

NADH/NADP+ transhydrogenases reversibly transfer a hydrogen atom between NADH and NADP+ (Table I). Upon membrane energization either by respiration or by ATP hydrolysis, the rate of reduction of NADP+ by NADH is increased severalfold, and the equilibrium constant for the reaction increased from 0.79 for the non-energy-dependent reaction to 480 (45). Presumably the physiological function of these transhydrogenases is to provide NADPH for biosynthesis and detoxification (46). To investigate whether either of these energy-linked transhydrogenase activities is increased upon hydrogen peroxide exposure, they were assayed in catalase-deficient E. coli cells after exposure to 50 µM H2O2 for 15 min (Table II). The assays included an NADH-regenerating system. The respiration-driven transhydrogenase activity was increased almost 3-fold, but the ATP-dependent transhydrogenase activity showed no increase. Likewise, no induction of NADH dehydrogenase activity was observed.

In addition to alkylhydroperoxidase (19) and o-dianisidine peroxidase (47), E. coli contains an NADH-dependent peroxidase (33). Although the former enzymes do not utilize H2O2 as substrate, the latter one does. The NADH-dependent peroxidase activity was elevated by only 38% after exposure to peroxide. However, NADPH-dependent peroxidase activity increased by 248% upon H2O2 challenge. NADH and NADPH oxidase activities were induced to only 140% of normal activity.

The turnover of NAD+ is very rapid in the cell, and it is 4-fold faster under aerobic conditions than anaerobic conditions (48). Moreover, in mammalian cells, DNA breaks caused by oxidative stresses can deplete NAD+ via the NAD+ glycohydrolase activity of poly(ADP-ribose) polymerase, whereas in E. coli, DNA ligase hydrolyzes NAD+. Thus, it was proposed that the turnover of NAD+ might have important functions under conditions of oxidative stress (48). Three enzymatic activities initiate NAD+ turnover in E. coli: NAD+ glycohydrolase, NAD+ pyrophosphatase, and DNA ligase. However, none of the activities of these enzymes changed after hydrogen peroxide exposure (Table II). Finally, NAD+ kinase activity was unchanged. In conclusion, the changes in the enzyme activities indicate that H2O2 is depleted by NADPH-dependent peroxidases in the absence of catalase and predict that cellular NADPH levels might be increased relative to those of NADH.

Measurement of Nicotinamide Nucleotide Pools in E. coli—The determination of nicotinamide nucleotide pools is complex because of the sensitivity of NAD(P)H to degradation or oxidation during isolation. Lundquist and Olivera (49) estimated the concentrations of NAD(P)+ in E. coli but did not include the reduced nicotinamide nucleotides. Bochner and Ames (50) used an acid extraction procedure for purification of the nicotinamide nucleotides which probably resulted in oxidation of NAD(P)H prior to quantitation. Therefore, we modified the method used for brain cells by Klaidman et al. (37) to measure the effect of oxidative stress upon the free pools in E. coli (Table III). Cells were lysed by NaOH in the presence of cyanide and EDTA. EDTA serves to chelate metal ions that could oxidize NAD(P)H, whereas cyanide forms adducts with NAD(P)+ to stabilize these species against degradation in basic solution, and these adducts can conveniently be quantitated by absorbance at 327 nm.


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  		TABLE III
Nicotinamide nucleotide pools 15 min after H2O2 challenge

All measurements are the average of three independent experiments. In all cases, the standard deviations were less than 10% of the mean values shown.

 

Prior to exposure to H2O2, NADPH accounted for 2 and 7% of the free nicotinamide nucleotides in aerobically grown or anaerobically grown E. coli, respectively. 15 min after exposure to 0.5 mM H2O2, however, the NADPH level increased by about half, whereas the NADPH:NADH ratio increased by 5.7-fold. After exposure to 5.0 mM H2O2, the effects were even more dramatic: although the steady-state level of NADPH dropped to half, and the ratio of NADP(H) to NAD(H) remained unchanged, the ratio of NADPH to NADH increased by 92-fold. After exposure to 10 mM H2O2, the concentration of NADH dropped below the detectable level. Clearly, the reduced pools dramatically shift to favor NADPH under oxidative stress, which correlates well with the induced activity of NADPH-generating enzymes and NADPH-dependent peroxidases after oxidative stress.

Kinetics of NAD(P)H Oxidation by Fe3+—The shift in reduced nicotinamide nucleotide pools from NADH to NADPH during oxidative stress could indicate that NADH can reduce Fe3+ to Fe2+ more rapidly than NADPH and therefore drive the Fenton reaction more efficiently than NADPH. To test this hypothesis, the initial rates of NADH and NADPH oxidation by Fe3+ were determined (Table IV). Because dioxygen can gain an electron to form the superoxide radical, which is itself capable of promoting oxidative damage (51), the reactions were measured under both aerobic and anaerobic conditions.


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  		TABLE IV
Rates of NAD(P)H oxidation by Fe3+

Measurements were performed at 25 °C with 160 µM NAD(P)H, as described under "Experimental Procedures."

 

Under anaerobic conditions, the rate of NADH oxidation by Fe3+ was 5.5-fold faster than that of NADPH, whereas under aerobic conditions, the initial rates were slightly faster, and NADH remained the preferred substrate by roughly 6-fold (Table IV). When H2O2 was added to the reaction, the oxidation rates increased dramatically, and the initial oxidation rate of NADH was 16-fold that of NADPH. These rates decreased only slightly in the presence of 100 mM ethanol, which can quench free hydroxyl radical (10), indicating that free hydroxyl radicals were not a major reactant in the oxidation. The presence of organic material that could effectively compete for freely diffusable hydroxyl radical might resemble the cellular environment more closely. Thus, altering the reductant pool to favor NADPH over NADH under oxidative stress would result in a major decrease in the potential to reduce Fe3+ and hence in levels of Fe2+-generated oxidants.

Analysis of Complexes of NAD(P)H with Fe3+ and Ga3+ by NMR Spectroscopy—NADH and NADPH differ only by the presence of a 2'-phosphate group on the adenine ribose of NADPH. Their structural properties in solution are also similar, with both molecules existing in an equilibrium between an extended form, in which the reduced nicotinamide ring and the adenine ring at opposite ends of the molecule, and a folded conformation in which the two hydrophobic rings are stacked (41). At 22 °C, 36% of either species is in the folded form (40). In addition, the reduction potentials of the two molecules are nearly identical (0.32 V (52)). Thus the different rates of oxidation by Fe3+ must be caused by different localization of Fe3+ on the nucleotides, by an electronic difference by which the 2'-phosphate raises the energy of an electron transfer to Fe3+, and/or by a shift in equilibrium between the linear and folded conformations.

To study such differences, NMR titrations of NADH and NADPH with Fe3+ were carried out. Titrations were also done with Ga3+, which is similar to Fe3+ in ionic radius (0.645 and 0.620 Å, respectively (53)), and has a similar hydrolysis equilibrium constant for the deprotonation of an aqua ligand (pKh = 2.6 and 2.2, respectively (54)). (The pKh reflects the tendency of metal ions to coordinate to hard donor atoms such as oxygen versus softer donor atoms such as nitrogen.) In addition, both Ga3+ and Fe3+ are labile metals with ligand exchange occurring at the rate of 403 and 160 s-1, respectively (55). One important difference between these metal ions, however, is that Ga3+ has no unpaired electrons and is therefore diamagnetic, as opposed to the paramagnetic high spin Fe3+, which has five unpaired electrons. This property of Ga3+ makes it ideally suited to NMR studies at high metal ion concentrations (56-59), whereas Fe3+ can be utilized effectively only at very low concentrations because of paramagnetic broadening of the resonances. One advantage of the Fe3+ paramagnetic broadening of NMR resonances, however, is the ability to determine Fe3+ localization in the molecule because signal broadening is proximity-dependent. Finally, Ga3+ cannot be reduced to Ga2+, so binding could be studied independently of metal ion reduction.

In studies of ATP, dAMP, and calf thymus DNA, the adenine ring was observed to contain two favored metal ion binding sites, the adenine N1 and N7 atoms (60-62). In the case of ATP, N1 is the predominant localization site only at pH 3-4.5, whereas binding of Fe3+ to the adenine N7 is favored at neutral pH (60). When we examined broadening of the proton resonances by Fe3+ of 10 mM ADP-ribose (a molecule similar to NADH but lacking a nicotinamide ring) with 0.01, 0.05, and 0.10 eq of Fe3+, broadening was slightly greater for the adenine H8 proton than for the H2 proton. This differential broadening indicates that Fe3+ interaction with the adenine N7 site was slightly stronger than with N1 (Fig. 1). Analysis of additional ribose resonances was not possible because of the highly overlapping signals from the {alpha} and {beta} anomers of this compound. Examining the proton broadening of 10 mM NMNH with 0.01 and 0.05 eq of Fe3+ or 50 mM NMNH with 0.05 eq of Fe3+, only general proton resonance broadening was observed with little selectivity (Fig. 2, A-C).



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  		FIG. 1.
Broadening of ADP-ribose adenosine proton resonances by Fe3+. FeCl3·6 H2O in the equivalents shown was added to a 10 mM ADP-ribose solution, and spectra were acquired as described under "Experimental Procedures." The numbering scheme is given in Fig. 5.

 


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  		FIG. 2.
Broadening of NAD(P)H and NMNH proton and carbon resonances by Fe3+. FeCl3·6 H2O was added to the nucleotide solutions, and spectra were acquired as described under "Experimental Procedures." A, 1H NMR resonances observed with 10 mM nucleotide, 0.1 mM Fe3+. B, 1H NMR resonances observed with 10 mM nucleotide, 0.5 mM Fe3+. C, 1H NMR resonances observed with 100 mM NADH or NADPH and 5 mM Fe3+, or with 50 mM NMNH and 2.5 mM Fe3+. D, 13C NMR resonances observed with 100 mM NADH or NADPH and 1 mM Fe3+, or with 50 mM NMNH and 0.5 mM Fe3+. The numbering scheme for NAD(P)H is given in Fig. 5.

 



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  		FIG. 5.
Conclusions for localization of Fe3+ upon NAD(P)H. Sites of major broadening of 1H, 31P, and 13C NMR resonances by Fe3+ are indicated in purple for NADH (A) and NADPH (B). C, differences in Fe3+ localization upon NADPH compared with NADH. Red indicates the sites of greater Fe3+ localization, orange indicates sites of slightly less localization, and green indicates sites of much less localization (weak sites of interaction).

 

When a comparison was made for Fe3+-induced proton resonance broadening for NADH and NADPH (Fig. 2) at 10 mM nucleotide and 0.01 eq of Fe3+ (Figs. 2A and 3), strongly selective broadening occurred for the adenine H8 proton of NADH, but only slight broadening was seen for the adenine H8 proton of NADPH. Meanwhile, sharpening was observed for the adenine H2 proton of NADPH, but not the adenine H2 proton of NADH. This signal sharpening is most likely a result of conformational change in the molecule upon the addition of the metal ion. Less broadening of the reduced nicotinamide ring and nicotinamide ribose protons was also observed for NADPH than NADH. In essence, these results indicate that the adenine N7 is the major site for Fe3+ binding to NADH (as was the case for ATP), but for NADPH there was no binding to the adenine N1 and less general binding to the reduced nicotinamide ring and the nicotinamide ribose. The smaller broadening seen for the H8 resonance of NADPH might indicate that at lower Fe3+ concentrations, Fe3+ is localizing at this site to a lesser extent.



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  		FIG. 3.
Broadening of the adenine 1H NMR H8 and H2 resonances for NADH and NADPH by Fe3+. FeCl3·6 H2O was added to 10 mM NAD(P)H in D2O as indicated, and spectra were acquired as described under "Experimental Procedures." With 0.01 eq of Fe3+, broadening of the adenine H8 resonances was 54% for NADH and 11% for NADPH.

 

These conclusions were confirmed with 10 mM nicotinamide nucleotide and 0.05 eq of Fe3+ (Figs. 2B and 3). With NADH there was general broadening, but with NADPH there was less binding of the adenine N1 (as observed by the smaller amount of broadening at the adenine H2 proton) and less broadening of the reduced nicotinamide and nicotinamide ribose protons. Finally, at 100 mM nucleotide and 0.01 eq of Fe3+ (50 mM NMNH), the most notable effect was reduced broadening of the nicotinamide protons for NADPH versus NADH (Fig. 2C). It should be noted that at 0.05 eq of Fe3+, general broadening of all NMR resonances was observed, both because of the higher Fe3+ concentration and because of the formation of radical species as Fe3+ oxidized the nucleotides. Such broadening was most noticeable for NADH (Fig. 2) because of the faster rate of NADH oxidation. For ADP-ribose, which cannot be oxidized, the addition of up to 0.10 eq of Fe3+ showed only selective resonance broadening (Fig. 1) compared with the general broadening seen for NADH (Fig. 2).

13C NMR resonance broadening (Fig. 2D) suggests that Fe3+ localizes near the nicotinamide amide of NADH (as observed by broadening of the nicotinamide C7 resonance). This selectivity was somewhat less for NADPH, and absent for NMNH. It should be noted that this Fe3+ interaction at the nicotinamide C7 is seen primarily at the high concentrations (100 mM) necessary to obtain 13C NMR spectra. Likewise, broadening of the carbon resonances on the adenine ring is stronger for NADH than NADPH (Fig. 2D), indicating greater interaction of Fe3+ with the adenine ring of NADH than that of NADPH. Addition of 0.01 eq of Fe3+ also resulted in extreme broadening of the phosphorus resonances (>80%) of the pyrophosphate backbone of NADH and NADPH and the 2'-phosphate of NADPH (data not shown), indicating that Fe3+ also localizes at the negatively charged phosphate groups.

Addition of Ga3+ to solutions of NADH resulted in the immediate formation of a white precipitate, preventing NMR studies. However, addition of Ga3+ to NADPH solutions resulted in precipitate formation only at high concentrations. As noted under "Experimental Procedures," elemental analysis of the precipitates indicated that they contained roughly two Ga3+/NAD(P)H molecule. Evidently, Ga3+ coordinates to NAD(P)H, most likely at the negatively charged pyrophosphate backbone, creating a neutral, insoluble species. Despite the difficulty with precipitation, spectra of NADPH were obtained from 1H NMR titrations with up to 2 eq of Ga3+ (Fig. 4A). Significant shifting of the adenine H8 and H2 resonances as well as the adenine ribose H2' resonance was observed with increasing Ga3+ concentration, suggesting that Ga3+, like Fe3+, localizes around the adenine N7 and N1, although in this case with roughly equal affinity at the two sites. Above 1.5 eq of Ga3+ signal broadening occurred because of precipitation.



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  		FIG. 4.
NMR titrations of NADPH with Ga3+. A, shifting of the adenine H8 and H2 (left) and the adenine ribose H2' (right) resonances in a 1H NMR titration of 1 mM NADPH with Ga3+ in D2O. B, shifting of the 2'-phosphate (left) and pyrophosphate backbone (right) resonances in a 31P NMR titration of 10 mM NADPH with Ga3+ in D2O.

 

Interaction of Ga3+ with NADPH was also examined with 31P NMR (Fig. 4B). Strong shifting of the 2'-phosphate resonance was observed ({delta} = 3.9) up to 1.0 eq of Ga3+, whereas the pyrophosphate shifting was relatively weak ({delta} = 0.12). Consistent shifting of the 2'-phosphate resonance occurred with Ga3+ up to 0.8 eq, as seen also for shifting of the adenine ribose H2' resonance (Fig. 4A). Shifting of the latter resonance indicates that the Ga3+ has a high affinity for this site in addition to the adenine N7 and N1 sites. Also noteworthy in the pyrophosphate backbone resonance at high equivalence of Ga3+ is a small shift from a single resonance into two distinguishable peaks, indicating that the two phosphorus atoms become chemically inequivalent. Such splitting could indicate that one oxygen atom of only one of the backbone phosphates was coordinating a Ga3+ ion. In contrast, similar experiments with NMNH indicated that Ga3+ did not localize on the reduced nicotinamide ring but only at the terminal phosphate group. Complete tables of the 1H, 13C, and 31P NMR resonance broadening for Ga3+ titrations with NADPH and NMNH are included in the supplemental material (Tables S1 and S2).

Our interpretation of the NMR results is summarized schematically in Fig. 5. Both NADH and NADPH have major Fe3+ binding sites at the adenine N7. NADH also has weaker interactions with adenine N1 and the reduced nicotinamide ring. In contrast, NADPH shows strong binding at the 2'-phosphate group, presumably at the expense of binding to the adenine N1 and reduced nicotinamide ring and to a lesser extent, the adenine N7. In essence, the 2'-phosphate group of NADPH diminishes interactions with the less specific sites seen for NADH, as expected for the addition of a strong site for Fe3+ localization.

Estimation of Kd for NADPH and Ga3+—Because of the broadening of NAD(P)H NMR resonances by Fe3+ at substoichiometric amounts, Kd values could not be calculated for this interaction. However, a Kd for Ga3+ interactions with NADPH could be estimated from 1H NMR Ga3+ titration data of 1 mM NADPH (Fig. 6A). Fitting the shifts in the adenine ribose 2'-phosphate resonance for a one-site binding model, the Kd was calculated to be ~550 µM (Fig. 6B). Given this Kd, the percentage of 1 mM Ga3+ bound to 1 mM NADPH at the 2'-phosphate was calculated to be 48%. Because the 2'-phosphate is a relatively strong site of metal ion localization, this could be close to the total amount of NADPH-bound Ga3+.



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  		FIG. 6.
Estimation of Kd for Ga3+ binding to NADPH. A, spectra of the adenine H8, H2, and the adenine ribose H2' resonances of 1 mM NADPH in D2O with Ga3+ added as Ga(NO3)3. B, the Kd was determined from the shifting of the H2' resonance in the 1H NMR titration of NADPH with Ga3+.

 

Analysis of Complexes of NAD(P)H with Fe3+ and Ga3+ by Absorption Spectroscopy—The absorption spectra during a titration of 50 µM NADH or NADPH with Fe3+ are shown in Fig. 7. The absorption band at 260 nm is primarily caused by the adenine ring with a small contribution from nicotinamide ring, whereas the less intense band near 340 nm is solely the result of the nicotinamide ring (63). Addition of Fe3+ to either NADH (Fig. 7A) or NADPH (Fig. 7B) resulted in a decrease in the absorption at 340 nm and a shift of the maximum to higher wavelengths (lower energy). The hypochromism and bathochromic shift are likely caused by the formation of a charge-transfer species with Fe3+ which perturbs the energies of the reduced nicotinamide molecular orbitals. Similar hypochromic effects have been observed for the binding of metal complexes to DNA (64, 65). Titration of NADPH with Ga3+ under the same conditions (supplemental material, Fig. S2) caused no shift in the 340 nm band, and Ga3+ cannot form charge-transfer complexes. Only very slight hypochromicity (6%) was observed during the Ga3+ titration, suggesting that most of the change in the NAD(P)H absorption spectra brought about by Fe3+ results from the charge-transfer complex formation.



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  		FIG. 7.
Absorbance spectra of NADH and NADPH during titration with Fe3+ Nucleotide concentrations were 50 µM, and Fe3+ was added as Fe(NO3)3 to NADH (A) or NADPH (B). The inset shows the A340 values normalized to those before the addition of Fe3+.

 

The changes in A340 of NADH and NADPH as a function of Fe3+ concentration (Fig. 7B, inset), indicate that the interactions of Fe3+ with the two nucleotides differ considerably. Although the A340 of NADH decreased steadily with increased Fe3+, the A340 for NADPH reproducibly showed a rapid decrease then became constant. These differences suggest the presence of a second Fe3+ binding site peculiar to NADPH, presumably the 2'-phosphate group, or alternatively, that the charge-transfer complexes might differ for the two nucleotides. Indeed, absorption spectra at higher (500 µM) concentrations of nucleotides in the presence of Fe3+ exhibited absorbance maxima at 520 nm for NADPH and at 544 nm for NADH (supplemental material, Fig. S3), indicating a difference between the two charge-transfer complexes. These absorbances were maximal at 0.5 eq of Fe3+ and decreased over time.

Gutman and co-workers (66, 67) reported a charge-transfer band at 540 nm for both the (NADH)2Fe3+ and (NADPH)2Fe3+ complexes. Those experiments, however, were performed at pH 3.5 at which the pyrophosphate backbone oxygen atoms and two of the 2'-phosphate oxygen atoms of NADPH would have been protonated. Differential protonation of NAD(P)H in the respective experiments is the likely explanation for the changes in wavelength of the charge-transfer bands seen in our studies. In addition, although the charge-transfer absorbances observed here at pH 7.5 for NADPH (66) are approximately the same as reported by Gutman and Eisenbach (67) ({epsilon} = 800 versus 900 cm-1 M-1, respectively), the absorbances observed for NADH are half as intense ({epsilon} = 400 versus 875 cm-1 M-1, respectively). With the differences in pH and a partial occupancy of 20% for the Fe3+ binding to NAD(P)H, assuming that the Kd values of the Ga3+ and Fe3+ interactions are similar, the different intensity of these charge-transfer bands is not surprising.

The energy difference between the NADPH·Fe3+ and NADH·Fe3+ charge-transfer bands was calculated to be 2.4 kcal/mol, indicating that the charge-transfer species is lower in energy for NADH than for NADPH. Therefore, the rate for the reduction of Fe3+ to Fe2+ could be as much as 60-fold greater for NADH than for NADPH. This is consistent with the fact that oxidation rates of NADPH are up to 16-fold slower than those of NADH in the presence of Fe3+.

An Energy-minimized Structural Model for the Complex of Fe3+ with the Adenine Ring of NADH and NADPH—Based on the NMR and absorbance studies, a model for the interaction of Fe3+ with the adenine ring of NAD(P)H was constructed (Fig. 8). The most probable configuration of NAD(P)H as reported by Oppenheimer (40) was used as the starting structure for the folded conformation of the molecule, and then Fe3+ was placed at a distance of 2.1 Å from the adenine N7 (43). Because of steric hindrance between the bound Fe3+ and the adenine amino group which was encountered in the refinement of the model, a second bond between the Fe3+ and the amino group (2.3 Å) was added. The adenine amino protons are not observable by NMR spectroscopy, so direct M3+ interactions at this site cannot be observed. However, studies of M2+ interactions with adenine analogs showed significant interactions between the metal ion and the neighboring amino group (68). Attempts to add a Fe3+ ion bound to the adenine N1 resulted in severe steric hindrance with the adenine amino group, so it is not surprising that Fe3+ localization at this site was weaker than for adenine N7.



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  		FIG. 8.
Energy-minimized model of Fe3+ bound to the adenine ring of NADPH in the folded conformation. Fe3+ (yellow) is shown bound to the N7 and amine of the adenine ring. Note that the 2'-phosphate is directed away from the adenine binding site and the reduced nicotinamide ring.

 

To keep the model in a stacked conformation, the distance between the reduced nicotinamide ring and the adenine ring was restrained to 3.6 Å. Although the Fe3+ is bound to the adenine ring and not to the redox-active nicotinamide ring, the folded form of the molecule presumably brings the reduced nicotinamide ring close enough to the Fe3+ to promote electron transfer. From analysis of the NMR spectra of NADPH·Ga3+, the percentage of NADPH in the folded conformation does not change upon metal ion addition (as determined using the method described under "Experimental Procedures"), so the metal ion apparently does not hold the adenine and reduced nicotinamide rings in close proximity. The effect of Fe3+ on the folded/linear equilibrium of NAD(P)H could not be measured because of the relatively low equivalents of Fe3+ used in these titrations.

In solution at pH 7.5, the Fe3+ ion would have octahedral geometry, most likely binding a combination of water molecules and/or hydroxyl ions to complete the coordination sphere. In addition to interactions at the adenine N7 and the amine, it is also possible that one of the oxygen atoms from the pyrophosphate backbone could coordinate the Fe3+, which would explain the inequivalency seen for the backbone phosphorus atoms at high Ga3+ concentration in the NADPH 31P NMR titration (Fig. 3B).

As can be seen from Fig. 8, Fe3+ bound to the 2'-phosphate group of the NADPH would be directed away from the adenine N7 binding site and hinder charge-transfer from the reduced nicotinamide ring even in the stacked conformation. Of course, this model applies to the strong binding site of Fe3+ at the adenine N7, not to the other, weaker binding sites whose interactions are significantly diminished by the presence of the 2'-phosphate of NADPH.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Exposure of E. coli to H2O2 induced the activity of G6PD and the respiration-dependent NADH/NADP+ transhydrogenase, enzymes that synthesize NADPH. Additionally, activities of NADH-dependent oxidase and peroxidase as well as NADPH-dependent oxidase increased by about half, but NADPH-dependent peroxidase activity increased by 2.5-fold. These changes are consistent with the large measured increase in the ratio of the NADPH/NADH pools and with NADPH being a much slower reductant of Fe3+ than is NADH. This satisfying correlation between the induction of enzymes to produce NADPH, the increase in NADPH relative to NADH, and the much lower reactivity of NADPH with Fe3+ suggests that these changes are a cellular strategy to protect against oxidative damage and indirectly suggests that NADH is a primary cellular reductant responsible for reduction of Fe3+ to drive the Fenton cycle in vivo. NAD(P)H may not be solely responsible for the reduction of Fe3+ to Fe2+ and driving the production of reactive oxygen species, however, because reduced flavins have been implicated as reductants that drive the Fenton cycle, at least in nonrespiring E. coli cells (69). Nonetheless, we propose that in respiring cells the regulation of NADH and NADPH levels under oxidative stress conditions may enhance survival by storing reducing potential in the form of NADPH.

Consistent with our proposal, the induction of G6PD by H2O2 supports a growing understanding of the major role for this enzyme in defense against oxidative stress. G6PD deficiency in humans caused by point mutations in the G6PD gene results in the loss of intracellular NADPH and an enhanced sensitivity to oxidants (25). Persons carrying the mutations exhibit hemolytic anemia under conditions of oxidative stress by exogenous agents or infection (70). Severely impaired catalase activity resulting from the decrease in NADPH was proposed to be an underlying mechanism of oxidant susceptibility in these individuals (71). However, catalase-deficient humans show only minor adverse physiological effects such as an increased risk of diabetes (72).

In G6PD-null strains of mice or yeast, other NADPH-generating enzymes such as isocitrate dehydrogenase do not seem to compensate for the loss of G6PD in defenses against oxidative damage. Consistent with this observation, the activity of isocitrate dehydrogenase was not induced when E. coli was challenged with H2O2. Despite this lack of induction in E. coli, increasing the level of NADP+-dependent isocitrate dehydrogenase was found to increase resistance to oxidative damage of transformed mouse fibroblasts (73).

Zheng et al. (74) used DNA microarray analysis to profile transcriptional responses to hydrogen peroxide in E. coli. They listed the 30 most highly induced genes that showed levels of induction of more than 10-fold. 11 of these genes had unknown function, whereas none of the remaining 19 overlapped those whose protein activities were measured here. Of course, changes in transcript levels do not translate directly to changes in enzyme activity for a number of reasons, so that both approaches are necessary to obtain a comprehensive understanding of the responses to oxidative stress.

Because of the structural similarity of NADH and NADPH, the cause of the large difference in the rate of Fe3+ reduction between them must be the 2'-phosphate group on NADPH. The NMR data for both showed that M3+ ions localized primarily on the adenine N7 (and the nitrogen of the adenine amino group), with secondary interactions at the amide group of the reduced nicotinamide ring, the adenine N1, and the oxygen atoms in the pyrophosphate backbone. For NADPH, M3+ ions also localized on the 2'-phosphate group, as would be expected from the negative charge of this group. The presence of this 2'-phosphate on NADPH decreased metal binding at the adenine N7, N1, and the reduced nicotinamide ring compared with NADH. Because separate Kd measurements for M3+ association with the adenine N7 and the 2'-phosphate of NADPH could not be obtained, a direct comparison of strength of M3+ binding to these sites cannot be made. It is reasonable to assume, however, that metal ion binding at these two sites is competitive. The Fe3+ ion, when bound to the adenine N7 and the amino group, does not simultaneously bind the reduced nicotinamide ring, as evidenced by the NMR data. However, the equilibrium between the linear and folded forms of NAD(P)H would bring the reduced nicotinamide ring into close proximity of the adenine ring, enabling Fe3+ reduction.

It is also conceivable that the 2'-phosphate could change the electronic configuration of the nicotinamide nucleotide and increase the energy required to transfer an electron from NADPH to Fe3+ compared with NADH. Evidence for the higher energy of charge-transfer from NADPH to Fe3+ compared with NADH was deduced from differences in the absorption spectra during titrations of NADH versus NADPH with Fe3+. The presence of the charge-transfer band at a lower wavelength (higher energy) for NADPH indicates that the charge-transfer process is energetically less favorable for NADPH than for NADH. Presently, our results indicate that both structural and electronic differences between NADPH and NADH are responsible for the lower rate of Fe3+ reduction, and further studies are required to determine the individual contributions of these factors.

It is generally accepted that NAD+ serves primarily for the generation of ATP, whereas NADPH serves primarily as an electron or hydride donor in reductive biosynthetic reactions. We propose an additional purpose for the ubiquitous existence of the two similar nicotinamide nucleotides: by increasing the NADPH:NADH ratio under oxidative stress, an order of magnitude decrease in oxidative damage to cellular components via iron-mediated Fenton oxidants would result because of the slower reduction of Fe3+ by NADPH.


   FOOTNOTES
 
* This work was supported by in part by National Institutes of Health Grants RO1GM19020 and P30ES01986. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

The on-line version of this article (available at http://www.jbc.org) contains Tables S1 and S2 and Figs. S1, S2, and S3. Back

{ddagger} Supported by National Institutes of Health Training Grant T32ES07075. Present address: Dept. of Chemistry, Clemson University, 481 Hunter Laboratories, Clemson University, Clemson, SC 29634. Back

§ Present address: Pfizer, Inc., 4125 Sorrento Valley Blvd., San Diego, CA 92121. Back

Present address: Scimagix, Inc., 2855 Campus Dr., Suite 100, San Mateo, CA 94403. Back

|| To whom correspondence should be addressed: Division of Biochemistry and Molecular Biology, Barker Hall, University of California, Berkeley, CA 94720-3202. Tel.: 510-642-7583; Fax: 510-643-3388; E-mail: slinn{at}socrates.berkeley.edu.

1 The abbreviations used are: G6PD, glucose-6-phosphate dehydrogenase; HPLC, high performance liquid chromatography; M3+, Fe3+ or Ga3+; NMNH, nicotinamide mononucleotide, reduced form. Back


   ACKNOWLEDGMENTS
 
We are indebted to James A. Imlay, S. Michael Chin, Yong Zhang Luo, and Leighanne Olson for providing the foundation experiments for this work. We thank Dr. Kenneth Raymond for the use of his UV-visible spectrometers.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Chance, B., Sies, H., and Boveris, A. (1979) Physiol. Rev. 59, 527-605[Free Full Text]
  2. Fridovich, I. (1978) Science 201, 875-880[Abstract/Free Full Text]
  3. Cerutti, P. A. (1985) Science 227, 375-381[Abstract/Free Full Text]
  4. Tyrrell, R. M., and Pidoux, M. (1989) Photochem. Photobiol. 49, 407-412[Medline] [Order article via Infotrieve]
  5. Prieto-Alamo, M. J., Abril, N., and Puyeo, C. (1993) Carcinogenesis 14, 237-244[Abstract/Free Full Text]
  6. Barja, G. (2002) Ageing Res. Rev. 1, 397-411[CrossRef][Medline] [Order article via Infotrieve]
  7. Halliwell, B. (2001) Drugs Aging 18, 635-716
  8. Markesbery, W. R., and Carney, J. M. (1999) Brain Pathol. 9, 133-146[Medline] [Order article via Infotrieve]
  9. Olsinki, R., Gackowski, D., Foksinski, M., Rozalski, R., Rozalski, K., and Jaruga, P. (2002) Free Radic. Biol. Med. 33, 192-200[CrossRef][Medline] [Order article via Infotrieve]
  10. Imlay, J. A., Chin, S. M., and Linn, S. (1988) Science 240, 640-642[Abstract/Free Full Text]
  11. Halliwell, B., and Gutteridge, J. M. C. (1990) Methods Enzymol. 186, 1-85[CrossRef][Medline] [Order article via Infotrieve]
  12. Qian, S. Y., and Buettner, G. R. (1999) Free Radic. Biol. Med. 26, 1447-1456[CrossRef][Medline] [Order article via Infotrieve]
  13. Lloyd, R., Hanna, P. M., and Mason, R. P. (1997) Free Radic. Biol. Med. 22, 885-888[CrossRef][Medline] [Order article via Infotrieve]
  14. Imlay, J. A., and Linn, S. (1988) Science 240, 1302-1309[Abstract/Free Full Text]
  15. Peskin, A. V. (1996) Free Radic. Biol. Med. 20, 313-318[CrossRef][Medline] [Order article via Infotrieve]
  16. Sinha, B. K., Mimnaugh, E. G., Rajagopalan, S., and Meyers, C. E. (1989) Cancer Res. 49, 3844-3848[Abstract/Free Full Text]
  17. Mezzetti, A., Di Ilio, C., Calafione, A. M., Aceto, A., Marzio, L., Frederici, G., and Cuccurello, F. (1990) J. Mol. Cell Cardiol. 22, 935-938[CrossRef][Medline] [Order article via Infotrieve]
  18. Kirkman, H. N., and Gaetani, G. F. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 4343-4347[Abstract/Free Full Text]
  19. McKie, J. H., and Douglas, K. T. (1991) FEBS Lett. 279, 5-8[CrossRef][Medline] [Order article via Infotrieve]
  20. Hillar, A., Nicholls, P., Switala, J., and Loewen, P. C. (1994) Biochem. J. 300, 531-539[Medline] [Order article via Infotrieve]
  21. Greenberg, J. T., and Demple, B. (1986) J. Bacteriol. 168, 1026-1029[Abstract/Free Full Text]
  22. Imlay, J. A., and Linn, S. (1987) J. Bacteriol. 169, 2967-2976[Abstract/Free Full Text]
  23. Demple, B., and Amabile-Cuevas, C. F. (1991) Cell 67, 837-839[CrossRef][Medline] [Order article via Infotrieve]
  24. Nunoshiba, T., Hidalgo, E., Amabile-Cuevas, C. F., and Demple, B. (1992) J. Bacteriol. 174, 6054-6060[Abstract/Free Full Text]
  25. Scott, M. D., Zuo, L., Lubin, B. H., and Chiu, D. T.-Y. (1991) Blood 77, 2059-2064[Abstract/Free Full Text]
  26. Pandolfi, P. P., Sonati, F., Rivi, R., Mason, P., Grosveld, F., and Luzzatto, L. (1995) EMBO J. 14, 5209-5215[Medline] [Order article via Infotrieve]
  27. Nogae, I., and Johnston, M. (1990) Gene (Amst.) 96, 161-169[CrossRef][Medline] [Order article via Infotrieve]
  28. Slekar, K. H., Kosman, D. J., and Culotta, V. C. (1996) J. Biol. Chem. 271, 28831-28836[Abstract/Free Full Text]
  29. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve]
  30. Zercz, C. R., Moul, D. E., Gomez, E. G., Lopez, V. M., and Andreoli, A. J. (1987) J. Bacteriol. 169, 184-188[Abstract/Free Full Text]
  31. Worthington, C. C. (1988) Worthington Manual: Enzymes and Related Biochemicals, Worthington Biochemical Company, Freehold, NJ
  32. Cavari, B. Z., Avi-Dor, Y., and Grossowicz, N. (1968) J. Bacteriol. 96, 751-759[Abstract/Free Full Text]
  33. Coves, J., Eschenbrenner, M., and Fontecave, M. (1991) Biochem. Biophys. Res. Commun. 178, 54-59[CrossRef][Medline] [Order article via Infotrieve]
  34. Kornberg, A., and Pricer, W. E. (1950) J. Biol. Chem. 182, 763-778[Free Full Text]
  35. Olivera, B. M., and Lehman, I. R. (1967) Proc. Natl. Acad. Sci. U. S. A. 57, 1700-1704[Free Full Text]
  36. Barbieri, J. T., Moloney, B. K., and Mendo-Mueller, L. M. (1989) J. Bacteriol. 171, 4362-4369[Abstract/Free Full Text]
  37. Klaidman, L. K., Leung, A. C., and Adams, J. D., Jr. (1995) Anal. Biochem. 228, 312-317[CrossRef][Medline] [Order article via Infotrieve]
  38. Norland, S., Hedal, M., and Tumyr, O. (1987) Microb. Ecol. 13, 95-101
  39. Rai, P., Cole, T. D., Wemmer, D. E., and Linn, S. (2001) J. Mol. Biol. 312, 1089-1101[CrossRef][Medline] [Order article via Infotrieve]
  40. Oppenheimer, N. J., Arnold, L. J., Jr., and Kaplan, N. O. (1978) Biochemistry 17, 2613-2619[CrossRef][Medline] [Order article via Infotrieve]
  41. Oppenheimer, N. J. (1987) in Pyridine Nucleotide Coenzymes (Dolphin, D., Poulson, R., and Avramovic, O., eds) Vol. A, pp. 185-230, John Wiley and Sons, New York
  42. Kong, J., White, C. A., Krylov, A. I., Sherrill, D., Adamson, R. D., Furlani, T. R., Lee, M. S., Lee, A. M., Gwaltney, S. R., Adams, T. R., Ochsenfeld, C., Gilbert, A. T. B., Kedziora, G. S., Rassolov, V. A., Maurice, D. R., Nair, N., Shao, Y., Besley, N. A., Maslen, P. E., Dombroski, J. P., Daschel, H., Zhang, W., Korambath, P. P., Baker, J., Byrd, E. F. C., Van Voorhis, T., Oumi, M., Hirata, S., Hsu, C.-P., Ishikawa, N., Florian, J., Warshel, A., Johnson, B. G., Gill, P. M. W., Head-Gordon, M., and Pope, J. A. (2000) J. Comput. Chem. 21, 1532-1548[CrossRef]
  43. Obrey, S. J., Bott, S. G., and Barron, A. R. (2000) J. Chem. Cryst. 30, 61-63
  44. Storz, G., Tartaglia, L. A., Farr, S. B., and Ames, B. N. (1990) Trends Genet. 6, 363-368[CrossRef][Medline] [Order article via Infotrieve]
  45. Lee, C.-P., and Ernster, L. (1964) Biochim. Biophys. Acta 81, 187-190
  46. Bragg, P. D., Davies, P. L., and Hou, C. (1972) Biochem. Biophys. Res. Commun. 47, 1248-1255[CrossRef][Medline] [Order article via Infotrieve]
  47. Claiborne, A., and Fridovich, I. (1979) J. Biol. Chem. 254, 4245-4254[Abstract/Free Full Text]
  48. Park, U. E., Olivera, B. M., Hughes, K. T., Roth, J. R., and Hillyard, D. R. (1989) J. Bacteriol. 171, 2173-2180[Abstract/Free Full Text]
  49. Lundquist, R., and Olivera, B. M. (1971) J. Biol. Chem. 246, 1107-1116[Abstract/Free Full Text]
  50. Bochner, B. R., and Ames, B. N. (1982) J. Biol. Chem. 257, 9759-9769[Abstract/Free Full Text]
  51. Gutteridge, J. M. C., and Halliwell, B. (2000) Ann. N. Y. Acad. Sci. 899, 136-147[Medline] [Order article via Infotrieve]
  52. Stryer, L. (1988) Biochemistry, 3rd Ed., p. 400, W. H. Freeman and Co., New York
  53. Shannon, R. D. (1976) Acta Crystallogr. Sect. A 32, 751-767[CrossRef]
  54. Baes, C. F., Jr. (1976) The Hydrolysis of Cations, p. 27, John Wiley and Sons, New York
  55. Hugi-Cleary, D., Helm, L., and Merbach, A. E. (1987) J. Am. Chem. Soc. 109, 4444-4450
  56. Kersting, B., Telford, J. R., Meyer, M., and Raymond, K. N. (1996) J. Am. Chem. Soc. 118, 5712-5721
  57. Stephan, H., Freund, S., Meyer, J. M., Winkelmann, G., and Jung, G. (1993) Liebigs Ann. Chem. 43-48
  58. Wasielewski, E., Atkinson, R. A., Abdallah, M. A., and Kieffer, B. (2002) Biochemistry 41, 12488-12497[CrossRef][Medline] [Order article via Infotrieve]
  59. Atkinson, R. A., Salah El Din, A. L. M., Kieffer, B., Lefevre, J.-F., and Abdallah, M. A. (1988) Biochemistry 37, 15965-15973
  60. Du, F., Mao, X.-A., Li, D.-F., and Liao, Z.-R. (2001) J. Inorg. Biochem. 83, 101-105[CrossRef][Medline] [Order article via Infotrieve]
  61. El-Mahdaoui, L., and Tajmir-Riahi, H. A. (1995) J. Biomol. Struct. Dyn. 13, 69-86[Medline] [Order article via Infotrieve]
  62. Sorokin, V. A., Sysa, I. V., Valeev, V. A., Gladchenko, G. O., Degtyar, M. V., and Blagoi, Y. P. (1997) J. Mol. Struct. 408/409, 233-236
  63. Rizzo, V., Pande, A., and Luisi, P. L. (1987) in Pyridine Nucleotide Coenzymes (Dolphin, D., Poulson, R., and Avramovi, O., eds) Vol. A, pp. 99-161, John Wiley and Sons, New York
  64. Liu, G. D., Liao, J. P., Huang, S. S., Shen, G. L., and Yu, R. Q. (2001) Anal. Sci. 17, 1031-1036[CrossRef][Medline] [Order article via Infotrieve]
  65. Cusumano, M., and Giannetto, A. (1997) J. Inorg. Biochem. 65, 137-144[CrossRef][Medline] [Order article via Infotrieve]
  66. Gutman, M., Margalit, R., and Schejter, A. (1968) Biochemistry 7, 2778-2785[CrossRef][Medline] [Order article via Infotrieve]
  67. Gutman, M., and Eisenbach, M. (1973) Biochemistry 12, 2314-2317[CrossRef][Medline] [Order article via Infotrieve]
  68. Kapinos, L. E., Holy, A., Guenter, J., and Siegel, H. (2001) Inorg. Chem. 40, 2500-2508[CrossRef][Medline] [Order article via Infotrieve]
  69. Woodmansee, A. N., and Imlay, J. A. (2002) J. Biol. Chem. 277, 34055-34066[Abstract/Free Full Text]
  70. Gerli, G. C., Beretta, L., Bianchi, M., Agostoni, A., Gualandri, V., and Orsini, G. B. (1982) Scand. J. Haematol. 29, 141-146[Medline] [Order article via Infotrieve]
  71. Scott, M. D., Wagner, T. C., and Chiu, D. T.-Y. (1993) Biochim. Biophys. Acta 1181, 163-168[Medline] [Order article via Infotrieve]
  72. Goth, L., and Eaton, J. W. (2000) Lancet 356, 1820-1821[CrossRef][Medline] [Order article via Infotrieve]
  73. Lee, S. M., Koh, H.-J., Park, D.-C., Song, B. J., Huh, T.-L., and Park, J.-W. (2002) Free Radic. Biol. Med. 32, 1185-1196[CrossRef][Medline] [Order article via Infotrieve]
  74. Zheng, M., Wang, X., Templeton, L. J., Smulski, D. R., LaRossa, R. A., and Starz, G. (2001) J. Bacteriol. 183, 4562-4570[Abstract/Free Full Text]

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