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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Free Radic Res. Author manuscript; available in PMC 2013 August 14.
Published in final edited form as:
PMCID: PMC3690130

Reversible inactivation of dihydrolipoamide dehydrogenase by mitochondrial hydrogen peroxide


Under oxidative stress conditions, mitochondria are the major site for cellular production of reactive oxygen species (ROS) such as superoxide anion and H2O2 that can attack numerous mitochondrial proteins including dihydrolipoamide dehydrogenase (DLDH). While DLDH is known to be vulnerable to oxidative inactivation, the mechanisms have not been clearly elucidated. The present study was therefore designed to investigate the mechanisms of DLDH oxidative inactivation by mitochondrial reactive oxygen species (ROS). Mitochondria, isolated from rat brain, were incubated with mitochondrial respiratory substrates such as pyruvate/malate or succinate in the presence of electron transport chain inhibitors such as rotenone or antimycin A. This is followed by enzyme activity assay and gel-based proteomic analysis. The present study also examined whether ROS-induced DLDH oxidative inactivation could be reversed by reducing reagents such as DTT, cysteine, and glutathione. Results show that DLDH could only be inactivated by complex III- but not complex I-derived ROS; and the accompanying loss of activity due to the inactivation could be restored by cysteine and glutathione, indicating that DLDH oxidative inactivation by complex III-derived ROS was a reversible process. Further studies using catalase indicate that it was H2O2 instead of superoxide anion that was responsible for DLDH inactivation. Moreover, using sulfenic acid-specific labeling techniques in conjunction with two-dimensional Western blot analysis, we show that protein sulfenic acid formation (also known as sulfenation) was associated with the loss of DLDH enzymatic activity observed under our experimental conditions. Additionally, such oxidative modification was shown to be associated with preventing DLDH from further inactivation by the thiol-reactive reagent N-ethylmaleimide. Taken together, the present study provides insights into the mechanisms of DLDH oxidative inactivation by mitochondrial H2O2.

Keywords: brain, dihydrolipoamide dehydrogenase, H2O2, mitochondria, reactive oxygen species, reversible inactivation, sulfenic acid, sulfenation


Dihydrolipoamide dehydrogenase (DLDH) is the third catalytic enzyme of mitochondrial α-keto acid dehydrogenase complexes including pyruvate dehydrogenase complex, α-ketoglutarate dehydrogenase complex, and branched chain amino acid dehydrogenase complex [1]. It is also a component in the glycine cleavage system [2]. In vivo, DLDH catalyzes the reoxidation of dihydrolipoamide using NAD+ as the electron acceptor [3]. In vitro, however, the enzyme can also catalyze the reduction of lipoamide back to dihydrolipoamide using NADH as the electron donor [3]. DLDH also possesses diaphorase activity that can transport electrons from NADH to a variety of artificial electron acceptors such as nitro blue tetrazolium (NBT) [4] and dichlorophenolindophenol (DCPIP) [3].

DLDH is a redox-sensitive, multifunctional oxidoreductase and is very susceptible to oxidative modification. It has been reported that DLDH can be attacked by lipid peroxidation byproducts [5], can be nitrated by reactive nitrogen species [6,7], and can also be carbonylated by reactive oxygen species (ROS) [8]. DLDH has two redox-reactive cysteine residues at its active center that are indispensible for its catalytic function [2,9]. However, as redox-reactive cysteine residues are susceptible to oxidative modifications [1012], the two cysteine residues could also render DLDH vulnerable to oxidative inactivation [4,13]. Indeed, it has been reported that DLDH can undergo poly(ADP-ribosyl)ation and s-nitrosylation [1419] by reactive nitrogen species under a variety of experimental conditions. Additionally, depending on experimental systems, DLDH not only can augment oxidative stress [2027], but can also attenuate oxidative stress [28,29]. Interestingly, DLDH may also play a role in protective response as its expression has been shown to be significantly upregulated, respectively, in estrogen signaling [30] and in human heart failure [31].

Mitochondria are the major source of endogenous ROS [32] and both complexes I and III in the electron transport chain have been recognized as the enzyme systems that generate ROS [33]. While DLDH from a variety of organisms can be oxidatively modified with a loss in enzyme activity by exogenous oxidants [3437], the mechanisms of its inactivation by endogenous, mitochondria-generated ROS remain unknown. Moreover, in proteomic studies whereby DLDH was identified to be oxidatively modified [58,15], no effort was ever made to examine the effects of such modifications on enzyme activity. The purpose of this study was, therefore, to elucidate the mechanisms of DLDH oxidative inactivation by mitochondrial ROS. Specifically, using isolated mitochondria as a stand-alone system, we wanted to address whether DLDH is sensitive to oxidative inactivation by complex I- or complex III-derived ROS, whether the inactivation is reversible, and what would be the actual reactive species and the nature or the adduct of the oxidative modifications.

Materials and methods

Animals and chemicals

Adult male Sprague-Dawley rats obtained from Harlan (Indianapolis, IN) were used throughout the studies. Bis-Tris was purchased from Calbiochem (La Jolla, CA, USA). Tricine and ε-amino-N-caproic acid were purchased from MP Biochemicals. Acrylamide/bisacrylamide, ammonium persulfate, and CBB R-250 were purchased from Bio-Rad laboratories (Richmond, CA, USA). BSA, NADH, EDTA, lipoamide, N-ethylmaleimide (NEM) and biotin-NEM, cysteine, glutathione (GSH), sodium arsenite, and NBT chloride tablets were purchased from Sigma (St. Louis, MO, USA). Serva Blue G was from Serva (Heidelberg, Germany). Bicinchoninic acid protein assay kit was purchased from Pierce (Rockford, IL, USA). Rabbit anti-DLDH polyclonal antibodies (IgG) were from US Biological (Swampscott, MA, USA) and goat anti-rabbit IgG conjugated with horseradish peroxidase were from Invitrogen (San Diego, CA, USA). Hybond-C membrane and an ECL immunochemical detection kit were obtained from GE Healthcare (Piscataway, NJ, USA). Dihydrolipoamide was prepared using sodium borohydride reduction of lipoamide as previously described [3,38]. DCP-Bio-1, a specific sulfenic acid probe [39], was purchased from KetaFAST (Boston, MA).

Preparation of brain mitochondria

Rat brain mitochondria were isolated as previously described [40] with slight modifications [4,13]. Briefly, brains were removed rapidly and homogenized in 15 ml of ice-cold, mitochondrial isolation buffer containing 0.32 M sucrose, 1 mM EDTA and 10 mM Tris-HCl, pH 7.1. The homogenate was centrifuged at 1,330 g for 10 min and the supernatant was saved. The pellet was resuspended in 0.5 (7.5 ml) volume of the original isolation buffer and centrifuged again under the same conditions. The two supernatants were combined and centrifuged further at 21,200 g for 10 min. The resulting pellet was resuspended in 12% Percoll solution that was prepared in mitochondrial isolation buffer and centrifuged at 6,900 g for 10 min. The resulting supernatant was then carefully removed by vacuum. The obtained soft pellet was resuspended in 10 ml of the mitochondrial isolation buffer and centrifuged again at 6,900 g for 10 min. All isolated mitochondria were used fresh and protein concentrations were determined by bicinchoninic acid assay [41].

In vitro mitochondrial oxidative stress

Mitochondrial oxidative stress in vitro was induced by supplementing mitochondria with respiratory substrates such as pyruvate/malate or succinate in the presence of electron transport chain inhibitors such as rotenone or antimycin A, a condition that is known to enhance mitochondrial ROS generation [13,42,43]. Mitochondrial incubation was carried out as previously described [44]. Briefly, mitochondria (0.25 mg/ml) were incubated at 25°C for 60 min in incubation buffer (110 mM mannitol, 10 mM KH2PO4, 60 mM Tris, 60 mM KCl, and 0.5 mM EGTA, pH 7.4) in the presence of 50 μM rotenone or 50 μM antimycin A. The mixture was then supplemented with either complex I substrates pyruvate/malate (5 mM each) or complex II substrate succinate (10 mM). Control mitochondria were incubated under the same conditions in the absence of any substrates and inhibitors. At the end of the incubation, mitochondria were pelleted by centrifugation at 8,000 g for 10 min followed by enzyme assays or sulfenic acid labeling. For further incubation of the mitochondrial samples with reducing reagents such as DTT, cysteine, and GSH, 10 mM of each of the chemicals was added to the mixture and the sample was incubated at room temperature for an additional 30 min followed by measurement of enzyme activities. For evaluation of the effect of catalase on DLDH oxidative modification induced by antimycin A/succinate, broken mitochondria, prepared by resuspending in 50 mM potassium phosphate buffer (pH 7.4) followed by sonication, were used so that catalase can readily get access to any formed hydrogen peroxide.

Measurement of total ROS and superoxide anion

Levels of total mitochondrial ROS following incubation with substrates and electron transport chain inhibitors were measured by the fluorescence probe DCFH as previously described with slight modifications [45]. Following in vitro oxidative stress, 1 μM DCFH (in dimethyl sulfoxide) was incubated with 1 ml of 50 μg mitochondrial proteins for 1 hr at room temperature. Fluorescence intensity was read by a fluorometer equipped with a 96-well plate reader at an excitation wavelength of 485 nm and an emission wavelength of 535 nm. Measurement of superoxide anion by MitoSOX (from Invitrogen) was performed as previously described [46]. Additionally, superoxide generation by sub-mitochondrial particles was also measured by the method that involves SOD-inhibitable reduction of acetylated cytochrome c [47].

Determination of enzyme activities

Dehydrogenase activity was measured by DLDH catalyzed, NAD+-dependent oxidation of dihydrolipoamide [3,38]. The final volume of reaction was 1 ml, and the mixture contained 100 mM potassium phosphate, pH 8.0, 1.0 mM EDTA, 0.6 mg/ml BSA, 3.0 mM NAD+, 5–10 μg/ml mitochondrial extract and 3.0 mM dihydrolipoamide. A solution containing all the assay components except dihydrolipoamide was used as the blank. The reaction was initiated by the addition of dihydrolipoamide and the change in absorbance at 340 nm was followed at room temperature. DLDH diaphorase activity was performed using blue native polyacrylamide gel electrophoresis (BN-PAGE) as previously described [4].

Labeling of protein sulfenic acids with biotin-maleimide

Labeling and detection of protein sulfenic acids was performed as previously described [48]. Mitochondrial pellet, immediately following in vitro incubation, was solubilized in a thiol-group blocking buffer containing 100 mM sodium acetate (pH 7.0), 20 mM NaCl, 1% SDS and 100 mM N-ethylmaleimide (NEM). The protein mixture was incubated on a rotator at room temperature for 2 hrs followed by clarification of the mixture by centrifugation at 13,000 g for 10 min. Excess NEM in the supernatant was removed by gel filtration using PD-10 columns. This was followed by addition of 0.1 mM biotin-maleimide and 20 mM sodium arsenite (both final concentrations) to the eluate. The sample was further incubated on a rotator at room temperature for 30 min. Proteins were then precipitated by 10% TCA (final concentration) on ice for 10 min followed by centrifugation on a bench top centrifuge at 3,000 rpm for 5 min. The pellet was washed three times with ethyl acetate: ethanol (1:1, v/v). Protein pellet after the third wash was used for polyacrylamide gel electrophoresis as described below.

Polyacrylamide gel electrophoresis and Western blot analysis

One dimensional Laemmli SDS-PAGE (7.5% resolving gel) was performed except that the running buffer used was that of Tricine SDS-PAGE (100 mM Tris, 100 mM Tricine, 0.1% SDS, pH 8.3) [49]. Two dimensional-PAGE was performed as previously described [19,50]. Proteins on gels were stained with Coomassie colloid blue [51], and Western blot analysis was performed according to standard procedures.

Mass spectrometry analysis of DLDH cysteine modifications

For MS analysis of cysteine oxidative modifications in DLDH, gel bands resolved by BN-PAGE were excised and destained with 50 mM ammonium bicarbonate prepared in 50% methanol. This was followed by rehydration and trypsin digestion according to standard procedures. Peptides were analyzed using nano-LC/MS/MS system comprising a nano-LC pump and a LTQ-FT mass spectrometer (Thermo Electron Corporation, San Jose, CA) equipped with a nanospray ion source. Approximately 5 to 20 fmoles of tryptic digest samples were dissolved in 5% acetonitrile with 0.1% formic acid and injected onto a C18 nanobore LC column for nano-LC/MS/MS and identification of peptides. A linear gradient LC profile was used to separate and elute peptides with a constant total flow rate of 350 nL/minute. The gradient consisted of 5 to 70% solvent B in 78 minutes (solvent B: 80% acetonitrile with 0.1% formic acid; solvent A: 5% acetonitrile with 0.1% formic acid). Mascot database searches were performed using Bioworks Browser 3.2 software (Thermo Electron Corporation, San Jose, CA). Identified peptides were generally accepted only when the MASCOT ion score value exceeded 20.

Gel image documentation and data analysis

All images were scanned by an EPSON PERFECTION 1670 scanner. Densitometric analysis of gel bands was performed using Image J software. For enzyme activities, data were presented as mean ± SEM of triplicate assays. P < 0.05 was considered significant using Weltch’s t-test.


DLDH inactivation by complex III-derived ROS

To investigate the mechanisms of DLDH inactivation by mitochondrial ROS, isolated mitochondria were incubated with respiratory substrates in the presence of inhibitors of electron transport chain. This approach is an in vitro incubation system that is known to elevate mitochondrial ROS production [13,42,43]. In the present study, pyruvate/malate were used as complex I substrates and succinate was used as complex II substrate. The inhibitors used were rotenone for complex I and antimycin A for complex III, respectively. Complex III inhibition by antimycin A would cause ROS production by both complex III and complex I as this inhibition would cause accumulation and leakage of electrons at both of the two complexes [33,52]. In contrast, complex I inhibition by rotenone would only induce ROS production by complex I as electrons only accumulate and leak at complex I [33,52]. Following incubation, mitochondrial extracts were prepared and DLDH activities were measured. Results show that, regardless of the substrates that were used, DLDH only exhibited inactivation of its dehydrogenase activity when antimycin A was present (Figure 1A) and antimycin A itself (50 μM), in the absence of a respiratory substrate, did not impair DLDH activity (Figure 1B). Additionally, DLDH diaphorase activity analyzed by BN-PAGE shows a similar inactivation pattern to that of dehydrogenase activity (Figure 2). These results indicate that DLDH could only be inactivated by complex III-derived ROS under our experimental conditions.

Figure 1
(A) DLDH dehydrogenase activity following in vitro mitochondrial oxidative stress induced by electron transport chain inhibitors rotenone or antimycin A in the presence of pyruvate/malate (complex I substrates) or succinate (complex II substrate). *<0.05 ...
Figure 2
DLDH diaphorase activity following in vitro mitochondrial oxidative stress induced by electron transport chain inhibitors rotenone or antimycin A in the presence of pyruvate/malate (complex I substrates) or succinate (complex II substrate). A shows diaphorase ...

The above observation that DLDH could only be inactivated by complex III-derived ROS suggests that complex I either does not produce ROS or produces ROS that can not get access to DLDH. To determine which of the two situations actually occurred in our experiments, we measured mitochondrial ROS by DCFH assay following the same incubation conditions as described in Figures 1 and and2.2. Results in Figure 3A show that both complexes I and III produced a significant amount of ROS. These results indicate that complex I indeed produced ROS in the presence of rotenone or antimycin A, but these ROS could not inactivate DLDH. Additionally, when the superoxide-specific probe MitoSox [46] was used to measure the rate of superoxide production, it was found that the rate of superoxide production was linearly correlated with antimycin A-concentrations (Figure 3B) whereby succinate concentration (10 mM) remained constant. Moreover, this linear relationship was further confirmed when mitochondrial superoxide production was measured by the method involving SOD-inhibitable reduction of acetylated cytochrome c using submitochondrial particles as the source of electron transport chain [47] (Figure 3C), which further demonstrates an authentic involvement of superoxide anion in our experimental system.

Figure 3
(A) Levels of mitochondrial ROS determined by DCFH assay. Following incubation of mitochondria under the indicated conditions, 50 μg mitochondrial proteins was incubated with 1 μM DCFH (prepared in DMSO) at room temperature for 60 min. ...

DLDH is inactivated by H2O2

It is well known that the initial reactive oxygen species generated by complex III, when stimulated by AA/succinate, is superoxide anion [42]. In the presence of mitochondrial superoxide dismutase whereby whole mitochondrial extract is used, the released superoxide can be quickly dismutated to yield H2O2 [52]. On the other hand, for a cysteine residue to undergo oxidative modification, it has to exist in the thiolate anion form at physiological pH [53]. Therefore it is unlikely that superoxide anion that owns a negative charge can directly attack a thiolate anion that is also negatively charged. We reasoned that it is H2O2 that inactivates DLDH under our experimental conditions. To test this possibility, we incubated broken mitochondria with antimycin A and succinate in the presence of catalase (1 mg/ml), which was followed by DLDH enzyme activity assay. Result in Figure 4A shows that catalase completely abolished DLDH inactivation, demonstrating that DLDH was indeed inactivated by H2O2 under our experimental conditions.

Figure 4
(A) Catalase abolished DLDH inactivation when broken mitochondria were incubated with 50 μM antimycin A and 10 mM succinate for 30 min at room temperature; catalase used was 1 mg/ml. (B) Loss of dehydrogenase activity by H2O2 could be restored ...

DLDH inactivation by H2O2 can be reversed by cysteine and glutathione

To determine whether DLDH inactivation by complex III-generated H2O2 was reversible or not, we further added reducing reagents to the mitochondria incubation mixture. We chose DTT, cysteine, and glutathione as our reducing reagents as we suspected that the inactivation could be due to DLDH cysteine modifications. Results in Figure 4B show that DLDH inactivation could be completely reversed by cysteine and GSH, but not by DTT. Similarly, loss of DLDH diaphorase activity could also be restored by cysteine and GSH (Figure 5). These results suggest that DLDH inactivation by complex III H2O2 is caused by its cysteine modifications and is a reversible process.

Figure 5
Loss of diaphorase activity by H2O2 could be restored by cysteine and GSH. Following incubation of mitochondria with succinate and antimycin A, 10 mM of each of the three reducing reagents was added and the sample was further incubated at room temperature ...

DLDH inactivation by H2O2 involves sulfenic acid formation

Based on facts that glutathione reductase is structurally similar to DLDH [1] and glutathione reductase undergoes sulfenation under oxidative stress conditions [11], we suspected that our observed reversible DLDH inactivation by H2O2 may also be caused by protein sulfenic acid formation or sulfenation as this modification is a reversible process that can regulate protein function [54,55]. To determine whether this is the case, we first took a biotin switch approach [48] whereby protein sulfenic acids were reduced by arsenite back to free cysteine residues and the newly generated cysteine thiol groups were then labeled by biotin-maleimide (NEM). This was followed by 2D analysis of biotinylated proteins and DLDH localization by anti-DLDH Western blot. Results in Figure 6 show that DLDH was indeed sulfenated in the presence of succinate and antimycin A (panel 3). Additionally, we demonstrated that the arsenite-reducing method is indeed specific toward protein sulfenic acids as arsenite treatment performed after pre-incubation of samples with dimedone, a sulfenic acid specific reagent [56], largely abolished the immunoblot signals (panel 4, the middle lane).

Figure 6
Two-dimensional gel electrophoretic analysis of DLDH sulfenation. Sulfenated proteins were treated with arsenite and labeled with biotin-NEM followed by 2D analysis. (1): a representative 2D map visualized by Coomassie blue staining; (2): pattern of protein ...

Loss of DLDH activity correlates with increase in the level of DLDH sulfenation

If sulfenation is involved in the loss of DLDH activity, a correlation between loss of DLDH activity and increase in DLDH sulfenation would be expected. To determine whether this is the case, we then incubated mitochondrial samples with a sulfenic acid-specific probe DCP-Bio1 [39] after oxidative stress triggered by AA/succinate. This was followed by measurement of DLDH activity and separation of DLDH from other mitochondrial proteins by BN-PAGE [4]. The corresponding DLDH band on blue native gel was then excised and further analyzed by Western blot probed with streptavidin-HRP followed by densitometric quantitation of the signal intensities. Figure 7A shows a significant inactivation of DLDH by the system of AA/succinate; Figure 7B shows comparison of Western blot analysis of DLDH sulfenation between control and AA/succinate-treated samples; Figure 7C shows quantitatively that there was a significant increase in the level of sulfenation after treatment with AA/succinate. Taken together, these results indicate that loss of DLDH activity is inversely related with increase in the level of DLDH sulfenation.

Figure 7
Loss of DLDH activity is correlated with increase in DLDH sulfenation. (A) Inactivation of DLDH after incubation with AA (50 μM) and succinate (10 mM); control samples contain neither AA nor succinate. (B) DLDH sulfenation detected by DCP-Bio1. ...

Mass spectrometry analysis of DLDH cysteine residues that are oxidized under oxidative stress conditions

To determine which cysteine residues in DLDH underwent oxidative modifications that might be involved in the loss of enzyme activity, we performed mass spectrometry (MS) analysis. Following BN-PAGE resolution of both control and oxidized samples, DLDH-containing gel bands were excised and subjected to MS analysis. Results show that no sulfenic acid or sulfinic acid formation could be detected. In contrast, sulfonic acid formation on cys-449 was detected for both control and oxidized samples; while that on cys277 could only be detected in the oxidized samples. These results indicate that cys-449 likely underwent auto-oxidation during sample preparations and cys-277 was over-oxidized only after in vitro oxidative stress. Modifications of these two cysteine residues are unlikely to impair DLDH function as there have been no reports that cys-227 and cys-449 are redox-sensitive. It should be noted that the status of the two redox-sensitive cysteine residues (cys-45 and cys-50) at the enzyme’s active center could not be analyzed as the peptides containing the two cysteine residues failed to be recovered.

DLDH oxidative inactivation by H2O2 is resistant to further inactivation by N-ethylmaleimide (NEM)

We have previously shown that DLDH can lose its activity upon treatment with thiol-reactive reagent such as NEM [4]. If DLDH oxidative inactivation in our preparation indeed occurred due to sulfenic acid formation on one of the two catalytic cysteine residues (Cys-45 or Cys-50) located at the active center, then this modification should prevent DLDH against further inactivation by NEM that is known to react primarily with either of the two cysteine residues [5759]. To test this possibility, we treated mitochondria with varying concentrations of NEM following incubation with antimycin A/succinate. Result in Figure 8A shows that DLDH oxidative inactivation by mitochondrial H2O2 indeed protected DLDH from further inactivation by NEM. Interestingly, NEM could not further inactivate the enzyme even at a 2 mM concentration. The reason for this remains unknown, but it is not without precedent as glutathionylated α-ketoglutarate dehydrogenase is also resistance to further inactivation by NEM [60]. In contrast, such a resistance could not be observed when similar mitochondrial preparations were further treated by guanidine chloride that can induce protein denaturation [61] and is known to bind to multiple sites on DLDH [62] (Figure 8B). These results imply that DLDH sulfenation in our system indeed occurred on one of the two catalytic, redox-reactive cysteine residues.

Figure 8
DLDH oxidative inactivation by mitochondrial H2O2 prevented against further inactivation by NEM (A) but not guanidine chloride (B). Mitochondrial oxidative stress and DLDH inactivation was induced by incubation with antimycin A/succinate as described ...


The major findings of the present study are that, under our experimental conditions using isolated mitochondria as a stand-alone system, DLDH could only be inactivated by H2O2 derived from complex III-, but not from complex I; and this inactivation could only be reversed by cysteine and GSH, but not by DTT. The present study has also demonstrated that DLDH inactivation by H2O2 likely involved protein sulfenic acid formation that could prevent DLDH from further alkylation by NEM. Additionally, as sulfenic acid formation is a reversible process [54], our results could also suggest that sulfenic acid formation may serve as a switch for DLDH activity under oxidative stress conditions.

It should be pointed out that in our in vitro oxidative stress system, both complexes I and III were capable of generating superoxide anion/H2O2 in the presence of anti-mycin A, regardless of the substrates that were supplemented (Figure 3). However, it was found that DLDH could only be inactivated by complex III-derived H2O2. Such results suggest that DLDH is physically associated with complex III. Indeed, when BN-PAGE-resolved DLDH gel band was processed by LC-Nano MS/MS for protein identification, ubiquinol cytochrome c reductase subunit 2 (a component of complex III) was found in the DLDH gel band while no complex I components could be detected [63]. Therefore, DLDH is only accessible to, and may only be inactivated by, complex III-derived H2O2. Alternatively, as it is well known that H2O2 can easily diffuse from one place to another in mitochondria, our observation that DLDH could not be inactivated by complex I-generated H2O2 suggests that there may be an indirect mechanism involving H2O2 but not by H2O2 directly.

Interestingly, while our data clearly demonstrate the reversibility of DLDH inactivation by mitochondrial H2O2, our data also show that, among the reducing reagents that were tested, only cysteine and GSH could restore the enzyme’s activity following oxidative inactivation. In contrast, DTT, a very efficient thiol protector [64], could not do so under our experimental conditions. These data are in agreement with previous findings that both GSH [65] and cysteine [66] can readily enter mitochondria, while DTT can not [67]. Therefore, DTT can not get access to the sulfenated cysteine residue(s) when intact mitochondria are oxidatively stressed. It should be noted that when pure DLDH isolated from pig heart was inactivated by metal-catalyzed oxidation, the inactivation could indeed be reversed by DTT, at least partially [68], indicating that DTT is able to get access to the oxidatively modified amino acid residues when purified DLDH is oxidized.

Rat DLDH contains a total of 10 cysteine residues [69]; and cys-45 and cys-50 are the two redox-sensitive cysteines located at the enzyme’s active center [13]. During catalysis, cys-45 binds substrate dihydrolipoamide while cys-50 interacts with FAD [2]. While we were able to determine two cysteine residues (cys-277 and cys-449) that were sulfonated, and one of them (cys-277) was sulfonated only after in vitro oxidative stress, we were unable to determine the status of oxidative modifications to cys-45 and cys-50 because the peptides containing the two cysteines failed to be recovered following the procedures of mass spectrometry analysis. Nevertheless, our results that DLDH oxidative inactivation prevented against further inactivation by the thiol-reactive reagent NEM would suggest that sulfenation occurred on one of the two catalytic cysteine residues (Figure 8A). Additionally, based on studies that cys-45 is more reactive than cys-50 toward thiol-reactive reagents [59], we reason that cys-45 would undergo sulfenation under our experimental conditions. This reasoning would also be in agreement with the findings that, in glutathione reductase that is structurally similar to DLDH [1], only the substrate binding cysteine residue can undergo oxidative modification when challenged by reactive nitrogen species [11]. Nevertheless, further studies will be needed to determine which of the two redox-sensitive cysteine residues at the active center is modified under oxidative stress conditions.

Additionally, if sulfenation is the main cause of DLDH inactivation, there should be a positive correlation between the level of DLDH sulfenation and the loss in DLDH activity. Indeed, after mitochondrial oxidative stress by AA/succinate, loss of DLDH activity was found to be inversely correlated with an increase in the level of DLDH sulfenation (Figure 7). Furthermore, we also attempted to test such a correlation using partially purified DLDH that was incubated with varying concentrations of H2O2 followed by labeling with DCP-Bio1. DLDH was partially purified from rat liver according to methods previously described [7072]. Enzyme activity was also measured spectrophotometrically after DCP-Bio1 labeling. Results in Supplementary Figure 1A to be found online at show that while DLDH activity decreased further with increasing concentrations of H2O2, DLDH sulfenic acid content, reflected by Western blot assay, actually decreased in an H2O2 concentration-dependent manner, indicating that purified DLDH had already been sulfenated and further oxidation by H2O2 over-oxidized the formed sulfenic acids, resulting in less and less labeling by DCP-Bio1 (Supplementary Figure 1B). In addition, while the maximum sulfenic acid signal intensity in the Western blot was obtained at 10 μM H2O2, nearly no inhibition of enzyme activity could be observed at this concentration of H2O2 (Supplementary Figure 1). Therefore, with the use of purified DLDH, assessment of a positive relationship between DLDH sulfenation and loss of enzyme activity did not yield expected results, presumably because of autooxidation during purification and overoxidation of protein sulfenic acids by H2O2. Nonetheless, our studies using partially purified DLDH further indicate a genuine involvement of transitional sulfenic acid formation in DLDH oxidative modifications.

While it has been established that protein sulfenic acid plays a central role in protein thiol oxidation and is mainly produced by reactions between redox-sensitive cysteine residues and hydrogen peroxide, alky hydroperoxides, as well as peroxynitrite [48,54,55,73,74]; it is also known that sulfenic acid is a key intermediate that can generate the most commonly reversible cysteine modification products including protein disulfides, mixed disulfides or S-glutathionylation [75]. Nevertheless, there is now accumulating evidence that stabilized sulfenic acids do exist and play a key redox regulatory role in many aspects of cell biology [53,76]. For example, protein S-sulfenation has been found to be a prerequisite for T cell activation, proliferation and function [77]. With respect to DLDH sulfenic acid formation in our system, the reason why DLDH sulfenic acids are stable enough to be detected remains unknown. Nonetheless, the stability of DLDH sulfenic acids could be controlled by a variety of factors such as solvent access, steric hindrance, the lack of proximal thiols, nearby available hydrogen-bonding interactions and/or the presence of positively charged side chains [74,78].

Finally, it should be alerted that the precise physiological or pathophysiological role of DLDH sulfenation is unknown at this time and could only be speculated. It is probable that this reversible modification, when occurring on one of the two cysteine residues at the enzyme’s active center, may protect DLDH from an irreversible oxidation and a permanent loss of enzyme activity (as shown in Figure 8A) provided that sulfenic acid would not be over-oxidized to form sulfinic or sulfonic acids that are irreversible. Moreover, DLDH sulfenation not only could act as a redox sink that preserves cellular antioxidant capacity, but could also serve as an on/off redox and metabolic switch used by cells to prevent against the occurrence of a widespread, severe, and irreversible cellular oxidative damage. Therefore, at early stage of oxidative stress, DLDH sulfenation is likely a trade-off mechanism by which mitochondria adapt to oxidative stress. It is also plausible that, under oxidative stress conditions whereby level of mitochondrial oxidant formation is elevated, DLDH inactivation via sulfenation could be part of mitochondrial oxidant-scavenging system in cellular metabolism, including mitochondrial superoxide dismutase and peroxiredoxins [79]. Hence, it is conceivable that DLDH sulfenation under oxidative stress conditions may be an adaptive or defensive response.

Supplementary Material

Supplmental Fig. 1


The authors thank Dr. Chad Nelson at the University of Utah for his assistance in mass spectrometry analysis of DLDH.

This work was supported in part by an NIH grant (AG022550) and a UNTHSC-UAEM seed grant (RI6044).


Declaration of interest

The authors report no declarations of interest. The authors alone are responsible for the content and writing of the paper.


1. Williams CH., Jr . Lipoamide dehydrogenase, glutathione reductase, thioredoxin reductase, and mercuric ion reductase-a family of flavoenzyme transhydrogenases. In: Muller F, editor. Chemistry and Biochemistry of Flavoenzymes. III. Boca Raton: CRC Press; 1992. pp. 121–212.
2. Vettakkorumakankav NN, Patel MS. Dihydrolipoamide dehydrogenase: structural and mechanistic aspects. Indian J Biochem Biophys. 1996;33:168–176. [PubMed]
3. Patel MS, Vettakkorumakankav NN, Liu TC. Dihydrolipoamide dehydrogenase: activity assays. Methods Enzymol. 1995;252:186–195. [PubMed]
4. Yan LJ, Yang SH, Shu H, Prokai L, Forster MJ. Histochemical staining and quantification of dihydrolipoamide dehydrogenase diaphorase activity using blue native PAGE. Electrophoresis. 2007;28:1036–1045. [PubMed]
5. Hussain SN, Matar G, Barreiro E, Florian M, Divangahi M, Vassilakopoulos T. Modifications of proteins by 4-hydroxy-2-nonenal in the ventilatory muscles of rats. Am J Physiol Lung Cell Mol Physiol. 2006;290:L996–1003. [PubMed]
6. Lee HM, Reed J, Greeley GH, Jr, Englander EW. Impaired mitochondrial respiration and protein nitration in the rat hippocampus after acute inhalation of combustion smoke. Toxicol Appl Pharmacol. 2009;235:208–215. [PMC free article] [PubMed]
7. Tyther R, Ahmeda A, Johns E, Sheehan D. Proteomic identification of tyrosine nitration targets in kidney of spontaneously hypertensive rats. Proteomics. 2007;7:4555–4564. [PubMed]
8. Tyther R, Ahmeda A, Johns E, Sheehan D. Protein carbonylation in kidney medulla of the spontaneously hypertensive rat. Proteomics Clin Appl. 2009;3:338–346. [PubMed]
9. Brautigam CA, Chuang JL, Tomchick DR, Machius M, Chuang DT. Crystal structure of human dihydrolipoamide dehydrogenase: NAD+/NADH binding and the structural basis of disease-causing mutations. J Mol Biol. 2005;350:543–552. [PubMed]
10. Barford D. The role of cysteine residues as redox-sensitive regulatory switches. Curr Opin Struct Biol. 2004;14:679–686. [PubMed]
11. Becker K, Savvides SN, Keese M, Schirmer RH, Karplus PA. Enzyme inactivation through sulfhydryl oxidation by physiologic NO-carriers. Nat Struct Biol. 1998;5:267–271. [PubMed]
12. Biswas S, Chida AS, Rahman I. Redox modifications of protein-thiols: emerging roles in cell signaling. Biochem Pharmacol. 2006;71:551–564. [PubMed]
13. Yan LJ, Thangthaeng N, Forster MJ. Changes in dihydrolipoamide dehydrogenase expression and activity during postnatal development and aging in the rat brain. Mech Ageing Dev. 2008;129:282–290. [PMC free article] [PubMed]
14. Pankotai E, Lacza Z, Muranyi M, Szabo C. Intra-mitochondrial poly(ADP-ribosyl)ation: potential role for alpha-ketoglutarate dehydrogenase. Mitochondrion. 2009;9:159–164. [PubMed]
15. Foster MW, Stamler JS. New insights into protein S-nitrosylation. Mitochondria as a model system. J Biol Chem. 2004;279:25891–25897. [PubMed]
16. Rhee KY, Erdjument-Bromage H, Tempst P, Nathan CF. S-nitroso proteome of Mycobacterium tuberculosis: enzymes of intermediary metabolism and antioxidant defense. Proc Natl Acad Sci U S A. 2005;102:467–472. [PubMed]
17. Ortega-Galisteo AP, Rodriguez-Serrano M, Pazmino DM, Gupta DK, Sandalio LM, Romero-Puertas MC. S-Nitrosylated proteins in pea (Pisum sativum L) leaf peroxisomes: changes under abiotic stress. J Exp Bot. 2012;63:2089–2103. [PMC free article] [PubMed]
18. Richardson AR, Payne EC, Younger N, Karlinsey JE, Thomas VC, Becker LA, et al. Multiple targets of nitric oxide in the tricarboxylic acid cycle of Salmonella enterica serovar typhimurium. Cell Host Microbe. 2011;10:33–43. [PMC free article] [PubMed]
19. Yan LJ, Liu L, Forster MJ. Reversible inactivation of dihydrolipoamide dehydrogenase by Angeli’s salt. Acta Biophysica Sinica. 2012;28:341–350. [PMC free article] [PubMed]
20. Bando Y, Aki K. Mechanisms of generation of oxygen radicals and reductive mobilization of ferritin iron by lipoamide dehydrogenase. J Biochem (Tokyo) 1991;109:450–454. [PubMed]
21. Sreider CM, Grinblat L, Stoppani AO. Catalysis of nitrofuran redox-cycling and superoxide anion production by heart lipoamide dehydrogenase. Biochem Pharmacol. 1990;40:1849–1857. [PubMed]
22. Gazaryan IG, Krasnikov BF, Ashby GA, Thorneley RN, Kristal BS, Brown AM. Zinc is a potent inhibitor of thiol oxidoreductase activity and stimulates reactive oxygen species production by lipoamide dehydrogenase. J Biol Chem. 2002;277:10064–10072. [PubMed]
23. Tahara EB, Barros MH, Oliveira GA, Netto LE, Kowaltowski AJ. Dihydrolipoyl dehydrogenase as a source of reactive oxygen species inhibited by caloric restriction and involved in Saccharomyces cerevisiae aging. Faseb J. 2007;21:274–283. [PubMed]
24. Ambrus A, Torocsik B, Tretter L, Ozohanics O, Adam-Vizi V. Stimulation of reactive oxygen species generation by disease-causing mutations of lipoamide dehydrogenase. Hum Mol Genet. 2011;20:2984–2995. [PubMed]
25. Zhang Q, Zou P, Zhan H, Zhang M, Zhang L, Ge RS, Huang Y. Dihydrolipoamide dehydrogenase and cAMP are associated with cadmium-mediated Leydig cell damage. Toxicol Lett. 2011;205:183–189. [PubMed]
26. Kareyeva AV, Grivennikova VG, Cecchini G, Vinogradov AD. Molecular identification of the enzyme responsible for the mitochondrial NADH-supported ammonium-dependent hydrogen peroxide production. FEBS Lett. 2011;585:385–389. [PMC free article] [PubMed]
27. Kareyeva AV, Grivennikova VG, Vinogradov AD. Mitochondrial hydrogen peroxide production as determined by the pyridine nucleotide pool and its redox state. Biochim Biophys Acta. 2012;1817:1879–1885. [PubMed]
28. Korotchkina LG, Yang H, Tirosh O, Packer L, Patel MS. Protection by thiols of the mitochondrial complexes from 4-hydroxy-2-nonenal. Free Radic Biol Med. 2001;30:992–999. [PubMed]
29. Igamberdiev AU, Bykova NV, Ens W, Hill RD. Dihydrolipoamide dehydrogenase from porcine heart catalyzes NADH-dependent scavenging of nitric oxide. FEBS Lett. 2004;568:146–150. [PubMed]
30. Nilsen J, Irwin RW, Gallaher TK, Brinton RD. Estradiol in vivo regulation of brain mitochondrial proteome. J Neurosci. 2007;27:14069–14077. [PubMed]
31. Li W, Rong R, Zhao S, Zhu X, Zhang K, Xiong X, et al. Proteomic analysis of metabolic, cytoskeletal and stress response proteins in human heart failure. J Cell Mol Med. 2012;16:59–71. [PMC free article] [PubMed]
32. Ames BN, Shigenaga MK. Oxidants are a major contributor to aging. Ann N Y Acad Sci. 1992;663:85–96. [PubMed]
33. St-Pierre J, Buckingham JA, Roebuck SJ, Brand MD. Topology of superoxide production from different sites in the mitochondrial electron transport chain. J Biol Chem. 2002;277:44784–44790. [PubMed]
34. Gutierrez-Correa J, Stoppani AO. Inactivation of heart dihydrolipoamide dehydrogenase by copper Fenton systems. Effect of thiol compounds and metal chelators. Free Radic Res. 1995;22:239–250. [PubMed]
35. Gutierrez-Correa J, Stoppani AO. Inactivation of myocardial dihydrolipoamide dehydrogenase by myeloperoxidase systems: effect of halides, nitrite and thiol compounds. Free Radic Res. 1999;30:105–117. [PubMed]
36. Gutierrez-Correa J. Trypanosoma cruzi dihydrolipoamide dehydrogenase as target of reactive metabolites generated by cytochrome c/hydrogen peroxide (or linoleic acid hydroperoxide)/phenol systems. Free Radic Res. 2010;44:1345–1358. [PubMed]
37. Gutierrez-Correa J, Stoppani AO. Myeloperoxidase-generated phenothiazine cation radicals inactivate Trypanosoma cruzi dihydrolipoamide dehydrogenase. Rev Argent Microbiol. 2002;34:83–94. [PubMed]
38. Patel MS, Hong YS. Lipoic acid as an antioxidant: the role of dihydrolipoamide dehydrogenase. In: Armstrong D, editor. Free Radical and Antioxidant Protocols. Totowa, NJ: Humana Press; 1998. pp. 337–346. [PubMed]
39. Poole LB, Klomsiri C, Knaggs SA, Furdui CM, Nelson KJ, Thomas MJ, et al. Fluorescent and affinity-based tools to detect cysteine sulfenic acid formation in proteins. Bioconjug Chem. 2007;18:2004–2017. [PMC free article] [PubMed]
40. Sims NR. Methods in Toxicology: Mitochondrial Dysfunction. 2. San Diego: Academic Press; 1993.
41. Smith PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, et al. Measurement of protein using bicinchoninic acid. Anal Biochem. 1985;150:76–85. [PubMed]
42. Turrens JF, Alexandre A, Lehninger AL. Ubisemiquinone is the electron donor for superoxide formation by complex III of heart mitochondria. Arch Biochem Biophys. 1985;237:408–414. [PubMed]
43. Turrens JF. Superoxide production by the mitochondrial respiratory chain. Biosci Rep. 1997;17:3–8. [PubMed]
44. Schonfeld P, Reiser G. Rotenone-like action of the branched-chain phytanic acid induces oxidative stress in mitochondria. J Biol Chem. 2006;281:7136–7142. [PubMed]
45. Yan LJ, Rajasekaran NS, Sathyanarayanan S, Benjamin IJ. Mouse HSF1 disruption perturbs redox state and increases mitochondrial oxidative stress in kidney. Antioxid Redox Signal. 2005;7:465–471. [PubMed]
46. Piacenza L, Irigoin F, Alvarez MN, Peluffo G, Taylor MC, Kelly JM, et al. Mitochondrial superoxide radicals mediate programmed cell death in Trypanosoma cruzi: cytoprotective action of mitochondrial iron superoxide dismutase overexpression. Biochem J. 2007;403:323–334. [PubMed]
47. Yan LJ, Christians ES, Liu L, Xiao X, Sohal RS, Benjamin IJ. Mouse heat shock transcription factor 1 deficiency alters cardiac redox homeostasis and increases mitochondrial oxidative damage. EMBO J. 2002;21:5164–5172. [PubMed]
48. Saurin AT, Neubert H, Brennan JP, Eaton P. Widespread sulfenic acid formation in tissues in response to hydrogen peroxide. Proc Natl Acad Sci U S A. 2004;101:17982–17987. [PubMed]
49. Yan LJ, Levine RL, Sohal RS. Effects of aging and hyperoxia on oxidative damage to cytochrome c in the housefly, Musca domestica. Free Radic Biol Med. 2000;29:90–97. [PubMed]
50. Yan LJ. Analysis of oxidative modification of proteins. Curr Protoc Protein Sci. 2009;Chapter 14(Unit 14):4. [PubMed]
51. Kang D, Gho YS, Suh M, Kang C. Highly sensitive and fast protein detection with Coomassie brilliant blue in sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Bull Korean Chem Soc. 2002;23:1511–1512.
52. Miwa S, St-Pierre J, Partridge L, Brand MD. Superoxide and hydrogen peroxide production by Drosophila mitochondria. Free Radic Biol Med. 2003;35:938–948. [PubMed]
53. Kettenhofen NJ, Wood MJ. Formation, reactivity, and detection of protein sulfenic acids. Chem Res Toxicol. 2010;23:1633–1646. [PMC free article] [PubMed]
54. Poole LB, Karplus PA, Claiborne A. Protein sulfenic acids in redox signaling. Annu Rev Pharmacol Toxicol. 2004;44:325–347. [PubMed]
55. Poole LB, Nelson KJ. Discovering mechanisms of signaling-mediated cysteine oxidation. Curr Opin Chem Biol. 2008;12:18–24. [PMC free article] [PubMed]
56. Poole LB. Formation and functions of protein sulfenic acids. Curr Protoc Toxicol. 2004;Chapter 17(Unit 17):1. [PubMed]
57. Nakamura M, Yamazaki I. One-electron transfer reactions in biochemical systems. VI. Changes in electron transfer mechanism of lipoamide dehydrogenase by modification of sulfhydryl groups. Biochim Biophys Acta. 1972;267:249–257. [PubMed]
58. Burleigh BD, Jr, Williams CH., Jr The isolation and primary structure of a paptide containing the oxidation-reduction active cystine of Escherichia coli lipoamide dehydrogenase. J Biol Chem. 1972;247:2077–2082. [PubMed]
59. Thorpe C, Williams CH., Jr Differential reactivity of the two active site cysteine residues generated on reduction of pig heart lipoamide dehydrogenase. J Biol Chem. 1976;251:3553–3557. [PubMed]
60. Applegate MA, Humphries KM, Szweda LI. Reversible inhibition of alpha-ketoglutarate dehydrogenase by hydrogen peroxide: glutathionylation and protection of lipoic acid. Biochemistry. 2008;47:473–478. [PubMed]
61. Monera OD, Kay CM, Hodges RS. Protein denaturation with guanidine hydrochloride or urea provides a different estimate of stability depending on the contributions of electrostatic interactions. Protein Sci. 1994;3:1984–1991. [PubMed]
62. Wilkinson KD, Williams CH., Jr Interactions of guanidinium chloride and pyridine nucleotides with oxidized and two-electron-reduced lipoamide dehydrogenase from Escherichia coli. J Biol Chem. 1979;254:863–871. [PubMed]
63. Yan LJ, Forster MJ. Resolving mitochondrial protein complexes using nongradient blue native polyacrylamide gel electrophoresis. Anal Biochem. 2009;389:143–149. [PMC free article] [PubMed]
64. Cleland WW. Dithiothreitol, a new protective reagent for SH groups. Biochemistry. 1963;3:480–482. [PubMed]
65. Shen D, Dalton TP, Nebert DW, Shertzer HG. Glutathione redox state regulates mitochondrial reactive oxygen production. J Biol Chem. 2005;280:25305–25312. [PubMed]
66. Stemmler TL, Lesuisse E, Pain D, Dancis A. Frataxin and mitochondrial FeS cluster biogenesis. J Biol Chem. 2010;285:26737–26743. [PMC free article] [PubMed]
67. Mesecke N, Terziyska N, Kozany C, Baumann F, Neupert W, Hell K, Herrmann JM. A disulfide relay system in the inter-membrane space of mitochondria that mediates protein import. Cell. 2005;121:1059–1069. [PubMed]
68. Gutierrez Correa J, Stoppani AO. Inactivation of lipoamide dehydrogenase by cobalt(II) and iron(II) Fenton systems: effect of metal chelators, thiol compounds and adenine nucleotides. Free Radic Res Commun. 1993;19:303–314. [PubMed]
69. Strausberg RL, Feingold EA, Grouse LH, Derge JG, Klausner RD, Collins FS, et al. Generation and initial analysis of more than 15,000 full-length human and mouse cDNA sequences. Proc Natl Acad Sci U S A. 2002;99:16899–16903. [PubMed]
70. Millard SA, Kubose A, Gal EM. Brain lipoyl dehydrogenase. Purification, properties, and inhibitors. J Biol Chem. 1969;244:2511–2515. [PubMed]
71. Ide S, Hayakawa T, Okabe K, Koike M. Lipoamide dehydrogenase from human liver. J Biol Chem. 1967;242:54–60. [PubMed]
72. Koike M, Hayakawa T. Purification and properties of lipoamide dehydrogenases from pig heart alpha-keto acid dehydrogenase complexes. Methods Enzymol. 1970;18:298–307.
73. Charles RL, Schroder E, May G, Free P, Gaffney PR, Wait R, et al. Protein sulfenation as a redox sensor: proteomics studies using a novel biotinylated dimedone analogue. Mol Cell Proteomics. 2007;6:1473–1484. [PubMed]
74. Salsbury FR, Jr, Knutson ST, Poole LB, Fetrow JS. Functional site profiling and electrostatic analysis of cysteines modifiable to cysteine sulfenic acid. Protein Sci. 2008;17:299–312. [PubMed]
75. Rehder DS, Borges CR. Cysteine sulfenic acid as an intermediate in disulfide bond formation and nonenzymatic protein folding. Biochemistry. 2010;49:7748–7755. [PMC free article] [PubMed]
76. Roos G, Messens J. Protein sulfenic acid formation: from cellular damage to redox regulation. Free Radic Biol Med. 2011;51:314–326. [PubMed]
77. Michalek RD, Nelson KJ, Holbrook BC, Yi JS, Stridiron D, Daniel LW, et al. The requirement of reversible cysteine sulfenic acid formation for T cell activation and function. J Immunol. 2007;179:6456–6467. [PubMed]
78. Claiborne A, Miller H, Parsonage D, Ross RP. Protein-sulfenic acid stabilization and function in enzyme catalysis and gene regulation. FASEB J. 1993;7:1483–1490. [PubMed]
79. Cox AG, Winterbourn CC, Hampton MB. Mitochondrial peroxiredoxin involvement in antioxidant defence and redox signalling. Biochem J. 2010;425:313–325. [PubMed]