Inactivation of the 2-oxo acid dehydrogenase complexes
upon generation of intrinsic radical species
Victoria I. Bunik
1
and Christian Sievers
2
1
A.N.Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia;
2
Physiological Chemistry
Institute of Eberhard-Karls-University, Tuebingen, Germany
Self-regulation of the 2-oxo acid dehydrogenase complexes
during catalysis was studied. Radical species as side products
of catalysis were detected by spin trapping, lucigenin fluor-
escence and ferricytochrome c reduction. Studies of the
complexes after converting the bound lipoate or FAD
cofactors to nonfunctional derivatives indicated that radicals
are generated via FAD. In the presence of oxygen, the 2-oxo
acid, CoA-dependent production of the superoxide anion
radical was detected. In the absence of oxygen, a protein-
bound radical concluded to be the thiyl radical of the
complex-bound dihydrolipoate was trapped by a-phenyl-
N-tert-butylnitrone. Another, carbon-centered, radical was
trapped in anaerobic reaction of the complex with 2-oxo-
glutarate and CoA by 5,5¢-dimethyl-1-pyrroline-N-oxide
(DMPO). Generation of radical species was accompanied by
the enzyme inactivation. A superoxide scavenger, superoxide
dismutase, did not protect the enzyme. However, a thiyl
radical scavenger, thioredoxin, prevented the inactivation. It
was concluded that the thiyl radical of the complex-bound
dihydrolipoate induces the inactivation by 1e

Abbreviations: E1, 2-oxo acid dehydrogenase; E2, dihydrolipoamide acyltransferase; E3, dihydrolipoamide dehydrogenase; DTPA, diethylene-
triaminepentaacetic acid; DMPO, 5,5¢-dimethyl-1-pyrroline-N-oxide; MNP, 2-methyl-2-nitrosopropane; PBN, a-phenyl-N-tert-butylnitrone;
POBN, a-(4-pyridyl-1-oxide)-N-tert-butylnitrone; ROS, reactive oxygen species; SOD, superoxide dismutase; ThDP, thiamin diphosphate.
(Received 31 May 2002, accepted 23 August 2002)
Eur. J. Biochem. 269, 5004–5015 (2002) Ó FEBS 2002 doi:10.1046/j.1432-1033.2002.03204.x
Depending on the particular 2-oxo acid, R- is CH
3
-(pyru-
vate), HOOC-(CH
2
)
2
- (2-oxoglutarate), CH
3
-CH(CH)
3
-
CH
2
- (2-oxoisovaleriate). Multiple copies of the substrate-
specific 2-oxo acid dehydrogenase (E1), dihydrolipoamide
acyltranferase (E2), and dihydrolipoamide dehydrogenase
(E3) are organized in highly ordered structures [1–4]. E1
catalyzes the rate-limiting step of the whole process [5–7]
and the active sites are coupled through the interacting
network of the lipoyl moieties [3,8]. The two electrons in the
catalytic intermediate of reactions 4 and 5, the 2e
)
reduced
E3, initially were thought to be distributed between flavin

reactive oxygen species (ROS) should not be neglected. ROS
have attracted increasing attention as cellular messengers
involved in differentiation, apoptosis and aging [25–27].
Further, treatment of cells with H
2
O
2
results in selective
oxidation of the 2-oxo acid dehydrogenase complexes [28],
thatcouldalsotakeplacein vivo upon site-specific
production of ROS via the complex-bound FAD. In vitro
and in situ inactivation of the 2-oxo acid dehydrogenase
complexes upon addition of organic hydroperoxides [29–31]
or a superoxide anion radical-generating system [32,33]
correlate with specific targeting and/or impaired function of
the complexes observed in many disorders linked to
mitochondrial and cell damage. These disorders include
poisoning with environmental toxicants [34,35], Alzheimer’s
[36,37] and Parkinson’s [38] diseases, Wernicke–Korsakoff
syndrome [39] and others. However, the mechanisms of
oxidative damage of the complexes and cellular protection
against this damage are not properly understood.
Because the redox state of cellular thiols and disulfides is
an important factor in cellular protection against oxidative
damage, we suggested that the 2-oxo acid dehydrogenase
complexes may be significant not only as catalytic systems,
but also as microcompartments of important biological
thiols, lipoate and CoA. The oligomeric complex cores form
an inner cavity for CoA. Depending on the source and type,
the cores consist of 24 (cube) or 60 (pentagonal dodeca-

CoA, diphenyliodonium chloride, succinyl-CoA, b-D-glu-
cose, N-ethyl maleimide, R,S-lipoamide, MNP, DTPA,
glutathione disulfide, copper-zinc superoxide dismutase
(from bovine erythrocytes, 3500 UÆmg
)1
), glucose oxidase
(from Aspergillus niger, 250 UÆmg
)1
), cytochrome c (from
horse heart, type VI) were from Sigma (Deisenhofen,
Germany). 2-Oxoglutarate and pyruvate were from Merck
(Darmstadt, Germany). DMPO, PBN, POBN and lucigenin
were from Molecular Probes (Leiden, the Netherlands).
Catalase (from bovine liver, 260 000 UÆmg
)1
), ThDP,
NAD
+
, and NADH were from Roche Molecular Biochemi-
cals (Mannheim, Germany). Recombinant thioredoxin
from Escherichia coli was from Calbiochem-Novabiochem
GmbH (Bad Soden, Germany). R,S-Dihydrolipoamide was
obtained from R,S-lipoamide by sodium borohydride
reduction as in [47].
Enzyme isolation, assays and modification
2-oxoglutarate and pyruvate dehydrogenase complexes
from pig heart were isolated according to [52] with the
modifications described earlier [46]. The pyruvate dehy-
drogenase complex from E. coli wasisolatedasin[53].
Overall, E1 and E3 enzymatic activities were assayed

M
)in
0.1
M
potassium phosphate buffer, pH 7.0. The residual
activity of the E3 component after one hour of modification
was no more than 4% of its initial value. The modified
complexes were separated from the low molecular mass
compounds by desalting on a HiTrap
TM
5 mL column
(Pharmacia, Uppsala, Sweden) in 0.1
M
potassium phos-
phate, pH 7.0.
EPR spectroscopy
Room temperature EPR spectra were recorded in a quartz
flat EPR sample cell at X-Band using a Bruker 300E EPR
spectrometer (Karlsruhe, Germany). The spectrometer was
operated at modulation amplitude 1 mT, modulation
frequency 100 kHz, microwave energy 20 mW. The reac-
tions took place in 0.05
M
potassium phosphate buffer,
pH 7.0. The stock solutions of MNP, DMPO, POBN and
PBN were freshly prepared and kept protected from light.
Controls in each experiment indicated that none of the
components alone produced the EPR signals studied.
Model reactions of thiyl radical trapping PBN took place
in a reaction medium containing excess dithiothreitol and

to 80% saturation. In anaerobic experiments, manipula-
tions of the samples were performed in the glove box. The
precipitating agents were added to the probe after recording
its EPR spectrum. The precipitated protein was centrifuged
for several minutes in a sealed Eppendorf tube, the pellet
washed with saturated ammonium sulfate and the centri-
fugation repeated. A stable and measurable EPR signal to
compare the reactions in the presence and absence of
O
2
under otherwise equal conditions was achieved by
employing high initial concentration of the 2-oxo acid
dehydrogenase complexes (30–200 mgÆmL
)1
), so that the
concentrations of substrates were comparable to that of the
protein redox centers.
Lucigenin-dependent fluorescence
This was measured with a Berthold LB 9505C Lumino-
meter (Germany). Reaction mixtures of 1 mL contained
2.5 mgÆmL
)1
2-oxoglutarate dehydrogenase complex,
27 l
M
lucigenin, 5 m
M
CoA, 5 m
M
2-oxoglutarate and

+
reduction 2–3 lmolÆmin
)1
Æmg
)1
) catalyzing
transformation of either 2-oxoglutarate and CoA (2 m
M
each) or NADH (8 m
M
). The same mixture was used as a
blank except that SOD (0.016 mgÆmL
)1
) was added.
Reactions were started with CoA or NADH. Under these
conditions initial rates of cytochrome c reduction in the
presence of SOD were no more than 20% of those obtained
without SOD.
Enzyme inactivation studies
Time-dependent changes in the activity of the enzyme
complex upon preincubation with its substrates and/or
products was studied during a 20-min preincubation
period in 0.1
M
potassium phosphate buffer, pH 7.0, at
the following final concentrations: enzyme complex,
3mgÆmL
)1
; 2-oxoglutarate, 2 m
M

radical intermediates by converting them to more stable
radicals. Nitrone (PBN, POBN, DMPO) and nitroso
(MNP) spin traps are known to react with short-lived
radicals, resulting in relatively long-lived nitroxide radical
adducts. Together with the characteristic properties of the
adducts formed, differential reactivity of spin traps to
radicals enables selective trapping and identification of the
original radical species.
The spin trap MNP is presumed to efficiently form
adducts with catalytic radical intermediates, as it is small
enough to reach enzyme active sites without major steric
hindrance. Aerobic incubation of MNP with the pyruvate or
5006 V. I. Bunik and C. Sievers (Eur. J. Biochem. 269) Ó FEBS 2002
2-oxoglutarate dehydrogenase complexes and their respect-
ive 2-oxo acid substrates, CoA and NAD
+
, led to formation
of MNP/
•
H, or t-butyl-hydronitroxide. This product of one-
electron reduction [25] was detected from the four line EPR
spectrum with a
N
¼ a
H
¼ 1.44 mT and its characteristic
change observed in 50% D
2
O due to t-butyl-deuteronitrox-
ide (a

doublet due to a hydrogen in the b-position to the nitrogen.
As in the reaction with MNP, the EPR signal persisted
after omitting NAD
+
from the reaction medium (Fig. 1,
spectrum 2). However, omitting either 2-oxo acid or CoA
(i.e. the components leading to the complex-bound dihydro-
lipoate) prevented the adduct formation.
Paramagnetic species were generated in an enzyme-
dependent manner both in the presence of the 2-oxo acid,
CoA (forward reaction) and when the 2-oxo acid dehy-
drogenase complexes catalyzed oxidation of NADH or
external dihydrolipoamide (backward reaction). In any
case, radicals were produced upon reduction of both the
E2-bound dihydrolipoate and E3. The particular contribu-
tion of these components to the production of radicals was
differentiated through their selective inactivation. N-Ethyl
maleimide modification of the lipoate residues of E2 led to
complete loss of the overall activity (Reactions 1–5) due to
E2 inactivation, while the E1 and E3 activities were fully
preserved. With this modified complex, no PBN adduct was
observed in the presence of 2-oxo acid and CoA, but it did
produce PBN adducts upon incubation with the E3
substrates, dihydrolipoate or NADH. Modification of the
tightly bound flavin cofactor of E3 with diphenyliodonium
chloride inactivated E3. This complex gave no PBN adduct
when incubated with either 2-oxo acid and CoA or NADH.
Thus, the E3-bound FAD was responsible for the formation
of radical species at the expense of either complex-bound or
free dihydrolipoate or NADH.

adducts. Probably, the thiyl radical of CoA was trapped
under these conditions. An indirect relationship between the
superoxide and PBN adducts is further supported by the
fact that the hyperfine splitting constants of the aerobic
PBN adducts formed in the forward (Table 1, N 1) and
backward (Table 1, N 2) reactions differed. Hence, secon-
dary reactions with the initially produced superoxide or its
PBN adduct must be invoked to explain formation of the
stable PBN adducts characterized by the EPR spectra
showninFig.1.
Direct detection of the superoxide anion radical
produced by the complexes
Production of superoxide anion radical by the complexes
was also examined by methods other than spin trapping.
Increased luminescence of lucigenin (bis-N-methylacridi-
nium) upon its reaction with superoxide is used for specific
detection of the latter in a number of biological systems [56].
Up to a 10-fold increase in the luminescence occured upon
incubation of the 2-oxoglutarate dehydrogenase complex
with 2-oxoglutarate and CoA in the presence of lucigenin.
However, lucigenin itself may increase formation of super-
oxide in the presence of enzymes that are capable of
catalyzing 1e
–
reduction of lucigenin directly [57], and E3
catalyzes 1e
–
reduction of various compounds [58–60].
Therefore to quantify production of superoxide radical by
Fig. 1. EPR spectra of PBN adducts recorded under equal settings after

identified radical
adducts
N
PBN E, 2-oxoglutarate
or pyruvate, CoA, O
2
Matched to thiyl radicals 1.56 ± 0.02 0.33 ± 0.01 Stable 1
E, NADH, O
2
Matched to alkyl radicals 1.6 ± 0.004 0.33 ± 0.01 Stable 2
E, 2-oxoglutarate or
pyruvate, CoA, anaerobic
Matched to thiyl radicals 1.57 ± 0.01 0.33 ± 0.01 Transient species,
stable at low substrate
3
E, NADH, anaerobic Matched to thiyl radicals 1.59 ± 0.01 0.33 ± 0.005 Transient species,
stable at low substrate
4
Reactive oxygen species
•
OH
•
OOH
1.53–1.56
1.48
0.27–0.295
0.275–0.295
s to min
Unstable (min)
77–79

9
10
Alkyl (RC
•
) Methyl
•
1.58–1.65 0.36–0.37 Stable 74, 80, 81 11
of linoleic acid 1.62–1.63 0.26–0.3 Stable 82, 83 12
Thiyl (RS
•
) of DTT
of glutathione
of cysteine
of protein Cys in 5
M
urea
1.56
1.56
1.56–1.57
1.6
0.32
0.32
0.34–0.35
0.36
min
min
min under oxidizing
conditions; hours without O
2
Stable

DMPO E, 2-oxoglutarate,
CoA, anaerobic
Matched to carbon-
centered radical
1.58 2.24 Stable 21
Thiyl (RS
•
) of cysteine
of glutathione
of N-acetylcysteine
of CoA
of lipoate
1.52–1.53
1.50–1.54
1.50
1.54
No adduct
1.70–1.73
1.62–1.63
1.68
1.62
min
transition metals with
SH excess catalyze decay
min
min
87
50, 87, 88
87
14

)1
Æmg
)1
. This corresponds to 0.3–0.4% of
the overall NAD
+
-reductase activity of the complex
(reactions 1–5).
Radical species and the catalysis-induced inactivation
of E1
Generation of ROS was documented in the current study
under conditions shown to induce catalysis-dependent
inactivation of the complexes [48,49]. Therefore we exam-
ined the possibility that the inactivation (Fig. 2A) was due
to the superoxide anion radical generated by the system. No
protection from the aerobic inactivation was observed in the
presence of SOD. Besides, the complexes were inactivated
by 2-oxo acid and CoA also under anaerobic conditions
(Fig. 2B). Thus, the enzyme inactivation was not caused by
the ROS produced. On the other hand, radical species were
deteced in the absence of oxygen too (Fig. 3). The spectrum
obtained under anaerobic conditions created with glucose
oxidase, glucose and catalase (Fig. 3, spectrum 2) showed a
weaker doublet at higher field, indicative of adduct decay
during the field sweep. As glucose is a known radical
scavenger [63] and in our experiments it indeed decreased
the signal of the PBN adducts already formed, a more
detailed study was performed under anaerobic conditions
created in a glove box.
Several properties of the anaerobic and aerobic PBN

reduction of the complex.
According to the backward catalytic process effected by
NADH (reactions 5 and 4), the protein-bound radical
species (Fig. 4B) could arise from either the E3 redox-active
disulfide and FAD or E2-bound lipoate residues. From
those, only the latter may show no nitroxide immobiliza-
tion, as the lipoyl-lysine side chains are mobile and protrude
from the complex core [1–4]. In particular, their essentially
free rotation was observed upon spin labeling of the lipoyl
Fig. 2. Inactivation of 2-oxoglutarate dehydrogenase complex in the
presence of 2-oxoglutarate and CoA under aerobic (A) and anaerobic (B)
conditions. Concentration of substrates: 0.15 m
M
(1), 1.5 m
M
(2).
Concentration of the complex used (9 mgÆmL
)1
 4.5 l
M
) corres-
ponds to  0.3 m
M
sites for substrate and/or reducing equivalents
(24E1 + 24E2 + 12E3-S
2
+12E3-FAD).
Fig. 3. Spectra of PBN adducts obtained upon anaerobic incubation
of 2-oxoglutarate dehydrogenase complex with 2-oxoglutarate and CoA.
1: Anaerobiosis created in glove box, reaction took place for 18 min in

hyperfine splitting constants (Table 1, N 3 and 4) charac-
teristic of the PBN-trapped thiyl radicals (Table 1, N
13,14,15) allow us to conclude that under anaerobic
conditions PBN traps radicals of the complex-bound
dihydrolipoate residues.
Detection of the E2-bound dihydrolipoate thiyl radical
correlated with the inactivation of the complexes by 2-oxo
acid plus CoA (Fig. 2B). In the absence of O
2
, both the
inactivation (Fig. 2B) and the EPR signal stability
decreased with increasing concentration of substrates, i.e.
upon full reduction of the catalytic centers. Dismutation of
the thiyl radicals and reduction of their PBN adducts within
the network of interacting lipoyl moieties of the E2 core
provides a good explanation for these phenomena.
Considering possible mechanisms of the inactivation of
the complex by the thiyl radical of dihydrolipoate, we took
into account that (a) the overall activity (Fig. 2) is decreased
due to the irreversible inactivation of E1 [49], and (b) the
thiyl radical of the lipoyl residue should efficiently interact
with the E1 catalytic intermediate, as the lipoyl-bearing
domain of E2 is designed for this interaction (reaction 2). In
this case, 1e
–
oxidation of the carbanion in the E1 active site
(a product of reaction 1) by highly electrophilic thiyl radical
of the dihydrolipoyl residue of E2 had to be expected. The
reaction was confirmed by anaerobic spin trapping with
DMPO. This spin trap is known to be unreactive towards

thioredoxin protection was obvious not only when assay-
ing the overall activity (reactions 1–5), but also upon
generation of the paramagnetic species. In the medium
without thioredoxin, the EPR signal intensity reached
saturation after the complex inactivation (10–15 min of
incubation with the substrates, Fig. 2A). When thioredoxin
was added, the initial rate of the EPR signal increase was
the same, but no saturation was obvious during half-an-
hour. Thus, as increased productivity of the complexes in
Fig. 5. DMPO spin trapping of anaerobic reaction medium containing
2-oxoglutarate dehydrogenase complex (9 mgÆmL
-1
), 2-oxoglutarate
and CoA (4 m
M
each).
Fig. 4. Localization of PBN adducts obtained upon incubation of E.coli
pyruvate dehydrogenase complex (27 mgÆmL
-1
)withNADH(0.4m
M
)
under aerobic (A) and anaerobic (B) conditions. 1: Before protein pre-
cipitation. 2: Protein fraction. 3: Non-protein fraction. Fractionation is
described in Materials and methods.
5010 V. I. Bunik and C. Sievers (Eur. J. Biochem. 269) Ó FEBS 2002
generation of radical species was observed in the presence
of thioredoxin.
Another argument for the presence of an oxidizing sulfur-
centered radical species under aerobic conditions is provided

inactivate E1. However, any combination of the substrates
and/or products providing concomitant presence of the E1
catalytic intermediate and complex-bound dihydrolipoate
(2-oxo acid + CoA; NADH + 2-oxoacid; dihydrolipo-
amide + succinyl-CoA) did lead to inactivation. In partic-
ular, the appearance of the ThDP adduct with the substrate
moiety during the complex-catalyzed succinyl-CoA hydro-
lysis [72] accounts for the inactivation by dihydrolipoamide
in the presence of succinyl-CoA (Table 2).
DISCUSSION
Generation of radical species during catalysis by 2-oxo acid
dehydrogenase multienzyme complexes has been documen-
ted in this work by EPR spectroscopy, ferricytochrome c
reduction and lucigenin fluorescence. The superoxide anion
radical is produced upon the E3-catalyzed 1e
–
oxidation of
the E2-bound dihydrolipoate intermediate. The thiyl radical
of the E2-bound dihydrolipoate formed in this reaction
causes the 1e
–
oxidation of the 2-oxo acid adduct with
ThDP through the site-directed action on E1. This results in
the carbon-centered radical fragment in the E1 active site
and the enzyme inactivation. The inactivation is prevented
by thioredoxin which is a known scavenger of thiyl radicals
[69].
The present work shows that production of radical
species by the 2-oxo acid dehydrogenase complexes under-
lies several phenomena of regulatory significance: (a)

)1
Æmg
)1
), or purified FAD-containing
monooxygenase (3–6 nmolÆmin
)1
Æmg
)1
) [62]. Inability of
the FAD-modified complex to generate paramagnetic
species rules out the direct oxidation of the accumulated
complex-bound dihydrolipoate intermediate as a source of
the superoxide and indicates that the integral complex
structure is required for the radical species production.
The results of the current study and the site-specific
reactivity of O
2
• –
also bear on consideration of the concept
of metabolons, i.e. intracellular compartmentalized func-
tional units. The regulatory potential of superoxide radical
production by the 2-oxoglutarate dehydrogenase complex
should greatly increase in the microenvironment of the citric
acid cycle metabolon. This implies close arrangement of the
2-oxoglutarate dehydrogenase complex and transition
metal-dependent enzymes, such as aconitase and fumarase,
both rapidly reacting with O
2
• –
. Aconitase has been shown

Succinyl-CoA No inactivation
NADH No inactivation
2-oxoglutarate, CoA 0.10 ± 0.01
Succinyl-CoA, dihydrolipoamide 0.03 ± 0.01
NADH + 2-oxoglutarate 0.04 ± 0.01
Ó FEBS 2002 Radicals upon oxidation of 2-oxo acids (Eur. J. Biochem. 269) 5011
PBN. However, the 1e
–
oxidation of the reduced complex
by oxygen implicates the redox equilibrium involving the
E3- and E2-bound thiyl radicals and the E3-bound flavin
semiquinone. Under these circumstances, appearance of a
strongly oxidizing thiyl radical of dihydrolipoate (E° of a
number of RS
•
/RS
–
or RS
•
/RSH couples are approximately
+0.75 or +1.33 V, respectively [71]) is supported by the
enzyme spectral changes, the SOD- and O
2
-insensitive
inactivation of the E1 catalytic intermediate (Fig. 2,
Table 2) and the protection by thioredoxin from such an
inactivation. Pro-oxidant action of thiyl radicals is avoided
in the presence of thioredoxin, because the free radical
species of thioredoxin, both disulfide and thiyl, are unreac-
tive [69]. In our system, migration of one electron between

bound dihydrolipoate radicals is less efficient than their
interaction with E1, and (b) the dihydrolipoate radical
intermediate survives long enough to be of regulatory
significance. By enabling E1 inactivation in response to the
absence of E3 substrate, the dihydrolipoate radical inter-
mediate transfers information from E3 to E1. As a result,
superoxide production by the complexes is restricted unless
thioredoxin is added. Thioredoxin modulates the self-
regulation of the complexes through abolition of the
deleterious action of the dihydrolipoate thiyl radicals on
E1. This provides an increased performance of the
complexes not only in the overall reaction, but also in
superoxide production.
Thus, the energy-providing oxidative decarboxylation of
2-oxoacids may influence mitochondrial metabolism also by
means of the pro-oxidant action of the dihydrolipoate
intermediate propagated through E3 (superoxide and thiyl
radicals production) and E1 (catalytic intermediate radical
production and inactivation). While formation of the
intrinsic thiyl radical is deleterious for the 2-oxo acid
oxidation, it also is an antioxidant defense mechanism,
preventing the superoxide production by the complexes.
External regulation of these processes by a cellular thiol-
disulfide oxidoreductase, thioredoxin, points to the link
between the complexes and thioredoxin-dependent
pathways in mitochondria. For instance, they may form
an antioxidant defense system, analogous to recently
discovered in mycobacteria where the 2-oxoglutarate dehy-
drogenase complex provides reducing equivalents to the
peroxiredoxin alkyl hydroperoxide reductase through a

drogenase complex of Escherichia coli: mechanism for rate
enhancement in a multimeric structure. Proc.Natl.Acad.Sci.USA
75, 5386–5390.
6. Akiyama, S.K. & Hammes, G.G. (1980) Elementary steps in the
reaction mechanism of the pyruvate dehydrogenase multienzyme
complex from Escherichia coli: kinetics of acetylation and deace-
tylation. Biochemistry 19, 4208–4213.
7. Waskiewicz, D.E. & Hammes, G.G. (1980) Elementary steps in
the reaction mechanism of the a-ketoglutarate dehydrogenase
multienzyme complex from Escherichia coli: kinetics of succiny-
lation and desuccinylation. Biochemistry 23, 3136–3143.
8. Reed, L.J. & Hackert, M.L. (1990) Structure-function relation-
ships in dihydrolipoamide acyltransferase. J. Biol. Chem. 265,
8971–8974.
9. Massey, V., Gibson, Q.H. & Veeger, C. (1960) Intermediates in the
catalytic action of lipoyl dehydrogenase (diaphorase) Biochem. J.
77, 341–351.
10. Massey, V. (1963) Lipoyl dehydrogenase. Enzymes 7, 275–306.
11. Searls, R.L., Peters, J.M. & Sanadi, D.R. (1961) a-Ketoglutaric
dehydrogenase. X. On the mechanism of dihydrolipoyl dehy-
drogenase reaction. J. Biol. Chem. 236, 2317–2322.
12. Matthews, R.G., Ballou, D.P., Thorpe, C. & Williams, C.H. Jr
(1977) Ion pair formation in pig heart lipoamide dehydrogenase.
J. Biol. Chem. 252, 3199–3207.
13. Templeton, D.M., Hollebone, B.R. & Tsai, C.S. (1980) Magnetic
circular dichroism studies on the active-site flavin of lipoamide
dehydrogenase. Biochemistry 19, 3969–3873.
14. Docampo, R., Moreno, S.J. & Mason, R.P. (1987) Free
radical intermediates in the reaction of pyruvate: ferredoxin
5012 V. I. Bunik and C. Sievers (Eur. J. Biochem. 269) Ó FEBS 2002

production of superoxide anion radicals in the reaction of reduced
flavins and flavoproteins with molecular oxygen. Biochem.
Biophys. Res. Communs. 36, 891–898.
23. Ballou, D., Palmer, G. & Massey, V. (1969) Direct demonstration
of superoxide anion production during the oxidation of reduced
flavin and of its catalytic decomposition by erythrocuprein.
Biochem. Biophys. Res. Communs. 36, 898–904.
24. Kunz, W.S. & Kunz, W. (1985) Contribution of different enzymes
to flavoprotein fluorescence of isolated rat liver mitochondria.
Biochim. Biophys. Acta 841, 237–246.
25. Khan, A.U. & Wilson, T. (1995) Reactive oxygen species as cel-
lular messengers. Chem. Biol. 2, 437–445.
26. Skulachev, V.P. (2000) Mithochondria in the programmed death
phenomena; a principle of biology. ÔIt is better to die than to be
wrongÕ. JUBMB Life 49, 365–373.
27. Sastre, J., Pallardo, V. & Vina, J. (2000) Mitochondrial oxidative
stress plays a key role in aging and apoptosis. JUBMB Life. 49,
427–435.
28. Cabiscol, E., Piulats, E., Echave, P., Herrero, E. & Ros, J. (2000)
Oxidative stress promotes specific protein damage in Sacharomices
cerevisiae. J. Biol. Chem. 275, 27393–27398.
29. Humphries, K.M. & Szweda, L.I. (1998) Selective inactivation of
a-ketoglutarate dehydrogenase and pyruvate dehydrogenase:
reaction of lipoic acid with 2-hydroxy-2-nonenal. Biochemistry 37,
15835–15841.
30. Rokutan, K., Kawai, K. & Asada, K. (1987) Inactivation of
2-oxoglutarate dehydrogenase in rat liver mitochondrial by its
substrate and T-butyl hydroperoxide. J. Biochem. 101, 415–422.
31. Millar, H.A. & Leaver, C.J. (2000) The cytotoxic lipid peroxida-
tion product, 4-hydroxy-2-nonenal, specifically inhibits dec-

Neurol. 35, 204–210.
39. Butterworth, R.F., Kril, J.J. & Harper, C.G. (1993) Thiamine-
dependent enzyme changes in the brains of alcoholics: Relation-
ship to the Wernike–Korsakoff syndrome alcohol. Clin.Exp.Res.
71, 1084–1088.
40. Hackert, M.L., Oliver, R.M. & Reed, L.J. (1983) A computer
model analysis of the active-site coupling mechanism in the pyru-
vate dehydrogenase complex of Escherichia coli. Proc. Natl Acad.
Sci. USA 80, 2907–2911.
41. Collins, J.H. & Reed, L.J. (1977) Acyl Group and electron pair
relay system: a network of interacting lipoyl moieties in the
pyruvate and a-ketoglutarate dehydrogenase complexes from
Escherichia coli. Proc. Natl. Acad. Sci. USA 74, 4223–4227.
42.Guest,J.R.,Ali,S.T.,Artymiuk,P.,Ford,G.C.,Green,J.&
Russel, G.C. (1990) Site-directed mutagenesis of dihy-
drolipoamide acetyltransferase and post-translational modifica-
tion of its lipoyl domains. In Biochemistry and Physiology of
Thiamin Diphosphate Enzymes (Bisswanger, H. & Ullrich, J., eds),
pp. 176–193. Chemie, Weinheim.
43. Scott, B.C., Aruoma, O.I., Evans, P.J., O’Neill, C., Van der Vliet,
A., Cross, C.E., Tritschler, H. & Halliwell, B. (1994) Lipoic and
dihydrolipoic acids as antioxidants. A critical evaluation. Free
Rad. Res. 20, 119–133.
44. Packer, L., Witt, E.H. & Tritschler, H.J. (1995) Alpha-lipoic acid
as a biological antioxidant. Free Rad Biol. Med. 19, 227–250.
45. Biewenga, G.Ph, Haenen, G.R.M.M. & Bast, A. (1997) The
pharmacology of the antioxidant lipoic acid Gen. Pharmacol. 29,
315–331.
46. Bunik, V. & Follmann, F. (1993) Thioredoxin reduction depen-
dent on a-keto acid oxidation by a-keto acid dehydrogenase

residues in the pyruvate dehydrogenase multienzyme complex of
Escherichia coli. Biochem. J. 187, 393–401.
55. Kalyanaraman, B. & Mason, R.P. (1979) The reduction of
nitroso-spin traps in chemical and biological systems. A cau-
tionary note. Tetrahedron Lett. 50, 4809–4812.
56. Li, Y., Stansbury, K.H., Zhu, H., & Trush, M.A. (1999)
Biochemical characterization of lucigenin (bis-N-methylacridi-
nium) as a chemiluminescent probe for detecting intramito-
chondrial superoxide anion radical production. Biochem. Biophys.
Res. Commun. 262, 80–87.
57. Skatchkov, M.P., Sperling, D., Hink, U., Aenggard, E. &
Muenzel, T. (1998) Quantificantion of superoxide radical forma-
tion in intact vascular tissue using a Cypridina luciferin analog as
an alternative to lucigenin. Biochem. Biophys. Res. Commun. 248,
383–386.
58. Leskovac, V., Svircevic, J., Trivic, S., Popovic, M. & Radulovic,
M. (1989) Reduction of aryl-nitroso compounds by pyridine and
flavin coenzymes. Int. J.Biochem. 21, 825–834.
59. Vienozinskis, J., Butkus, A., Cenas, N. & Kulys, J. (1990) The
mechanism of the quinone reductase reaction of pig heart lipoa-
mide dehydrogenase. Biochem. J. 269, 101–105.
60. Sreider, C.M., Grinblat, L. & Stoppani, A.O.M. (1992) Reduction
of nitrofuran compounds by heart lipoamide dehydrogenase: role
of flavin and the reactive disulfide groups. Biochem. Internat. 28,
323–334.
61. McCord, J.M. & Fridovich, I. (1968) The reduction of cytochrome
c by milk xantine oxidase. J. Biol. Chem. 243, 5753–5760.
62.Rosen,G.M.,Finkelstein,E.&Rauckman,E.J.(1982)A
method for the detection of superoxide in biological systems. Arch.
Biochem. Biophys. 215, 367–378.

ymol. 186, 168–180.
72. Frey, P.A., Flournoy, D.S., Gruys, K. & Yang, Y S. (1989)
Intermediates in reductive transacetylation catalyzed by pyruvate
dehydrogenase complex. Ann. NY Acad. Sci. 573, 21–35.
73. Konstantinov, A.A., Peskin, A.V., Popova, E.Yu, Khomutov,
G.B. & Ruuge, E.K. (1987) Superoxide generation by the
respiratory chain of tumor mitochondria. Biochim. Biophys. Acta
894, 1–10.
74. Vasquez-Vivar, J., Kalyanaraman, B. & Kennedy, M.C. (2000)
Mitochindrial aconitase is a source of hydroxyl radical. An elec-
tron spin resonance investigation. J. Biol. Chem. 275, 14064–
14069.
75. Bonomi, F., Cerioli, A. & Pagani, S. (1989) Molecular aspects of
the removal of ferritin-bound iron by
DL
-dihydrolipoate. Biochim.
Biophys. Acta V. 994, 180–186.
76.Bryk,R.,Lima,C.D.,Erdjument-Bromage,H.,Tempst,P.&
Nathan, C. (2002) Metabolic enzymes of mycobacteria linked to
antioxidant defense by a thioredoxn-like protein. Science 295,
1073–1077.
77. Kotake, Y. & Janzen, E.G. (1991) Decay and fate of the hydroxyl
radical adduct of a-phenyl-N-tert-butylnitrone in aqueous media.
J. Am. Chem. Soc. 113, 9503–9506.
78. Harbour, J.R., Chow, V. & Bolton, J.R. (1974) An electron spin
resonance study of the spin adducts of OH and HO
2
radicals with
nitrones in the ultraviolet photolysis of aqueous hydrogen per-
oxide solutions. Can. J. Chem. 52, 3549–3553.

nitroxides. Anal. Chem. 71, 715–721.
87. Ross, D., Albano, E., Nilsson, U. & Moldeus, P. (1984) Thiyl
radicals formation during peroxidase-catalyzed metabolism of
acetaminophen in the presence of thiols. Biochem. Biophys. Res.
Commun. 125, 109–115.
5014 V. I. Bunik and C. Sievers (Eur. J. Biochem. 269) Ó FEBS 2002
88. Mason, R.P. & Ramakrishna Rao, D.N. (1990) Thyil free radical
metabolites of thiol drugs, glutathione, and proteins. Methods
Enzymol. 186, 318–329.
89. Buettner, G.R. (1984) Thiyl free radical production with hem-
atoporphyrin derivative, cysteine and light: a spin-trapping study.
FEBS Lett. 177, 295–299.
90. Saez,G.,Thornalley,P.J.,Hill,H.A.O.,Hems,R.&Bannister,
J.V. (1982) The production of free radicals during the autoxidation
of cysteine and their effect on isolated rat hepatocytes. Biochim.
Biophys. Acta 719, 24–31.
91. Sankarapandi, S. & Zweier, J.L. (1999) Evidence against the
generation of free hydroxyl radicals from the interaction of cop-
per, zinc-superoxide dismutase and hydrogen peroxide. J. Biol.
Chem., 274, 34576–34583.
Ó FEBS 2002 Radicals upon oxidation of 2-oxo acids (Eur. J. Biochem. 269) 5015