Báo cáo khoa học: 2-Oxo acid dehydrogenase complexes in redox regulation Role of the lipoate residues and thioredoxin pot

MINIREVIEW
2-Oxo acid dehydrogenase complexes in redox regulation
Role of the lipoate residues and thioredoxin
Victoria I. Bunik
A.N.Belozersky Institute of Physico-Chemical Biology, Moscow State University, Russia
A number of cellular systems cooperate in redox regulation,
providing metabolic responses according to changes in the
oxidation (or reduction) of the redox active components of
a cell. Key systems of central metabolism, such as the 2-oxo
acid dehydrogenase complexes, are important participants
in redox regulation, because their function is controlled by
the NADH/NAD
+
ratio and the complex-bound dihydro-
lipoate/lipoate ratio. Redox state of the complex-bound
lipoate is an indicator of the availability of the reaction
substrates (2-oxo acid, CoA and NAD
+
) and thiol-disulfide
status of the medium. Accumulation of the dihydrolipoate
intermediate causes inactivation of the first enzyme of the
complexes. With the mammalian pyruvate dehydrogenase,
the phosphorylation system is involved in the lipoate-
dependent regulation, whereas mammalian 2-oxoglutarate
dehydrogenase exhibits a higher sensitivity to direct regula-
tion by the complex-bound dihydrolipoate/lipoate and
external SH/S-S, including mitochondrial thioredoxin.
Thioredoxin efficiently protects the complexes from self-
inactivation during catalysis at low NAD
+
. As a result,

2-oxoglutarate dehydrogenase complex may provide succi-
nyl-CoA for phosphorylation of GDP and ADP under
conditions of restricted NAD
+
availability. This may be
essential upon accumulation of NADH and exhaustion of
the pyridine nucleotide pool. Concomitantly, thioredoxin
stimulates the complex-bound dihydrolipoate-dependent
production of reactive oxygen species. It is suggested that
this side-effect of the 2-oxo acid oxidation at low NAD
+
in vivo would be overcome by cooperation of mitochondrial
thioredoxin and the thioredoxin-dependent peroxidase,
SP-22.
Keywords: dihydrolipoate; 2-oxo acid dehydrogenase
complex; radical species; redox state; thioredoxin.
Introduction
Regulation of metabolism dependent on the cellular redox
state has attracted increasing attention [1–5]. The redox
state is characterized by the degree of oxidation or reduction
of various redox-active species of a cell. Among these
species, pyridine nucleotides and thiol/disulfide compounds
are of special significance, as they interconnect many
enzymes of the multifaceted metabolic network. On the
one hand, the steady-state ratios of NAD(P)H/NAD(P)
+
and SH/S-S mediate the redox regulation through direct
effects on proteins. Activities of many enzymes depend on
the redox state of the pyridine nucleotide pools, while
proteins with essential SH/S-S groups can be regulated by

post-translational modification involving cellular thiols and
disulfides [1,5–7]. On the other hand, the NAD(P)H/
NAD(P)
+
and SH/S-S ratios are intimately related to the
cellular level of ROS. Reduced pyridine nucleotides and
thiols participate both in ROS formation and degradation.
The former process is effected by different NAD(P)H
oxidases [8] and upon thiol oxidation [9,10], while the
glutathione- and thioredoxin-dependent peroxidase reac-
tions use NADPH and thiols to scavenge hydrogen
peroxide and limit formation of radical species [11]. At
low concentrations, ROS are essential participants of the
cellular redox regulation [3,11,12]. Their extremely high
reactivity allows for the fast local modification of proteins.
This can provide transient oxidative modifications against
the reducing potential of the medium, e.g. formation of
a protein disulfide bond in the reducing cytoplasm [13].
However, the high reactivity also leads to cell destruction if
cellular capacity to scavenge ROS is compromised. This
happens under pathological conditions where NAD(P)H/
NAD(P)
+
and SH/S-S ratios are decreased [14–17].
While experiments with intact cells enable us to assess the
net effects of redox perturbations that reflect integrated
metabolic responses, dissecting the mechanisms of these
overall responses requires investigation of the separate
components of the metabolic network. Among cellular
systems, the 2-oxo acid dehydrogenase multienzyme com-

plexes occupy key positions for redox regulation. In the
overall process (see reactions 1–5 in scheme below) involving
sequential action of 2-oxo acid dehydrogenase (E1),
dihydrolipoamide acyltransferase (E2) and dihydrolipo-
amide dehydrogenase (E3), they split a carbon-carbon
bond of the 2-oxo acid preserving its energy in acyl-CoA
Correspondence to V. Bunik, A.N.Belozersky Institute of
Physico-Chemical Biology, Moscow State University,
Moscow 119899, Russia.
Fax: + 7 095 939 31 81, Tel.: + 7 095 939 14 56,
E-mail:
Abbreviations: E1, 2-oxo acid dehydrogenase; E2, dihydrolipoamide
acyltransferase; E3, dihydrolipoamide dehydrogenase; EPR,
electron paramagnetic resonance; ROS, reactive oxygen species.
(Received 23 July 2002, revised 10 December 2002,
accepted 11 December 2002)
Eur. J. Biochem. 270, 1036–1042 (2003) Ó FEBS 2003 doi:10.1046/j.1432-1033.2003.03470.x
and NADH, where R defines pyruvate, 2-oxoglutarate or
the 2-oxo acids with branched carbon chain:
Thus, the catalytic action of the 2-oxo acid dehydro-
genase complexes directly influences the NADH/NAD
+
ratio and involves the important biological SH/S-S com-
pounds, lipoic acid and CoA, with one of them (lipoate)
being covalently bound to the complexes.
Sensitivity of the 2-oxo acid dehydrogenase complexes to
NADH/NAD
+
has long been recognized as a mechanism
of feedback control [18,19]. Being operative in vivo [20], it

increases fluxes through pyruvate dehydrogenase complex
under more oxidizing conditions [2]. However, the SH/S-S-
dependent regulation of the complexes, in particular, by the
complex-bound lipoate/dihydrolipoate ratio, has received
little attention, although some experiments suggested the
interaction of the complex-bound dihydrolipoate with
externaldisulfides[21,22].Atthesametime,thereare
numerous studies of the antioxidant properties of free
lipoate [23–25]. Free dihydrolipoate efficiently reduces
transcription factors [26] and thioredoxin [27], which are
well-known components of the redox regulation [5,11,28].
Because cellular lipoate/dihydrolipoate is mostly localized
to the 2-oxo acid dehydrogenase complexes, the interplay
between the complexes and other participants of redox
signaling is of great interest.
An essential feature of the complex-bound lipoate/
dihydrolipoate couple is that its redox state is linked to the
irreversible reaction of the 2-oxo acid oxidation. Because
of this, the lipoate redox state in vivo is defined by the
steady-state concentrations of the overall reaction sub-
strates (2-oxo acid, CoA and NAD
+
) rather than
thermodynamic equilibrium with cellular thiols. Another
essential feature of the lipoate residues within the 2-oxo
acid dehydrogenase complexes arises from the ÔcrowdingÕ
effect. Confinement of the complex-bound lipoate within
the volume of the enzyme complexes may result in unusual
properties of the compound, when compared to the same
quantity of the lipoate molecules distributed in bulk

solution. In particular, neighbouring sulfur greatly increa-
ses the stability of the thiyl radical in dithiols compared to
monothiols [29]. Additional stabilization of the lipoate
thiyl radicals may be expected within the network of the
interacting complex-bound lipoate residues. Thus, the
complexes provide an excellent opportunity to study in vitro
the consequences of cellular compartmentalization of
biologically active SH/S-S.
Taking into account multiple pharmacological effects of
free lipoate [23–25], we suggested that the lipoate clustering
within the 2-oxo acid dehydrogenase complexes may be
significant not only for the typical enzymatic catalysis, but
also for cellular redox regulation, including cellular protec-
tion against oxidative damage. Our idea that the complex-
bound lipoate possesses a function beyond its role as
a catalytic intermediate in the oxidative decarboxylation is
in good agreement with certain data about the complexes,
which have not previously been given an adequate explan-
ation. Even a limited number of organisms from which the
2-oxo acid dehydrogenase complexes have been isolated
have revealed structural variations of these systems that go
beyond those essential for catalytic performance. The
variations are linked to different lipoate content in the
complexes, including different degrees of oligomerization,
different stoichiometries of catalytic components (in parti-
cular, lipoyl moieties) and different localization of the
lipoate residues. They may be incorporated not only in the
established lipoate holder, E2, but also in other components
[30,31]. Depending on the source and presence of substrates,
the mostly investigated ÔclassicÕ complexes (i.e. those

containing lipoate residues in E2 only) are built around
oligomeric cores of 3, 24 or 60 E2 subunits. Different types
of E2 may bear up to three lipoate residues, which is
genetically determined [30,32,33]. Surprisingly, more than
half of the lipoyl moieties of the E2 oligomer [34–36], or two
of the three lipoyl-bearing domains of E2 [37,38], may be
removed without significant change in the overall activity
in vitro. Yet microorganisms possessing pyruvate dehydro-
genase complex with a decreased number of lipoyl domains
are at a physiological disadvantage. They exhibit decreased
growth rates and are eventually Ôwashed-outÕ from the
mixed population containing the mutant and wild type cells
[38,39]. The physiological behaviour, however, is main-
tained even with a reduced number of lipoyl groups as long
as the ÔreachÕ of the lipoyl moieties is not decreased. That is,
the mutant strain possessing E2 bearing the three lipoyl
domains with only the outermost one lipoylated (the two
inner lipoyl domains in this case do not contain the lipoyl
group) behaved identically to the wild type under the
conditions employed [38]. Because such a mutant complex
catalyzes pyruvate oxidation even 25% less efficiently than
the complexes which are unable to provide the normal
growth rates (with E2 containing one or two fully lipoylated
domains) [38], the physiological advantage of the E2 with
the three lipoyl domains cannot be ascribed entirely to the
catalytic role of these domains. Rather, the advantage
appears to depend on the ability of the lipoyl group to
protrude from the inner core of the complex, indicative of
the biological significance of the complex-bound lipoate
interaction with the surrounding medium.

This paper reviews experimental evidence for the involve-
ment of the complex-bound lipoate in such ÔparacatalyticÕ
reactions, i.e. those where the complex-bound lipoate
escapes the catalytic route (reactions 1–5). The reactions
underlie the SH/S-S-dependent regulation of the 2-oxo acid
dehydrogenase complexes on different levels. The basic level
corresponds to self-regulation of the complexes by the
complex-bound lipoate/dihydrolipoate ratio. Involvement
Ó FEBS 2003 Redox regulation of 2-oxo acid dehydrogenation (Eur. J. Biochem. 270) 1037
of external components in the lipoate-dependent reactions
extends this regulation to a higher level. For instance,
production of ROS by the complexes and interchange of the
complex-bound dihydrolipoate/lipoate with external thiols
and disulfides, including thioredoxin, may be important for
ROS-dependent signaling. Thus, the redox state of the
complex-bound lipoate creates a sensitive link between
the 2-oxo acid dehydrogenase reaction and surrounding
medium.
Self-regulation of the complexes
by the redox state of the lipoate residues
As follows from the scheme of the overall 2-oxo acid
dehydrogenase reaction (reactions 1–5), the steady-state
ratio of the complex-bound lipoate/dihydrolipoate is a func-
tion of (a) concentrations of the reaction substrates and
products, (b) kinetic properties of the component enzymes,
E1, E2 and E3, and (c) their stoichiometry and interactions
within the complex. During catalysis in the physiological
direction, the complex-bound lipoate is reduced to dihydro-
lipoate by the 2-oxo acid and CoA (reactions 1–3) and the
dihydrolipoate is reoxidized by NAD

+
in a FAD-dependent
process (reactions 4,5). The lipoate may be also reduced in
the backward reactions (5 and 4), upon preincubation with
NADH. In the mitochondrial 2-oxoglutarate dehydrogenase
complex, this induces strong cooperativity among the active
sites of its first component, 2-oxoglutarate dehydrogenase,
upon 2-oxoglutarate binding, and complicates the kinetic
dependence of the reaction rate on 2-oxoglutarate [40].
Isolated from the complex, the 2-oxoglutarate dehydro-
genase component did not show such changes in response
to NADH. However, the changes were observed after the
enzyme reduction with dihydrolipoate. Similar concentra-
tion of other cellular thiols, such as glutathione, cysteine or
CoA, were ineffective [40]. Thus, the lipoate residues of the
complex mediate the regulation of its first component by the
last product of the overall reaction, NADH.
Reduction of the lipoate residues of the complexes in
the forward direction, i.e. by 2-oxo acid and CoA
(reactions 1–3), when the following reoxidation by
NAD
+
(reaction 5) is restricted, is accompanied by an
irreversible inactivation of E1 [41]. The inactivation is
observed both in the presence and absence of O
2
.When
the complex-bound lipoate was reduced under anaerobic
conditions, the complex-bound thyil radical and a radical
fragment of 2-oxo acid were detected in spin trapping

experiments with a-phenyl-N-tert-butylnitrone and 5,5¢-
dimethyl-1-pyrroline-N-oxide, respectively [42]. Thus, the
E1 inactivation occurs upon 1e
–
reduction of the thiyl
radical of the complex-bound dihydrolipoate by the E1
catalytic intermediate (E1*S):
The resulting substrate-derived radical fragment (S
Æ
)
likely causes the observed E1 inactivation due to a site-
directed modification. Efficiency of reaction 6 is provided by
the protein–protein interactions evolved to enable the
catalytic 2e
–
reduction of the lipoate by E1*S (reaction 2).
In the absence of O
2
, the dihydrolipoate thiyl radical is
transiently formed upon equilibration of the complex
redox centers. In the presence of O
2
, the E3-bound FAD
catalyzes 1e
–
oxidation of the complex-bound dihydrolipo-
ate by oxygen, resulting in superoxide anion radical
production [42]. The thiyl radical of the complex-bound
dihydrolipoate is an intermediate of this side reaction
(Fig. 1). The superoxide production by the complexes is

competitive with the NAD
+
reduction. Under conditions
where less NAD
+
is available, more superoxide is
produced, and this leads to a higher steady-state concen-
tration of complex-bound thiyl radicals and a concomi-
tantly greater extent of enzyme inactivation by the 2-oxo
acid plus CoA. Saturation by NAD
+
protects from the
1e
–
oxidation of the dihydrolipoate intermediate by oxygen
and prevents inactivation [41]. Resistance of the overall
activity to the superoxide anion radical produced is
documented by the fact that superoxide dismutase does
not prevent the inactivation. This is in good accord with
the independence of the inactivation on the presence of
oxygen.
The dihydrolipoate-mediated inactivation of E1 at low
NAD
+
concentrations is more pronounced in mammalian
than bacterial complexes [43]. In contrast, inhibition of E3
by over-reduction with NADH is less efficient in mamma-
lian complexes [44]. Thus, the E3 inhibition seems to be the
main response of bacterial complexes to NADH accumu-
lation. The mammalian complexes develop the E1-directed

mechanisms of NADH- and dihydrolipoate-dependent
regulation. It is the initial stage of the substrate transfor-
mation which is then affected.
The 2-oxoglutarate dehydrogenase complex is more
sensitive to the 2-oxo acid, CoA-induced inactivation than
the pyruvate dehydrogenase complex [43]. This agrees with
the difference in the regulation of the two complexes by
Fig. 1. Production of superoxide anion radical by the 2-oxo acid dehy-
drogenase complexes. Oxidation of the complex-bound dihydrolipoate
by oxygen is catalyzed by the E3-bound FAD. Superoxide anion
radical is detected in the reaction medium by appearance of the EPR
signal corresponding to its reaction with the spin trap a-phenyl-N-tert-
butylnitrone. Appearance of the EPR signal is blocked either by the
modification of the complex-bound FAD or by addition of superoxide
dismutase.
1038 V. I. Bunik (Eur. J. Biochem. 270) Ó FEBS 2003
phosphorylation/dephosphorylation. The latter mechanism
controls the function of eukaryotic pyruvate dehydro-
genase, whereas 2-oxoglutarate dehydrogenase is not phos-
phorylated. Remarkably, the efficiency of the pyruvate
dehydrogenase phosphorylation depends on the state of the
complex-bound dihydrolipoate. Thus, the redox regulation
of the eukaryotic pyruvate dehydrogenase is mediated by
the phosphorylation/dephosphorylation system, which thus
becomes the main transducer of multiple metabolic signals.
Regarding the mammalian 2-oxoglutarate dehydrogenase
complex, the 2-oxo acid, CoA-dependent inactivation
through the complex-bound dihydrolipoate intermediate
appears to be the biologically relevant mechanism of redox
regulation. The concentrations of 2-oxoglutarate and CoA

determined in mitochondria [45] are saturating for the
complex, while NADH/NAD
+
ratio varies depending on
the metabolic state. Moreover, NAD
+
is a substrate of
many mitochondrial enzymes, with the competition between
them further reducing the effective NAD
+
concentration
available for 2-oxoglutarate oxidation. Hence, estimation of
the substrate ratio existing in vivo shows that it is in the
range where the self-inactivation of the 2-oxoglutarate
dehydrogenase complex upon accumulation of the dihydro-
lipoate intermediate may occur.
As a result, the complex-bound lipoate allows the starting
component of the complexes, E1, to respond to the state of
the mitochondrial NAD
+
and NADH pool. The E1
activity is regulated both upon accumulation of NADH
and decrease of NAD
+
. The E1 inactivation at low NAD
+
concentration prevents the side production of dihydrolipo-
ate-dependent reactive oxygen species (Fig. 1) at the expense
of the 2-oxo acid oxidation. Because a decrease in
concentration of mitochondrial pyridine nucleotides induces

antioxidant defense mechanisms [15,16], inactivation of the
2-oxo acid dehydrogenases under these conditions may be a
part of the integrated response. It also may explain the
reduction of the 2-oxoglutarate dehydrogenase complex
activity observed under different pathological states [46].
Thiol-disulfide exchange between
the complex-bound lipoate and external
thiols/disulfides
In studies of mitochondria, disulfides inhibited mitochond-
rial respiration at the level of the 2-oxoglutarate dehydro-
genase reaction [21,22]. Investigation of the purified
2-oxoglutarate dehydrogenase complex confirmed the inhi-
bition of the overall reaction by the low molecular mass
disulfides [41]. The data pointed to the exchange of redox
equivalents between the complexes and the medium,
involving the dihydrolipoate intermediate. Such an
exchange also enabled thiols or disulfides to protect the
complexes from the inactivation at low levels of NAD
+
[43,47]. With free disulfides which are substrates for E3
(R-lipoate), the flow of reducing equivalents from 2-oxo
acids to the disulfides was catalyzed by E3 [48,49]. Because
dihydrolipoate is an efficient reductant of thioredoxin [27]
which may further direct reducing equivalents to different
processes [28], the 2-oxo acid dehydrogenase reaction
coupled to free lipoate reduction in the presence of thio-
redoxin may be a source of reducing equivalents for not
only NADH-dependent, but also thioredoxin-dependent
pathways. For instance, the reaction provides reduction of
disulfides in proteins such as insulin and thioredoxin

reductase [48,49]. Recently, an antioxidant defense system
in mycobacteria was discovered where the 2-oxoglutarate
dehydrogenase complex provides reducing equivalents to
the peroxiredoxin alkyl hydroperoxide reductase through
a thioredoxin-like protein [50].
Discovery of a specific mitochondrial thioredoxin with
unknown protein targets [51] stimulated our interest in the
potential interplay between the 2-oxo acid dehydrogenase
complexes and thioredoxin via the complex-bound lipoate.
Unraveling an in vivo function of a thioredoxin species is
complicated by the high chemical reactivity of its dithiol/
disulfide group, as it allows thioredoxin to participate in
a number of redox processes in vitro. Study of cross-
reactivity of thioredoxins and potential target proteins
from different species helps to solve this problem through
revealing specific protein–protein interactions promoting
chemical reactions. In particular, mitochondrial thio-
redoxin rather inefficiently regulates the enzymes which
are known to depend on the thioredoxin action [52]. In
contrast, it efficiently protects the 2-oxo acid dehydro-
genase complexes from the 2-oxo acid, CoA-dependent
inactivation [47,52]. Studies using four types of the 2-oxo
acid dehydrogenase complexes and 11 thioredoxin species
support the biological relevance of this protection [43,47].
Mitochondrial complexes are much more sensitive to the
thioredoxin regulation than their bacterial counterparts.
This is due to a greater sensitivity of the mitochondrial
complexes to the 2-oxo acid, CoA-induced inactivation, as
the thioredoxin effect is related to alleviation of this
inactivation. On the other hand, among 11 thioredoxin

species with comparable activity in the nonspecific insulin
reduction test, mitochondrial thioredoxin is by far the
most effective in protecting the complexes. While some of
thioredoxins are inactive or even decrease the complex
activity, mitochondrial thioredoxin is protective down to
10
)7
M
concentrations. Correlation of the thioredoxin
effects and protein structures revealed the following
structural determinants of the specific action of mito-
chondrial thioredoxin on the complexes [47]: (a) active site
disulfide/dithiol group and the residues modulating its
properties, (b) the interaction between the a3/3
10
and a1
helices and the length of the a1 helix and (c) the three
charged residues on the thioredoxin surface opposite to
the active site, which significantly influence polarization of
the molecule. Experimentally observed effects of different
thioredoxins on the complexes (increase or decrease in the
complex activity, or none) correlate with the dipole
direction, while the effective thioredoxin concentrations
correlate with the dipole magnitude. It is known that
steering effects of the long-range interactions between the
electrostatic dipoles increase the number of effective
collisions, i.e. collisions which may be stabilized by
short-range interactions [53]. The observed correlation
between polarization of the thioredoxin molecule and
efficiency of its protection of the 2-oxo acid dehydroge-

nase complexes points to long-range interactions as
the basis for the effect of thioredoxin on the activity of
the complex. This relationship suggests coevolution of the
interacting proteins, which would not be possible if the
interactions were not relevant in vivo.
Ó FEBS 2003 Redox regulation of 2-oxo acid dehydrogenation (Eur. J. Biochem. 270) 1039
Thioredoxin protection from the 2-oxo acid, CoA-
induced inactivation of the dehydrogenase complexes, and
the recently published data on the high stability of the
thioredoxin thyil radical, which allows thioredoxin to
prevent the pro-oxidant action of the radical [54], support
the proposed mechanism of the inactivation (reaction 6).
Catalysing the dismutation of the dihydrolipoate thyil
radicals (Fig. 2), thioredoxin prevents their adverse action
upon E1. The main component of cellular thiol buffer,
glutathione, also protects the complexes in vitro [41].
However, unlike thioredoxin, low molecular mass thiols
do not specifically bind to the complexes and their thiyl
radicals are known to possess pro-oxidant action [10,55,56].
Hence, regarding the overall mitochondrial metabolism,
glutathione cannot be an efficient scavenger of the complex-
bound thiyl radicals of dihydrolipoate.
Protected by thioredoxin, the 2-oxoglutarate dehydro-
genase complex can produce energy at low NAD
+
not only
in the form of NADH, but also in the form of a macroergic
compound, succinyl-CoA. The latter supports the only
reaction of substrate phosphorylation in the Krebs cycle,
catalyzed by succinyl thiokinase. This may be especially

important in cases where leakage of pyridine nucleotides or
accumulation of NADH occurs due to disturbances in the
respiratory chain. However, switching off the self-regulation
of the complexes by the dihydrolipoate thiyl radical,
thioredoxin also stimulates the side reaction of the super-
oxide anion radical production by the complexes (Fig. 1). In
this regard, it is worth noting that mitochondrial thiore-
doxin is a substrate of mitochondrial thioredoxin peroxi-
dase, SP-22 [57]. Reduced by the complex-bound
dihydrolipoate and coupled to SP-22, thioredoxin would
be able to scavenge hydrogen peroxide, which is formed
after dismutation of the superoxide anion radical produced
by the complexes as shown below:
Thus, thioredoxin interaction with the 2-oxo acid dehy-
drogenase complexes under conditions of an increased
steady-state concentration of dihydrolipoate may provide
a dual positive effect: relief of the pro-oxidant action of
dihydrolipoate on E1 and scavenging of ROS produced by
E3. As a result, cooperation of the 2-oxo acid dehydro-
genase complexes, thioredoxin and SP-22 (reaction 7)
enables oxidation of 2-oxo acids under increased concen-
tration of the dihydrolipoate intermediate without accumu-
lation of ROS.
The antioxidant action of mitochondrial thioredoxin
upon the 2-oxoglutarate dehydrogenase may be involved in
the thioredoxin antiapoptotic action when cells are treated
with tert-butyl hydroperoxide [58]. Selective targeting of the
2-oxoglutarate dehydrogenase complex under oxidative
stress [59] and inactivation of the 2-oxoglutarate dehydro-
genase by tert-butyl hydroperoxide [60] favour this inter-

pretation.
Concluding remarks
Participationofthe2-oxoaciddehydrogenasecomplexes
in redox regulation is summarized in Fig. 3. The interac-
tion of the complexes with the surrounding medium may
be realized through the lipoate-dependent ÔparacatalyticÕ
reactions. Such reactions allow the 2-oxo acid dehydro-
genase complexes to transform a signal in the form of
metabolite concentrations into chemical reactions such as
ROS production, thioredoxin reduction and E1 modifi-
cation. This network of reactions provides not only self-
regulation of the complexes (Fig. 3, boxed region), but
also their interaction with the surrounding medium, which
may be used in different signaling pathways. Our data on
the interplay between the mitochondrial complexes and
thioredoxin favors participation of the complexes in the
redox-dependent signaling through the thioredoxin sys-
tem. Other forms of such participation are known. For
example, the E1 subunit of the pyruvate dehydrogenase
Fig. 3. Participation of the 2-oxo acid dehydrogenase complexes in the
redox regulation of metabolism. The redox state of the surrounding
medium is sensed by the complexes through the concentrations of
2-oxo acid, CoA, NAD(H) and oxygen. This external signal is trans-
formed into the ratio of the complex-bound lipoate/dihydrolipoate.
Dependent on the input, this redox couple may regulate the activity of
the starting component E1 (a), produce output to surrounding medium
in the form of ROS (b) and reduce thioredoxin and disulfides (c).
Thioredoxin interferes with the self-regulation of the complexes (a),
concomitantly stimulating ROS production (b). The latter may be
overcome in the presence of SP-22.

Fig. 2. Thioredoxin catalysis of the dismutation of the thiyl radicals of
the complex-bound dihydrolipoate intermediate.
1040 V. I. Bunik (Eur. J. Biochem. 270) Ó FEBS 2003
complex from Azotobacter vinelandii wasshowntobindto
the fpr promoter region DNA, which is activated upon
cellular response to oxidative stress [61]. In Escheri-
chia coli, this promoter is activated by the redox-depend-
ent transcription factor SoxS. In view of the redox-sensing
function of the lipoate/dihydrolipoate couple of the
complex (Fig. 3) and the intimate link between this
couple and E1, the E1 dissociation from the complex to
bind DNA may represent another form of the lipoate-
dependent response under conditions of oxidative stress.
Acknowledgements
This work was supported by grants from DFG (438 17-159-92),
Volkswagen (I/69766) and Alexander von Humboldt (IV RUS 1003594
STP) Foundations. Critical reading of the manuscript by Prof J. J.
Mieyal (Case Western Reserve University, Cleveland, USA) is greatly
acknowledged.
References
1. Gilbert, H. (1984) Redox control of enzyme activities by thiol/
disulfide exchange. Methods Enzymol. 107, 330–351.
2. De Graef, M.R., Alexeeva, S., Snoep, J.L. & De Mattos, M.J.T.
(1999) The steady-state internal redox state (NADH/NAD
+
)
reflects the external redox state and is correlated with catabolic
adaptation in Escherichia coli. J. Bacteriol. 181, 2351–2357.
3.Pani,G.,Colavitti,R.,Borrello,S.&Galeotti,T.(2000)
Redox regulation of lymphocyte signalling. JUBMB Life 49,

381–389.
4. Rusnak, F. & Reiter, T. (2000) Sensing electrons: protein phos-
phatase redox regulation. Trends Biochem. Sci. 25, 527–529.
5. Danon, A. (2002) Redox reactions of regulatory proteins: do
kinetics promote specificity? Trends Biochem. Sci. 27, 197–203.
6. Mieyal, J.J., Srinivasan, U., Starke, D.W., Gravina, S.A. &Mieyal,
P.A. (1995) Glutathionyl specificity of thioltransferases: mecha-
nistic and physiological implications. In Biothiols in Health and
Disease (Packer, L. & Cadenas, E., eds), pp. 305–372. Marcel
Dekker, Inc., New York, USA.
7. Klatt, P. & Lamas, S. (2000) Regulation of protein function by
S-glutathionylation in response to oxidative and nitrosative stress.
Eur. J. Biochem. 267, 4928–4944.
8. Cross, A.R. & Jones, O.T.G. (1991) Enzymic mechanisms of
superoxide production. Biochim. Biophys. Acta 1057, 281–298.
9. Romero, F.J., Ordonez, I., Arduini, A. & Cadenas, E. (1992) The
reactivity of thiols and disulfides with different redox states of
myoglobin. J. Biol. Chem. 267, 1680–1688.
10. Winterbourn, C.C. & Metodieva, D. (1994) The reaction of
superoxide with reduced glutathione. Arch. Biochem. Biophys. 314,
284–290.
11. Nordberg, J. & Arner, E.S.J. (2001) Reactive oxygen species,
antioxidants, and the mammalian thioredoxin system. Free Rad.
Biol. Med. 31, 1287–1312.
12. Ullrich, V. & Bachschmid, M. (2000) Superoxide as a messenger of
endotelial function. Biochem. Biophys. Res. Communs. 278, 1–8.
13. Aslund, F., Zheng, M., Beckwith, J. & Storz, G. (1999) Regulation
of the OxyR transcription factor by hydrogen peroxide and
the cellular thiol-disulfide status. Proc. Natl Acad. Sci. USA 96,
6161–6165.

14. Schulz, J.B., Lindenau, J., Seyfried, J. & Dichgans, J. (2000)
Glutathione, oxidative stress and neurodegeneration. Eur. J.
Biochem. 207, 4904–4911.
15. Sastre, J., Pallardo, F.V. & Vina, J. (2000) Mitochondrial oxida-
tive stress plays a key role in aging and apoptosis. JUBMB Life
49, 427–435.
16. Skulachev, V.P. (2001) NAD(P)+ decomposition and antioxidant
defense of the cell. FEBS Lett. 492, 1–3.
17. Lisa, F.D. & Ziegler, M. (2001) Pathophysiological relevance of
mitochondria in NAD
+
metabolism. FEBS Lett. 492, 4–8.
18. Massey, V. (1963) Lipoyl dehydrogenase. Enzymes 7, 275–306.
19. de Kok, A. & van Berkel, W.J.H. (1996) Lipoamide dehy-
drogenase. In a-Keto Acid Dehydrogenase Complexes (Patel, M.S.,
Roche, T.E. & Harris, R.A., eds), pp. 53–70. Birkhaeuser Verlag,
Basel-Boston-Berlin.
20.Snoep,J.L.,deGraff,M.R.,Westphal,A.H.,deKok,A.,de
Mattos, M.J.T. & Neijssel, O.M. (1993) Differences in sensitivity
to NADH of purified pyruvate dehydrogenase complexes of
Enterococcus faecalis, Lactococcus lactis, Azotobacter vinelandii
and Escherichia coli: implications for their activity in vivo. FEMS
Lett. 114, 279–284.
21. Skrede, S., Bremer, J. & Elsjarn, L. (1965) The effect of disulfides
on mitochondrial oxidation. Biochem. J. 95, 838–884.
22. Skrede, S. (1968) The mechanism of disulfide reduction by mito-
chondria. Biochem. J. 108, 693–699.
23. Biewenga, G.P., Haenen, G.R. & Bast, A. (1997) The pharma-
cology of the antioxidant lipoic acid. Gen. Pharmacol. 29, 315–331.
24. Packer, L., Witt, E.H. & Tritschler, H.J. (1995) Alpha-lipoic acid

as a biological antioxidant. Free Rad. Biol. Med. 19, 227–250.
25. 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.
26. Suzuki, Y.J., Mizuno, M., Tritschler, H.J. & Packer, L. (1995)
Redox regulation of NF-kappa B DNA binding activity by
dihydrolipoate. Biochem. Mol. Biol. Internat. 36, 241–246.
27. Holmgren, A. (1979) Thioredoxin catalyzes the reduction of
insulin disulfides by dithiothreitol and dihydrolipoamide. J. Biol.
Chem. 254, 9627–9632.
28. Masutani, H. & Yodoi, J. (2002) Thioredoxin. Overview. Methods
Enzymol. 347, 279–286.
29. Asmus, K D. (1990) Sulfur-centered free radicals. Methods
Enzymol. 186, 168–180.
30. de Kok, A., Hengeveld, A.F., Martin, A. & Westphal, A.H. (1998)
The pyruvate dehydrogenase complex from Gram-negative
bacteria. Biochim. Biophys. Acta 1385, 353–366.
31. De la Sierra, I.L., Pernot, L., Prange, T., Saludjian, P., Schiltz, M.,
Fourme, R. & Padron, G. (1997) Molecular structure of the
lipoamide dehydrogenase domain of a surface antigen from
Neisseria meningitidis. J. Mol. Biol. 269, 129–141.
32. Perham, R.N. (2000) Swinging arms and swinging domains in
multifunctional enzymes: catalytic machines for multistep reac-
tions. Ann. Rev. Biochem. 69, 961–1004.
33. Berg, A. & de Kok, A. (1997) 2-Oxo acid dehydrogenase multi-
enzyme complexes. The central role of the lipoyl domain. Biol.
Chem. 378, 617–634.
34. 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.
35. Danson, M.J., Ferscht, A.R. & Perham, R.N. (1978) Rapid
intramolecular coupling of active sites in the pyruvate dehy-
drogenase complex of Escherichia coli: mechanism for rate
enhancement in a multimeric structure. Proc.NatlAcad.Sci.USA
75, 5386–5390.
36. Collins, J.H. & Reed, L.J. (1977) Acyl group and electron pair
relay system: a network of interacting lipoyl moieties in the pyru-
vate and a-ketoglutarate dehydrogenase complexes from
Escherichia coli. Proc. Natl. Acad. Sci. USA 74, 4223–4227.
37.Guest,J.R.,Ali,S.T.,Artymiuk,P.,Ford,G.C.,Green,J.
& Russel, G.C. (1990) Site-directed mutagenesis of dihydro-
lipoamide acetyltransferase and post-translational modification of
Ó FEBS 2003 Redox regulation of 2-oxo acid dehydrogenation (Eur. J. Biochem. 270) 1041
its lipoyl domains. In Biochemistry and Physiology of Thiamin
Diphosphate Enzymes (Bisswanger, H. & Ullrich, J., eds), pp. 176–
193. Chemie, Weinheim, Germany.
38. Guest, J.R., Attwood, M.M., Machado, R.S., Matqi, K.Y., Shaw,
J.E. & Turner, S.L. (1997) Enzymological and physiological con-
sequences of restructuring the lipoyl domain content of the pyru-
vate dehydrogenase complex of Escherichia coli. Microbiology 143,
457–466.
39. Dave, E., Guest, J.R. & Attwood, M.M. (1995) Metabolic
engineering in Escherichia coli: lowering the lipoyl domain content
of the pyruvate dehydrogenase complex adversely affects the
growth rate and yield. Microbiology 141, 1839–1849.
40. Bunik, V.I., Buneeva, O.A. & Gomazkova, V.S. (1990) Change
in a-ketoglutarate dehydrogenase cooperative properties due to
dihydrolipoate and NADH. FEBS Lett. 269, 252–254.

41. Bunik, V. (2000) Increased catalytic performance of the 2-oxo acid
dehydrogenase complexes in the presence of thioredoxin, a thiol-
disulfide oxidoreductase. J. Molec. Catalysis B: Enzymatic 8,
165–174.
42. Bunik, V. & Sievers, C. (2002) Inactivation of the 2-oxo acid
dehydrogenase complexes upon generation of intrinsic radical
species. Eur. J. Biochem. 269, 5004–5015.
43. Raddatz, G., Kruft, V. & Bunik, V. (2000) Structural determinants
for the efficient and specific interaction of thioredoxin with 2-oxo
acid dehydrogenase complexes. Appl. Biochem. Biotechnol. 88,
77–96.
44. Matthews, R.G. & Williams, C.H. Jr (1976) Measurement of the
oxidation-reduction potentials for two-electron and four-electron
reduction of lipoamide dehydrogenase from pig heart. J. Biol.
Chem. 251, 3956–3964.
45. Russel, R.R. III & Taegtmeyer, H. (1992) Coenzyme A sequestra-
tion in rat hearts oxidising ketone bodies. J. Clin. Invest. 89, 968–
973.
46. Gibson, G.E., Park, L.C.H., Sheu, K F.R., Blass, J.P. &
Calingasan, N.Y. (2000) a-Ketoglutarate dehydrogenase complex
in neurodegeneration. Neurochem. Int. 36, 97–112.
47. Bunik, V., Raddatz, G., Lemaire, S., Meyer, Y., Jacquot, J P. &
Bisswanger, H. (1999) Interaction of thioredoxins with target
proteins: Role of particular structural elements and electrostatic
properties of thioredoxins in their interplay with 2-oxo acid
dehydrogenase complexes. Prot. Sci. 8, 65–74.
48. Bunik, V. & Follmann, F. (1993) Thioredoxin reduction depen-
dent on a-keto acid oxidation by a-keto acid dehydrogenase
complexes. FEBS Lett. 336, 197–200.
49. Bunik, V., Shoubnikova, A., Loeffelhardt, S., Bisswanger, H.,

Borbe, H.O. & Follmann, H. (1995) Using lipoate enantiomers
and thioredoxin to study mechanism 2-oxo acid-dependent
dihydrolipoate production by the 2-oxo acid dehydrogenase
complexes. FEBS Lett. 371, 167–170.
50. Bryk, R., Lima, C.D., Erdjument-Bromage, H., Tempst, P.
& Nathan, C. (2002) Metabolic enzymes of mycobacteria linked to
antioxidant defense by a thioredoxin-like protein. Science 295,
1073–1077.
51. Bodenstein-Lang, J., Buch, A. & Follmann, H. (1989) Animal and
plant mitochondria contain specific thioredoxins. FEBS Lett. 258,
22–26.
52. Bunik, V., Follmann, H. & Bisswanger, H. (1997) Activation of
mitochondrial 2-oxo acid dehydrogenases by thioredoxin. Biol.
Chem. 378, 1125–1130.
53. Janin, J. (1997) Kinetics of protein-protein recognition. Proteins
28, 153–161.
54. Hanine, L.C., El Conte, D., Jacquot, J P. & Houee-Levin, C.
(2000) Redox properties of protein disulfide bond in oxidized
thioredoxin and lysozyme: a pulse radiolysis study. Biochemistry
39, 9295–9301.
55. 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.
56. Harman, L.S., Mottley, C. & Mason, R.P. (1984) Free
radical metabolites of
L
-cysteine oxidation. J. Biol. Chem. 259,
5606–5561.
57. Watabe,S.,Hiroi,T.,Yamamoto,Y.,Fujioka,Y.,Hagesawa,H.,

Yago, N. & Takahashi, S.Y. (1997) SP-22 is a thioredoxin-
dependent peroxide reductase in mitochondria. Eur. J. Biochem.
249, 52–60.
58.Chen,Y.,Cai,J.,Murphy,T.J.&Jones,D.P.(2002)Over-
expressed human mitochondrial thioredoxin confers resistance to
oxidant-induced apoptosis in human osteosarcoma cells. J. Biol.
Chem. 277, 33242–33248.
59. Cabiscol, E., Piulats, E., Echave, P., Herrero, E. & Ros, J. (2000)
Oxidative stress promotes specific protein damage in Saccharo-
myces cerevisiae. J. Biol. Chem. 275, 27393–27398.
60. 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.
61. Regnstrom, K., Sauge-Merle, S., Chen, K. & Burgess, B.K. (1999)
In Azotobacter vinelandii, the E1 subunit of the pyruvate dehy-
drogenase complex binds fpr promoter region DNA and ferre-
doxin I. Proc. Natl Acad. Sci. USA 96, 12389–12393.
1042 V. I. Bunik (Eur. J. Biochem. 270) Ó FEBS 2003