ISSN 0006-2979, Biochemistry (Moscow), 2026, Vol. 91, No. 6, pp. 893-909 © Pleiades Publishing, Ltd., 2026.
893
REVIEW
Interplay between Neuronal Metabolism
and Signaling in Model Systems with Impaired
α-Ketoglutarate Dehydrogenase Complex Activity
Márton Kokas
1,a
, Attila Ambrus
1,b
, László Tretter
1,c
, and Tímea Komlódi
1,d
*
1
Department of Biochemistry, Semmelweis University, Budapest, H-1094, Hungary
a
e-mail: kokas.marton@semmelweis.hu
b
e-mail: ambrus.attila@semmelweis.hu
c
e-mail: tretter.laszlo@semmelweis.hu
d
e-mail: komlodi.timea@semmelweis.hu
Received March 5, 2026
Revised June 22, 2026
Accepted June 22, 2026
Abstract— The α-ketoglutarate dehydrogenase complex (KGDHC) serves as a master regulator of cell’s mo-
lecular machinery. Beyond its classical role as a rate-limiting enzyme in the tricarboxylic acid (TCA) cycle,
KGDHC has emerged as a critical redox sensor that can act as both a source and a target of reactive ox-
ygen species (ROS), thereby regulating cellular redox homeostasis. This review summarizes evidence from
genetically modified animal models and cell culture studies demonstrating that compromised KGDHC ac-
tivity affects neuronal metabolism, redox homeostasis, and cellular signaling. KGDHC dysfunction causes
mitochondrial failure, resulting in reduced ATP synthesis and activation of AMP-activated protein kinase
(AMPK). Although inhibition of KGDHC reduces mitochondrial ROS formation, it also disrupts physiological
ROS-dependent signaling mechanisms. In KGDHC-deficient mice, impaired ROS signaling and energy deficit
decrease brain adaptability, increase susceptibility to neurotoxins, and disrupt crucial pathways by down-
regulating PGC-1α and Nrf2. These alterations result in suppression of antioxidant defences and lead to
neuronal death in the hippocampus and memory impairment. Moreover, KGDHC dysfunction induces mito-
chondrial fragmentation and is strongly linked to excitotoxicity, further accelerating neuronal dysfunction.
As observed in heterozygous models, even partial KGDHC deficiency can exacerbate persisting cellular and
mitochondrial defects, leading to the development of more severe pathological conditions.
DOI: 10.1134/S0006297926600663
Keywords: α-ketoglutarate dehydrogenase complex, neuronal signaling, cognitive decline, reactive oxygen
species, succinylation
* To whom correspondence should be addressed.
INTRODUCTION
α-Ketoglutarate dehydrogenase complex (KGDHC),
also referred to as 2-oxoglutarate dehydrogenase
complex (EC 1.2.1.105), is a member of the α-keto
acid dehydrogenase enzyme family. It catalyzes a
highly regulated conversion of α-ketoglutarate (α-KG)
to succinyl-CoA with the generation of NADH in the
tricarboxylic acid (TCA) cycle. NADH supplies elec-
trons to the electron transport system (ETS), thereby
supporting mitochondrial ATP synthesis via oxidative
phosphorylation (OXPHOS), which accounts for >80 %
of the total cellular ATP production [1]. Succinyl-CoA
also serves as a substrate for mitochondrial sub-
strate-level phosphorylation, enabling ATP (or GTP)
synthesis even in the absence of oxygen [2], inde-
pendently of the ETS. Succinyl-CoA is also a precursor
in porphyrin synthesis. Compromised KGDHC activity
negatively affects mitochondrial substrate-level phos-
phorylation by limiting succinyl-CoA availability to
succinyl-CoA ligase [3].
KGDHC consists of three types of subunits:
1) α-ketoglutarate dehydrogenase (E1k; OGDH and
OGDHL genes), 2) dihydrolipoyl succinyltransferase/
transsuccinylase (E2k; DLST gene), and 3) dihydroli-
poamide dehydrogenase (E3; DLD gene). The structure
and reaction mechanisms of KGDHC are reviewed
in detail in [4-6]. Homozygous disruption of any
KOKAS et al.894
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
of these genes results in embryonic lethality, indicat-
ing an essential role of KGDHC in early embryonic de-
velopment [7-12]. In mammals, E2k and E3 are each
encoded by a single gene copy, whereas the E1k sub-
unit has two isoforms: ubiquitously expressed protein
encoded by OGDH and its brain-specific paralog en-
coded by OGDHL gene, which arose from a common
ancestral gene and is located on chromosome 10.
[13-15]. In rodent brain, the OGDH : OGDHL ratio is
approximately 60 : 40. The E1k/E2k and E1k/E3 ra-
tios are lower in the brain compared to the heart.
A recent study using triple-transgenic Alzheimer’s
disease (AD) mouse model reported OGDHL down-
regulation in the cortex and the CA1 subfield of the
hippocampus [16]. The E3 subunit is shared among
several multienzyme complexes, including the glycine
cleavage system (glycine decarboxylase complex) and
four distinct α-keto acid dehydrogenase complexes:
pyruvate dehydrogenase complex (PDHC), KGDHC,
branched-chain α-keto acid dehydrogenase complex,
and α-ketoadipate dehydrogenase complex (KADHC).
Importantly, KADHC also shares the E2k subunit and
has a highly homologous E1a subunit (also known
as dehydrogenase E1 and transketolase domain con-
taining 1, DHTKD1) with KGDHC in mammals [17, 18].
KADHC plays a crucial role in the degradation of
L-lysine and tryptophan by converting α-ketoadipate
into glutaryl-CoA and NADH. Recent structural analy-
sis using liquid chromatography/mass spectrometry
(LC-MS) and molecular dynamics simulations has
shown that E1a and E1k bind to different sites on
E2k, with different E1 residues contributing to the
complex formation [19]. In contrast to the embry-
onic lethality observed in models with knocked out
KGDHC-encoding genes, DHTKD1
−/−
mice are via-
ble [20], suggesting that KGDHC can at least partially
compensate for the loss of KADHC. For a comprehen-
sive structural analysis, see review [4].
KGDHC has also been shown to localize to the
nucleus, where it modulates gene expression via his-
tone succinylation [21], a key epigenetic modifica-
tion implicated in tumor proliferation and progres-
sion [22, 23].
Beyond its canonical role in energy production,
KGDHC is critically involved in amino acid metabo-
lism. Glutamate is converted to α-KG by glutamate
dehydrogenase (GDH) with the release of ammonia.
Conversely, α-KG can be transaminated to glutamate
in the reactions involving aspartate or alanine. Glu-
tamate is the principal excitatory neurotransmitter
governing essential functions, including learning,
cognition, and memory [24, 25]. However, excessive
glutamate accumulation can trigger excitotoxicity,
leading to neuronal damage, e.g., in pathologies such
as stroke and neurodegenerative diseases [26-28].
It was reported that KGDHC inhibition by succinyl
phosphonate (SP) promoted glutamate accumulation
and induced neuronal death in primary neuronal cul-
tures [29, 30]. SP-treated non-pregnant rats exhibited
elevated cerebellar concentrations of several amino
acids, including glutamate and alanine, compared to
untreated animals [31]. Recently, it was demonstrat-
ed that although thiamine pretreatment increased ve-
sicular glutamate levels, while pretreatment with SP
decreased them, these changes did not explain their
respective effects on excitotoxicity [32]. Instead, im-
paired KGDHC activity was found to promote gluta-
mate-induced excitotoxicity through the metabotropic
regulation of NMDA receptors in primary neuronal
cultures, indicating that susceptibility to excitotoxic
damage depends more on the neuronal responsive-
ness to glutamate than on the amount of glutamate
released.
In astrocytes, glutamate can be converted to glu-
tamine by glutamine synthase through incorporation
of a molecule of ammonia in an ATP-dependent reac-
tion. Astrocytes supply glutamine to neurons, where
it is converted back to glutamate by glutaminase. This
cycle proves a close cooperation between astrocytes
and neurons and demonstrates the tight metabolic
interplay between α-KG and amino acid metabolism.
KGDHC integrates energy metabolism with cel-
lular signaling pathways, as its substrate α-KG is a
co-substrate for the ketoglutarate-dependent diox-
ygenases (KGDDs). These enzymes primarily cata-
lyze the hydroxylation or demethylation of histones
and DNA, thereby controlling epigenetic modifica-
tions [33]. The most well-known role of KGDDs is the
regulation of hypoxia-inducible factor 1 (HIF-1), a
master transcriptional regulator of cellular response
to hypoxia. Under low-oxygen conditions, HIF-1 ac-
tivates the transcription of genes involved in angio-
genesis, glycolysis, and erythropoiesis [34, 35]. Despite
extensive evidence supporting these α-KG-dependent
mechanisms, to the best of our knowledge, no studies
have yet investigated these processes in KGDHC-defi-
cient animal models or cell cultures.
KGDHC dysfunction has been documented inAD,
as well as in several other neurodegenerative dis-
eases and brain injuries, indicating the vital role of
KGDHC in cell metabolism and signaling [5]. Inter-
estingly, the Swedish mutation (APP670/671) causing
early-onset familial AD is associated with reduced
immunoreactivity of E1k (~50% of control) and E2k
(~25% of control), but not of the shared E3 subunit,
in brain samples [36]. In contrast, an earlier study
in sporadic AD patients reported reduction (23 to
41%) of all three subunits in the temporal cortex.
Furthermore, E1k and E3 levels were reduced in the
parietal cortex, while only E3 was downregulated
in the hippocampus [37]. Reduced KGDHC immunos-
taining was also reported in the substantia nigra [38]
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 895
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
and pathophysiologically unaffected brain areas
of patients with Parkinson’s disease [39]. Important-
ly, regional and cell type-specific heterogeneity of the
KGDHC overall activity has been observed through-
out the brain [5]; hence, establishing direct associa-
tion between altered KGDHC activity and neurolog-
ical disorders remains challenging.
To move beyond purely observational studies,
the use of genetically modified animals has become
essential. Although cellular models provide valuable
mechanistic insights, animal models offer a more
comprehensive framework for investigating complex
relationship between cellular metabolism and sig-
naling, while also revealing systemic consequences
of impaired KGDHC activity, including cognitive de-
cline and behavioural changes. In this review, we
summarize key findings from genetically modified
KGDHC-deficient cell lines, rodent models, and other
organisms such as Drosophila. We also examine the
effects of KGDHC downregulation or dysfunction in-
duced by employing alternative approaches, including
the use of specific inhibitors, such as succinyl phos-
phonate (SP). Particular emphasis is placed on the
central role of KGDHC in bioenergetics, cellular sig-
naling, and redox homeostasis.
STUDIES IN GENETICALLY MODIFIED
ANIMALS AND CELL LINES
WITH KGDHC DYSFUNCTION
Behavioral consequences in KGDHC-deficient
rodent models. The generation of genetically mod-
ified cell lines and animal models with compro-
mised KGDHC activity has provided valuable tools
for investigating various roles of KGDHC in cellu-
lar and mitochondrial metabolism and signaling at
the cellular and whole-body level [8, 10, 11, 40-45].
Heterozygous knockout mice partially deficient by
E1k (OGDH
+/−
), E2k (DLST
+/−
), and E3 (DLD
+/−
) mice
and double knockout heterozygous E2k/E3 (DLST
+/−
/
DLD
+/−
) mice have been successfully generated [3, 7,
8, 40, 41, 43]; however, homozygous deletion of any of
these genes resulted in embryonic lethality [7-9]. Het-
erozygous DLST
+/−
or DLD
+/−
mice exhibited no overt
phenotypic alteration [8, 41], whereas middle-aged
DLST
+/−
/DLD
+/−
mice developed several neuropatho-
logical alterations, including microgliosis in the ce-
rebral cortex, neuronal death in the hippocampus,
and disruptions in the mitochondrial biogenesis and
dynamics [43]. These structural and cellular changes
indicate neuronal involvement and minor cognitive
decline that have also been detected in standard test-
ing paradigms [43]. Specifically, DLST
+/−
/DLD
+/−
mice
showed reduced locomotor activity and impaired
performance in fatigue-endurance running tests [43].
Furthermore, pharmacological inhibition of KGDHC
by injection of SP or triethyl succinyl phosphonate
(TESP, membrane-penetrating SP derivative) increased
anxiety behavior in rats compared to control ani-
mals [46]. In the study by Graf et al. [47], SP-treated
rats subjected to stress (ethanol injection) demonstrat-
ed decreased locomotor activity and prolonged sleep
duration relative to the controls [47]. These studies
indicate that partial KGDHC inhibition induced either
pharmacologically (e.g., with low SP dosage) or by
heterozygous knockout of KGDHC subunits, is largely
asymptomatic under normal physiological conditions.
However, it compromises adaptive and compensatory
mechanisms during stress or disease, leading to overt
pathophysiological manifestations. Thus, heterozygous
E1k knockout rats displayed no significant changes
in animal behavior, blood biochemistry, and tissue
histology compared to controls [10]. However, when
fed with a high-fat diet, these animals suffered from
liver dysfunction, further suppoting the notion that
additional stressors could lead to manifistations of
pathophysiological consequences of KGDHC deficien-
cy. It should be noted, however, that no neurological
outcomes or amyloid plaque formation have been as-
sessed in this study.
KGDHC-deficient Drosophila model. Bi-allelic
introduction of human disease-associated mutations
of OGDHL into Drosophila flies caused developmental
lethality and marked disturbances in the TCA cycle
metabolism [11]. The flies exhibited severe defects in
recovery and pronounced locomotor delays follow-
ing mechanical stress. Moreover, the concentration
of α-KG in the brains of OGDHL-deficient larvae was
1.5 times higher compared to the control. However,
interpretation of these findings is complicated by the
fact that Drosophila genome contains only one OGDH
gene. Recently, it was reported that introduction of
heterozygous (monoallelic) human mutant variants
of OGDHL in Drosophila caused no developmental
defects, but instead led to age-dependent locomotion
impairments [48]. Other heterozygous mutations were
associated with reduced KGDHC activity, abnormal
proteolytic cleavage, and impaired mitochondrial im-
port. Although heterozygous mutations in OGDHL are
generally insufficient to cause overt human disease,
they contribute to the development of late-onset neu-
rological disorders.
Metabolic consequences of KGDHC dysfunc-
tion. Metabolic and bioenergetic studies are essential
for understanding how KGDHC deficiency disrupts
the TCA cycle and cellular energy production. These
investigations provide mechanistic insight into the
contribution of impaired KGDHC activity to oxidative
stress and metabolic failure in affected tissues.
Synaptic and non-synaptic mitochondria respond
differently to KGDHC dysfunction. According to
KOKAS et al.896
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
Berndt et al. [49], synaptic mitochondria are signifi-
cantly more sensitive to KGDHC inhibition and dis-
play membrane depolarization and pronounced de-
cline in ATP production at only 30% inhibition [49].
In contrast, non-synaptic mitochondria are more ro-
bust and maintain their function up to 70% KGDHC
inhibition. This selective vulnerability of synaptic mi-
tochondria has been proposed as a key factor in the
early synaptic degeneration and neural disintegration
in the Parkinson’s disease and AD [49, 50].
Metabolic tracing experiments using labeled
glucose revealed increased glucose concentration in
the cortex of DLST
+/−
mice. However, no alterations
were detected in glucose levels in the cerebellum,
concentrations of lactate or alanine, or the content
of TCA cycle intermediates in the cortex [51]. These
findings further support that the heterozygous knock-
out of any individual KGDHC subunit is insufficient
to dramatically alter metabolism under resting con-
dition. Furthermore, homozygous DLST knockout
in H838 cells caused no significant changes in cell
growth [52], although these cells displayed deple-
tion of four-carbon TCA cycle intermediates (succi-
nate, fumarate, malate, and aspartate), alongside a
marked accumulation of 2-hydroxyglutarate (2-HG),
while α-KG levels remained unaffected compared to
controls [52]. In our studies, mitochondrial O
2
con-
sumption supported by α-KG was reduced by 28, 22
and 43% in the brain mitochondria isolated from
DLST
+/−
, DLD
+/−
, and DLST
+/−
/DLD
+/−
animals, respec-
tively, reflecting an additive inhibitory effect of the
double heterozygosity [3, 42, 43]. Furthermore, brain
mitochondria from DLST
+/−
and DLD
+/−
mice exhibit-
ed reduced oxygen consumption when energized with
glutamate + malate (GM) [3]. Similarly, OGDHL-defi-
cient neuroblastoma SH-SY5Y cells showed impaired
mitochondrial respiration (~60% of control) when cul-
tured in a medium supplemented with glucose, py-
ruvate, and glutamine [11]. Collectively, these results
emphasize KGDHC involvement in glutamate metab-
olism. In accordance with this, the rate of α-KG-sup-
ported ATP synthesis in the brain was reduced in all
KGDHC-deficient groups [43]. Kiss et al. [3] reported
that inhibition of adenine nucleotide translocase
(ANT) with carboxyatractyloside (CATR) in respira-
tion-impaired mitochondria energized with GM (with
or without β-hydroxybutyrate) caused a decrease in
the mitochondrial membrane potential (ΔΨ
m
) in mi-
tochondria from DLD
+/−
and DLST
+/−
/DLD
+/−
mice,
indicating that ANT was operating
in the reverse
mode, importing extramitochondrial ATP. In the con-
trol mitochondrial, however, CATR caused mitochon-
drial membrane hyperpolarization, reflecting normal
ANT function [3]. Furthermore, increased ATP efflux
was observed in DLD
+/−
brain mitochondria supplied
with GM. It can be concluded that impaired KGDHC
activity limits succinyl-CoA availability for sub-
strate-level phosphorylation via succinyl-CoA ligase,
resulting in lower ATP/ADP ratio in the mitochondrial
matrix. The drop in the ATP/ADP ratio shifts the ANT
reversal potential, resulting in the reversal of ANT
transport activity and utilization of extramitochon-
drial ATP [2, 53]. A decrease in ATP levels activates
AMP-activated protein kinase (AMPK), which is a cen-
tral regulator of intracellular signaling in the fasting
state (for review see [54, 55]). Both oxidative stress
and OXPHOS inhibitors can activate AMPK through
canonical pathways [56]. Although AMPK activation
is not unique to KGDHC (or even E3) dysfunction, its
role is often overlooked in discussions of mitochon-
drial energy failure. Klivenyi et al. [57] in 2004 and
Yang et al. [8] in 2009 reported that both DLD
+/−
and
DLST
+/−
mice exhibited increased vulnerability to neu-
rotoxins commonly used to mimic symptoms of Par-
kinson’s and Huntington’s diseases. Crossing DLST
+/−
mice with a transgenic mouse model for amyloid
deposition (Tg19959) increased amyloid plaque bur-
den in the offspring. In addition, DLST deficiency re-
sulted elevated the levels of Aβ oligomers and nitroty-
rosine, caused a higher incidence of memory deficit,
and accelerated decline in spatial learning in female
mice [58]. These studies suggested that KGDHC dys-
function compromises brain’s adaptative capacity and
contributes to the pathogeneses of neurodegenerative
diseases. Studies in SH-SY5Y neuroblastoma cells ex-
pressing DLST gene truncated starting from intron 7,
demonstrated that the product of this alternatively
spliced gene (termed MIRTD) contributes to the ETC
biogenesis [59].
ROS production and redox signaling upon
KGDHC impairment. ROS are inevitable by-prod-
ucts of aerobic respiration in cells and mitochondria.
Atphysiological concentrations, typically in the nano-
molar range, ROS act as signaling molecules, primarily
by oxidizing specific cysteine residues (a mechanism
referred to as oxidative eustress) [60, 61]. In contrast,
elevated ROS levels (e.g. ≥100 nM H
2
O
2
) cause oxida-
tive distress (damage) [61]. Superoxide and hydrogen
peroxide are two (patho)physiologically important
oxygen species implicated in signaling pathways reg-
ulating cell differentiation, proliferation, antioxidant
and stress responses, maintenance of ion gradients,
pH buffering capacity, and various anabolic and cat-
abolic reactions (see [61-64] for detail). Multiple in-
tracellular sites contribute to ROS production, with
NADPH oxidases and mitochondria representing the
predominant sources [64-67]. In mitochondria, more
than ten ROS-producing enzymes have been identi-
fied [64-66]. Among these, KGDHC may serve as a ma-
jor ROS-generating site in isolated mitochondria when
the NADH/NAD
+
ratio is elevated, such ad during dys-
function of the TCA cycle or the ETS [6, 41, 68-70].
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 897
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
Importantly, KGDHC is both a source and a target
of ROS [6, 71-73]. Covalently tethered dihydrolipoic
acid (LAH
2
), which acts as a E2k cofactor, can be ir-
reversibly inactivated by H
2
O
2
and lipid peroxidation
products [73, 74], leading to compromised KGDHC ac-
tivity and subsequent impairment of mitochondrial
bioenergetics, i.e., processes implicated in the devel-
opment of neurological disorders, among others [5].
LAH
2
in E2k can also be reversibly inactivated by
glutathionylation or nitrosylation [75-79]. The extent
of lipoic acid glutathionylation correlates with both
the degree of enzyme inhibition and its subsequent
reactivation [80]. Although these modifications tem-
porarily and locally reduce the KGDHC activity, they
likely simultaneously protect the enzyme from other,
potentially irreversible, oxidative modifications (see
above) [80, 81]. Indeed, both S-glutathionylation and
S-nitrosylation reduce the KGDHC-dependent H
2
O
2
formation, indicating that these reversible modifi-
cations (i) regulate redox signaling via KGDHC and
(ii) exert an antioxidant effect by protecting the thiol
group of LAH
2
in KGDHC from reactive species (e.g.,
4-hydroxy-2-nonenal) or irreversible oxidation by
H
2
O
2
during oxidative stress [78, 80-82].
Isolated brain mitochondria from heterozygous
DLST
+/−
mice did not exhibit a significant reduction
in H
2
O
2
production supported by either succinate
or α-KG, nor in the α-KG-supported H
2
O
2
formation
under conditions of the elevated NADH/NAD
+
ra-
tio [41-43]. In contrast, in DLD
+/−
and DLST
+/−
/DLD
+/−
mice, these rates were significantly reduced [42, 43].
Because DLST is essential for the KGDHC-associat-
ed ROS generation [72], these findings suggest that
the impairment of reverse electron transfer (RET)-
induced H
2
O
2
production initiated by succinate in
brain mitochondria from DLD
+/−
mice is attributable
to DLD present in its free form or associated with
DLD- binding complexes other than KGDHC.
Specific inhibition of the KGDHC subunit E1k has
been shown to reduce glutamate-induced ROS forma-
tion in neurons, as measured in situ using hydroethi-
dine, thus emphasizing the contribution of KGDHC
to glutamate-driven ROS generation [83]. Stimulation
of neurons with glutamate increases intracellular
calcium concentration, which not only induces the
KGDHC enzymatic activity, but also promotes its ROS
generating capacity [71].
According to current knowledge, RET-driven ROS
generation plays an important role in ischemia–reper-
fusion injury [84, 85]. During ischemia, succinate ac-
cumulates; upon reperfusion, its oxidation supports
RET-associated ROS formation, leading to oxidative
damage. A significant decrease in the RET-driven
ROS formation in the DLD
+/−
and DLST
+/−
/DLD
+/−
brain mitochondria suggests that partial impairment
of KGDHC activity might confer protection against
ischemia–reperfusion injury. However, it is important
to point out that although reduction in ROS formation
is beneficial, it may also disrupt essential ROS-medi-
ated physiological signaling mechanisms.
Notably, transient decrease in KGDHC activity
may trigger protective cellular responses such as
glutathionylation, whereas prolonged KGDHC inhibi-
tion leads to increased oxidative stress, calcium im-
balance, and cell death, as demonstrated in models
with impaired KGDHC activity [36]. In heterozygous
DLD
+/−
and DLST
+/−
/DLD
+/−
mice, significant reduc-
tion in H
2
O
2
production in brain mitochondria was
observed in situ using different mitochondrial respi-
ratory substrates [41-43]. However, Calingasan et al.
[86] reported increased lipid peroxidation in the sub-
granular zone of the hippocampal dentate gyrus, indi-
cating increased oxidative stress in DLST
+/−
mice [86]
and increased lipid peroxidation in the striatum of
DLD
+/−
mice [57], that were associated with reduced
neurogenesis and cognitive decline. Discrepancies be-
tween direct ROS measurements and oxidative dam-
age markers such as lipid peroxidation might be due
to the fact that ROS measurements reflect immediate
ROS production, while oxidative stress markers indi-
cate localized, cumulative oxidative damage. Conse-
quently, utilizing multiple complementary techniques
is necessary, as significant tissue-specific oxidative
damage may occur even when isolated mitochondrial
assays suggest reduced ROS generation.
Signaling, cognitive decline, neuronal death,
and neurodegenerative diseases. The role of ROS
in cellular signaling pathways, especially in processes
underlying memory and learning, has been intensive-
ly studied, with a major focus on glutamate receptor
activation in the hippocampus [87, 88]. It was shown
that glutamate receptor agonists increase mitochon-
drial ROS production in postsynaptic neurons, leading
to enhancement of protein kinase C (PKC) signaling
activity [87, 88]. Consistent with this, scavenging ROS
in slices prepared from the hippocampal CA1 subfield
reduced PKC signaling and impaired long-term poten-
tiation (LTP), a well-established cellular correlate for
learning and memory [89-92]. We recently reported
that the DLST
+/−
/DLD
+/−
mice described above suffered
from massive neuronal loss in the CA1 subfield [43];
which also displayed a stronger KGDHC immunore-
activity than the CA3 subfield [5]. These data suggest
that disrupted H
2
O
2
signaling in the brain of DLST
+/−
/
DLD
+/−
animals may lead to cognitive decline and
neuronal death via the PKC signaling pathway.
We further demonstrated that peroxisome prolif-
erator-activated receptor gamma coactivator 1-alpha
(PGC-1α) and nuclear factor erythroid-derived 2-like 2
(Nrf2, also referred to as NFE2L2; not to be confused
with the nuclear respiratory factor 2), which are
the key regulators of mitogenesis and antioxidant
KOKAS et al.898
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
Fig. 1. Consequences of KGDHC dysfunction for cellular signaling in neurodegeneration and ageing. DKO, double knockout
(DLST
+/–
/DLD
+/–
).
response, were downregulated in the brain of DLST
+/−
/
DLD
+/−
mice [43]. The tumor suppressor p53 is known
to regulate both PGC-1α and Nrf2, as well as super-
oxide dismutase 2 (SOD2) [93]. Aquilano etal. [94] re-
ported that glutathione depletion induced generation
of ROS and reactive nitrogen species (RNS) in SH-SY5Y
cells [94], leading to PGC-1α upregulation associat-
ed with the increased Nrf2-mediated expression of
SOD2 [93]. In line with that, brain mitochondria iso-
lated from DLST
+/−
/DLD
+/−
mice demonstrated reduced
H
2
O
2
production rates on different respiratory sub-
strates, while immunohistological staining confirmed
downregulated expression of both Nrf2 and PGC-1α,
highlighting the regulatory role of KGDHC-mediat-
ed ROS signaling in mitochondrial biogenesis [43]
(Fig. 1). Consistently with our results, Nrf2-deficient
mice exhibited reduced LTP accompanied by im-
pairments in memory and learning [95]. Moreover,
decreased PGC-1α expression has been observed in
early AD, suggesting that KGDHC dysfunction may be
crucial in the progression of certain neurodegenera-
tive diseases [96].
Mitochondrial dynamics. Neurons and glial cells
isolated from DLST
+/−
/DLD
+/−
mice displayed acceler-
ated mitochondrial fragmentation and reduced fusion
rates, indicating a shift toward impaired mitochon-
drial dynamics, which is in line with previous stud-
ies reporting elevated expression of dynamin-related
protein 1 (Drp1) and downregulation of mitofusin 2
(Mfn2) in the cortices of AD patients and in SH-SY5Y
neurons with reduced KGDHC activity [43, 97, 98].
Frank et al. [99] reported that even mild stress can
trigger elevated Drp1 levels in human cells. The bal-
ance between mitochondrial fusion and fission is es-
sential for post-mitotic neurons, as its disruption leads
to serious neurological pathologies (e.g., Charcot–
Marie–Tooth disease type 2A) [100]. Further support
comes from Toyoma et al. [101], who reported that
enhanced AMPK signaling (resulting from reduced
ATP production by OXPHOS) promotes mitochondrial
fission by increasing Drp1 recruitment to the mito-
chondria through phosphorylation of mitochondrial
fission factor. Collectively, these findings suggest that
mitochondrial dynamics, indirectly regulated by the
KGDHC activity, may represent a critical factor in the
progression of neurodegeneration.
Further studies are needed to clarify which
KGDHC-related pathways are more relevant under
different physiological and pathological conditions.
Notably, E1k reduction may increase the α-KG/succi-
nate ratio, which has been associated to antitumor
effects. In contrast, reduced E2k and E3 levels are
linked to mitochondrial fragmentation and dysfunc-
tion, leading to the downregulation of signaling path-
ways (e.g., Nrf2-mediated stress response).
Succinylation of lysine residues is a posttrans-
lational modification that controls several signal-
ing pathways [102]. Metabolic disturbances such as
TCA cycle dysregulation, decreased ATP production,
or ETC impairment, can suppress protein succinyla-
tion, whereas hypoxic conditions enhance it [103].
Accumulation of succinyl-CoA, for example due to
the impaired TCA cycle activity, promotes protein
succinylation and may further exacerbate patholog-
ical conditions associated with neuronal and/or met-
abolic disorders [104]. In cultured mouse neurons
and neuronal cell lines, selective KGDHC inhibition
significantly reduced both cytosolic and mitochon-
drial protein succinylation [105]. Conversely, reduced
KGDHC activity has been shown to decrease mito-
chondrial protein succinylation, while increasing
cytosolic succinylation, potentially due to the E2k
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 899
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
translocation from mitochondria to the cytosol upon
ETC impairment [106]. Enhanced extramitochondrial
protein succinylation, mediated by KGDHC has been
implicated in plaque and tangle formation in AD, con-
sistent with decreased KGDHC activity in this disease
[106]. Purified KGDHC can also function as a trans-
succinylase due to the succinyltransferase activity
of the E2k subunit, enabling it to succinylate other
TCA cycle enzymes, as well as PDHC. This means that
KGDHC can indirectly control acetylation via PDHC,
whereas PDHC can modulate the KGDHC activity via
acetylation. According to the Gibson’s group, the suc-
cinylation activity of KGDHC is more efficient than
non-enzymatic succinyl-CoA-dependent succinylation,
highlighting the functional importance of the E2k
subunit [105]. Furthermore, KGDHC activity was re-
duced by succination in Complex I deficiency Leigh
syndrome [107]. Succination is irreversible posttrans-
lational modification of cysteine thiol groups with
fumarate with the formation of S-(2-succinyl)cysteine
[108]. A decrease in the KGDHC activity limits succi-
nyl-CoA availability, thereby reducing substrate-level
phosphorylation and further aggravating cellular en-
ergy deficit [107].
Lipoylation. Beal’s group previously reported
that lipoic acid may exert beneficial effects in neu-
rodegenerative disorders, including Huntington’s
disease and amyotrophic lateral sclerosis (ALS) [109-
111]. A more recent study in genetically modified
mice and human striatal organoids demonstrated that
in Huntington’s disease, lipoylation of E2k is reduced,
with no significant alterations in the lipoylation of
the closely related E2p subunit of PDHC or expres-
sion levels of both KGDHC and PDHC subunits [112].
These findings point out the importance of posttrans-
lational modification of proteins in different patho-
physiological conditions, as well as suggest that lipo-
ic acid, a well-known antioxidant, may have a more
complex role in mitochondrial function rather than
acting as a ROS scavenger. However, further research
is needed to prove this hypothesis.
ALTERNATIVE APPROACHES
TO MODIFICATION OF KGDHC ACTIVITY
Succinyl phosphonate (SP) and its derivatives
act as highly potent and selective KGDHC inhibitors
by mimicking the structure of its natural substrate,
α-KG [113, 114]. By binding to E1k, SP effectively
blocks the TCA cycle, resulting in the elevated levels
of α-KG and pyruvate, significant reduction in mito-
chondrial ATP synthesis, and upregulated ROS gener-
ation. This metabolic blockade triggers compensatory
alterations in the amino acid metabolism, including
increase in amino acid levels [30, 31, 115], which in
turn affect cellular signaling pathways and ultimate-
ly contribute to neuronal cell death signaling [30].
Such metabolic shutdown is particularly effective in
slowing the growth of cancer cells, particularly glio-
blastoma, which often rely on KGDHC for survival
and biosynthesis [115]. Interestingly, at certain con-
centrations, SP can act as a neuroprotective agent
by preventing glutamate excitotoxicity [116, 117] and
glutamate- induced overproduction of ROS during
metabolic stress [83]. Importantly, low-dose SP ad-
ministration in rats induced no significant energy
insufficiency [47]. However, under stress condition,
e.g. ethanol administration, SP provoked energy defi-
cit accompanied by decreased locomotor activity, re-
duced muscle strength, and prolonged narcotic sleep
duration. In addition, SP and its phosphonoethyl es-
ter derivative (PESP) decreased the viability of mouse
B cell activated by lipopolysaccharide (LPS) and in-
terleukin-4 [118], consistent with findings in DLST
+/−
mice, which also showed reduced B cell viability.
Thiamine antivitamins (antimetabolites) such
as pyrithiamine and oxythiamine, are compounds
that structurally resemble thiamine (vitamin B1) but
interfere with its biological function and, therefore,
are widely used to model thiamine deficiency [119-
121]. These compounds target enzymes that require
thiamine pyrophosphate (active form of thiamine)
as a cofactor. Among these enzymes, KGDHC is con-
sidered the most critical link between thiamine de-
ficiency and neurodegeneration [122-128]. In animal
models, KGDHC downregulation due to administra-
tion of thiamine antivitamins induced localized lactic
acidosis and brain energy metabolism failure, pro-
moting glutamate-induced neuronal depolarization in
vulnerable areas and ultimately causing NMDA-recep-
tor mediated excitotoxicity followed by apoptotic cell
death [129, 130]. Other mechanisms, such as oxidative
stress and blood–brain barrier impairment, also con-
tribute to cell loss in thiamine deficiency [126, 129,
131]. Manifestations of thiamine deficiency include
severe metabolic, neurological, and cardiovascular
dysfunction, such as fatigue, cognitive decline, pares-
thesia, and muscle weakness [122, 132]. Chronic alco-
holism is a major risk factor and commonly leads to
thiamine deficiency, manifested as the Wernicke’s en-
cephalopathy [122, 130, 132]. Thiamine deficiency-in-
duced oxidative stress in rats caused localized brain
lesions and abnormal accumulation of amyloid pre-
cursor protein (APP) in damaged neurons, providing
a relevant model for elucidating the link between me-
tabolism, APP dysregulation, and neurodegeneration
in disorders such as Wernicke–Korsakoff syndrome
and AD [133].
Lipoate analogues are synthetic compounds cre-
ated by structural modification of naturally occurring
α-lipoic acid, a key cofactor in thiamine-dependent
KOKAS et al.900
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
Fig. 2. Consequences of KGHDC dysfunction for energy metabolism, signaling, and redox homeostasis.
enzymes. One of the best-known lipoate analogues
is the anticancer agent devimistat (CPI-613), which
acts by inhibiting PDHC and KGDHC, thereby induc-
ing metabolic collapse and subsequent death of tu-
mor cells [134-136]. Preclinical studies suggest that
combining CPI-613 with Bcl-xL inhibitors represents
a promising strategy for glioblastoma treatment [137].
siRNA (small interfering RNA) or shRNA (short
hairpin RNA) are used to selectively silence expres-
sion of KGDHC subunits (particularly the KGDH gene)
in order to study how cells rely on KGDHC-dependent
metabolic and signaling pathways. It has been shown
that KGDH-targeting siRNA or shRNA can impair pro-
liferation of KGDH-dependent cancer cell lines, e.g.,
KS1 human glioblastoma cells, by reducing utilization
of aspartate as an alternative energy source [138].
Circ-KGDH (also referred to as circ-OGDH) is
a circular RNA derived by back-splicing of KGDH
gene exons into a covalently closed loop structure.
CircRNAs primarily function as non-coding regulators
of gene expression by acting as microRNA “sponges”
and inhibiting their activity, interacting with
RNA-binding proteins, and modulating transcription-
al processes [139]. Emerging evidence suggests that
circRNAs accumulate mostly in the central nervous
system, where they play an important role in neuro-
nal function and synaptic plasticity [139]. Circ-KGDH
expression is significantly upregulated under hypoxic
conditions, including in the ischemic penumbra in the
middle cerebral artery occlusion (MCAO) mouse mod-
el, plasma of patients with acute ischemic stroke, and
SH-SY5Y cells in the oxygen-glucose deprivation/reox-
ygenation (OGD/R) model, where it aggravates neuro-
nal injury [140-142]. Circ-KGDH interacts with miR-
5112 (microRNA-5112) in primary cortical neurons,
leading to the upregulation of COL4A4 (α-4 chain of
type IV collagen) expression, which induces neuro-
nal injury. Additionally, hypoxia-stressed penumbral
tissue releases small extracellular vesicles containing
circ-KGDH, thus causing remote injury after cortical
ischemia [143, 144]. Consistently, the knockdown of
circ-KGDH increased neuronal cell viability under
ischemic conditions [141, 142]. Therefore, circ-KGDH
is both a promising therapeutic target for improving
neuronal survival and a potential blood-based bio-
marker for predicting the penumbra in patients with
acute ischemic stroke [145].
CONCLUSIONS
In conclusion, KGDHC dysfunction contributes to
neurological diseases by disrupting cellular bioener-
getics, mitochondrial dynamics, redox signaling, and
glutamate metabolism, ultimately leading to cogni-
tive decline and behavioural abnormalities (Fig. 2).
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 901
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
Animal models show that mild KGDHC inhibition or
partial (heterozygous) KGDHC deficiently cause no,
or only subtle, phenotypic changes under baseline
conditions. However, such KGDHC deficiency induces
severe cell vulnerability during metabolic stress. Sev-
eral key questions remain unresolved, such as precise
cell type-specific regulation of KGDHC activity or the
therapeutic window within which pharmacological
modulation of KGDHC is neuroprotective rather than
detrimental. Addressing these gaps will be essential to
better understanding the pathogenesis of major neu-
rological disorders and to identifying rational targets
for therapeutic intervention in neurodegeneration.
Contributions
M. K. and T. K. wrote the text of the review; A. A. and
L. T. edited the manuscript.
Funding
This work was supported by the Hungarian Scientific
Research Fund (OTKA grant 143627, to A.A.) and
National Research, Development and Innova-
tion Fund (TKP2021-EGA-25 grant, to A.A.). Project
no. TKP2021-EGA-25 has been implemented with the
support provided by the Ministry of Innovation and
Technology of Hungary from the National Research,
Development and Innovation Fund, financed under
the TKP2021-EGA funding scheme. This work was also
supported by the Cooperative Translational Research
Program (KTKP) of the Faculty of Medicine at Sem-
melweis University (to T.K.).
Ethics approval and consent to participate
This work does not contain any studies involving hu-
man or animal subjects.
Conflict of interest
The authors of this work declare that they have no
conflicts of interest.
Additional information
The figures were created with images provided by
Servier Medical Art
(https://smart.servier.com), li-
censed under CC BY 4.0.
REFERENCES
1. Fiskum, G. (1985) Mitochondrial damage during cerebral ischemia, Ann. Emerg. Med., 14, 810-815, https://
doi.org/10.1016/s0196-0644(85)80063-9.
2. Chinopoulos, C. (2011) Mitochondrial consumption of cytosolic ATP: not so fast, FEBS Lett., 585, 1255-1259,
https://doi.org/10.1016/j.febslet.2011.04.004.
3. Kiss, G., Konrad, C., Doczi, J., Starkov, A. A., Kawamata, H., Manfredi, G., Zhang, S. F., Gibson, G. E., Beal, M. F.,
Adam-Vizi, V., and Chinopoulos, C. (2013) The negative impact of alpha-ketoglutarate dehydrogenase com-
plex deficiency on matrix substrate-level phosphorylation, FASEB J., 27, 2392-2406, https://doi.org/10.1096/
fj.12-220202.
4. Szabo, E., Nagy, B., Czajlik, A., Komlodi, T., Ozohanics, O., Tretter, L., and Ambrus, A. (2024) Mitochondrial al-
pha-keto acid dehydrogenase complexes: recent developments on structure and function in health and disease,
Subcell Biochem., 104, 295-381, https://doi.org/10.1007/978-3-031-58843-3_13.
5. Hansen, G. E., and Gibson, G. E. (2022) The alpha-ketoglutarate dehydrogenase complex as a hub of plasticity
in neurodegeneration and regeneration, Int. J. Mol. Sci., 23, 12403, https://doi.org/10.3390/ijms232012403.
6. Bunik, V. I. (2019) Redox-driven signaling: 2-oxo acid dehydrogenase complexes as sensors and transmitters of
metabolic imbalance, Antioxid. Redox Signal., 30, 1911-1947, https://doi.org/10.1089/ars.2017.7311.
7. Johnson, M. T., Yang, H. S., Magnuson, T., and Patel, M. S. (1997) Targeted disruption of the murine dihydroli-
poamide dehydrogenase gene (Dld) results in perigastrulation lethality, Proc. Natl. Acad. Sci. USA, 94, 14512-
14517, https://doi.org/10.1073/pnas.94.26.14512.
8. Yang, L., Shi, Q., Ho, D. J., Starkov, A. A., Wille, E. J., Xu, H., Chen, H. L., Zhang, S., Stack, C. M., Calingasan, N. Y.,
Gibson, G. E., and Beal, M. F. (2009) Mice deficient in dihydrolipoyl succinyl transferase show increased vulner-
ability to mitochondrial toxins, Neurobiol. Dis., 36, 320-330, https://doi.org/10.1016/j.nbd.2009.07.023.
9. Amsterdam, A., Nissen, R. M., Sun, Z., Swindell, E. C., Farrington, S., and Hopkins, N. (2004) Identification
of 315 genes essential for early zebrafish development, Proc. Natl. Acad. Sci. USA, 101, 12792-12797, https://
doi.org/10.1073/pnas.0403929101.
10. Fan, Z., Li, L., Li, X., Zhang, M., Zhong, Y., Li, Y., Yu, D., Cao, J., Zhao, J., Xiaoming, D., Zhang, M., Jian-Guo, W.,
Liu, Z., Goscinski, M. A., Berge, V., Nesland, J. M., and Suo, Z. (2018) Generation of an oxoglutarate dehy-
drogenase knockout rat model and the effect of a high-fat diet, RSC Adv., 8, 16636-16644, https://doi.org/
10.1039/c8ra00253c.
11. Yap, Z.Y., Efthymiou,S., Seiffert,S., Vargas Parra,K., Lee,S., Nasca,A., Maroofian,R., Schrauwen,I., Pendziwiat,M.,
Jung, S., Bhoj, E., Striano, P., Mankad, K., Vona, B., Cuddapah, S., Wagner, A., Alvi, J. R., Davoudi-Dehaghani, E.,
Fallah, M. S., Gannavarapu, S., et al. (2021) Bi-allelic variants in OGDHL cause a neurodevelopmental spectrum
KOKAS et al.902
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
disease featuring epilepsy, hearing loss, visual impairment, and ataxia, Am. J. Hum. Genet., 108, 2368-2384,
https://doi.org/10.1016/j.ajhg.2021.11.003.
12. Whittle, E. F., Chilian, M., Karimiani, E. G., Progri, H., Buhas, D., Kose, M., Ganetzky, R. D., Toosi, M. B.,
Torbati, P. N., Badv, R. S., Shelihan, I., Yang, H., Elloumi, H. Z., Lee, S., Jamshidi, Y., Pittman, A. M.,
Houlden, H., Ignatius, E., Rahman, S., Maroofian, R., et al. (2023) Biallelic variants in OGDH encoding ox-
oglutarate dehydrogenase lead to a neurodevelopmental disorder characterized by global developmental de-
lay, movement disorder, and metabolic abnormalities, Genet. Med., 25, 100332, https://doi.org/10.1016/j.gim.
2022.11.001.
13. Szabo, P., Cai, X., Ali, G., and Blass, J. P. (1994) Localization of the gene (OGDH) coding for the E1k component
of the alpha-ketoglutarate dehydrogenase complex to chromosome 7p13-p11.2, Genomics, 20, 324-326, https://
doi.org/10.1006/geno.1994.1178.
14. Larsson, T. A., Olsson, F., Sundstrom, G., Lundin, L. G., Brenner, S., Venkatesh, B., and Larhammar, D. (2008)
Early vertebrate chromosome duplications and the evolution of the neuropeptide Y receptor gene regions, BMC
Evol. Biol., 8, 184, https://doi.org/10.1186/1471-2148-8-184.
15. Bunik, V. I., and Degtyarev, D. (2008) Structure-function relationships in the 2-oxo acid dehydrogenase family:
substrate-specific signatures and functional predictions for the 2-oxoglutarate dehydrogenase-like proteins,
Proteins, 71, 874-890, https://doi.org/10.1002/prot.21766.
16. Yao, L., Xu, X., Xu, Y., Li, C., Xie, F., Guo, M., Liu, Z., and Liu, X. (2022) OGDHL ameliorates cognitive impairment
and Alzheimer’s disease-like pathology via activating Wnt/beta-catenin signaling in Alzheimer’s disease mice,
Behav. Brain Res., 418, 113673, https://doi.org/10.1016/j.bbr.2021.113673.
17. Nemeria, N. S., Gerfen, G., Nareddy, P. R., Yang, L., Zhang, X., Szostak, M., and Jordan, F. (2018) The mitochon-
drial 2-oxoadipate and 2-oxoglutarate dehydrogenase complexes share their E2 and E3 components for their
function and both generate reactive oxygen species, Free Radic. Biol. Med., 115, 136-145, https://doi.org/10.1016/
j.freeradbiomed.2017.11.018.
18. Leandro, J., Dodatko, T., Aten, J., Nemeria, N. S., Zhang, X., Jordan, F., Hendrickson, R. C., Sanchez, R., Yu, C.,
DeVita, R. J., and Houten, S. M. (2020) DHTKD1 and OGDH display substrate overlap in cultured cells and form
a hybrid 2-oxo acid dehydrogenase complex in vivo, Hum. Mol. Genet., 29, 1168-1179, https://doi.org/10.1093/
hmg/ddaa037.
19. Ozohanics, O., Zhang, X., Nemeria, N. S., Ambrus, A., and Jordan, F. (2023) Probing the E1o-E2o and E1a-E2o
interactions in binary subcomplexes of the human 2-oxoglutarate dehydrogenase and 2-oxoadipate dehydroge-
nase complexes by chemical cross-linking mass spectrometry and molecular dynamics simulation, Int. J. Mol.
Sci., 24, 4555, https://doi.org/10.3390/ijms24054555.
20. Biagosch, C., Ediga, R. D., Hensler, S. V., Faerberboeck, M., Kuehn, R., Wurst, W., Meitinger, T., Kolker,S., Sauer, S.,
and Prokisch, H. (2017) Elevated glutaric acid levels in Dhtkd1-/Gcdh- double knockout mice challenge our
current understanding of lysine metabolism, Biochim. Biophys. Acta Mol. Basis Dis., 1863, 2220-2228, https://
doi.org/10.1016/j.bbadis.2017.05.018.
21. Wang, Y., Guo, Y. R., Liu, K., Yin, Z., Liu, R., Xia, Y., Tan, L., Yang, P., Lee, J. H., Li, X. J., Hawke, D., Zheng, Y.,
Qian, X., Lyu, J., He, J., Xing, D., Tao, Y. J., and Lu, Z. (2017) KAT2A coupled with the alpha-KGDH complex acts
as a histone H3 succinyltransferase, Nature, 552, 273-277, https://doi.org/10.1038/nature25003.
22. Li, X., Egervari, G., Wang, Y., Berger, S. L., and Lu, Z. (2018) Regulation of chromatin and gene expression
by metabolic enzymes and metabolites, Nat. Rev. Mol. Cell Biol., 19, 563-578, https://doi.org/10.1038/s41580-
018-0029-7.
23. Smestad, J., Erber, L., Chen, Y., and Maher, L. J., 3rd (2018) Chromatin succinylation correlates with ac-
tive gene expression and is perturbed by defective TCA cycle metabolism, iScience, 2, 63-75, https://doi.org/
10.1016/j.isci.2018.03.012.
24. Pal, M. M. (2021) Glutamate: the master neurotransmitter and its implications in chronic stress and mood dis-
orders, Front. Hum. Neurosci., 15, 722323, https://doi.org/10.3389/fnhum.2021.722323.
25. Meldrum, B.S. (2000) Glutamate as a neurotransmitter in the brain: review of physiology and pathology, J. Nutr.,
130, 1007S-1015S, https://doi.org/10.1093/jn/130.4.1007S.
26. Wu, W. L., Gong, X. X., Qin, Z. H., and Wang, Y. (2025) Molecular mechanisms of excitotoxicity and their rele-
vance to the pathogenesis of neurodegenerative diseases-an update, Acta Pharmacol. Sin., 46, 3129-3142, https://
doi.org/10.1038/s41401-025-01576-w.
27. Almohmadi, N. H., Al-Kuraishy, H. M., Al-Gareeb, A. I., Albuhadily, A. K., Abdelaziz, A. M., Jabir, M. S., Alexiou, A.,
Papadakis, M., and Batiha, G. E. (2025) Glutamatergic dysfunction in neurodegenerative diseases focusing
on Parkinson’s disease: role of glutamate modulators, Brain Res. Bull., 225, 111349, https://doi.org/10.1016/
j.brainresbull.2025.111349.
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 903
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
28. Blass, J. P., and Gibson, G. E. (1991) The role of oxidative abnormalities in the pathophysiology of Alzheimer’s
disease, Rev. Neurol (Paris), 147, 513-525.
29. Weidinger, A., Milivojev, N., Hosmann, A., Duvigneau, J. C., Szabo, C., Toro, G., Rauter, L., Vaglio-Garro, A.,
Mkrtchyan, G. V., Trofimova, L., Sharipov, R. R., Surin, A. M., Krasilnikova, I. A., Pinelis, V. G., Tretter, L.,
Moldzio, R., Bayir, H., Kagan, V. E., Bunik, V. I., and Kozlov, A. V. (2023) Oxoglutarate dehydrogenase complex
controls glutamate-mediated neuronal death, Redox Biol., 62, 102669, https://doi.org/10.1016/j.redox.2023.102669.
30. Trofimova, L. K., Araujo, W. L., Strokina, A. A., Fernie, A. R., Bettendorff, L., and Bunik, V. I. (2012) Con-
sequences of the alpha-ketoglutarate dehydrogenase inhibition for neuronal metabolism and survival:
implications for neurodegenerative diseases, Curr. Med. Chem., 19, 5895-5906, https://doi.org/10.2174/
092986712804143367.
31. Graf, A., Ksenofontov, A., and Bunik, V. (2021) Inhibition of 2-oxoglutarate dehydrogenase as a chemical model
of acute hypobaric hypoxia, Front. Med. (Lausanne), 8, 751639, https://doi.org/10.3389/fmed.2021.751639.
32. Goeschl, V., Hotka, M., Hochreiter, B., Hilber, K., Boehm, S., Kozlov, A. V., and Kubista, H. (2026) alpha-Ketoglu-
tarate dehydrogenase complex activity modulates glutamate excitotoxicity via metabotropic regulation of NMDA
receptors in primary cultures, J. Cell Sci., 139, jcs264420, https://doi.org/10.1242/jcs.264420.
33. Martinez, S., and Hausinger, R. P. (2015) Catalytic mechanisms of Fe(II)- and 2-oxoglutarate-dependent oxygen-
ases, J. Biol. Chem., 290, 20702-20711, https://doi.org/10.1074/jbc.R115.648691.
34. Jaakkola, P., Mole, D. R., Tian, Y. M., Wilson, M. I., Gielbert, J., Gaskell, S. J., von Kriegsheim, A., Hebestreit, H. F.,
Mukherji, M., Schofield, C. J., Maxwell, P. H., Pugh, C. W., and Ratcliffe, P. J. (2001) Targeting of HIF-alpha
to the von Hippel-Lindau ubiquitylation complex by O2-regulated prolyl hydroxylation, Science, 292, 468-472,
https://doi.org/10.1126/science.1059796.
35. Semenza, G. L. (2012) Hypoxia-inducible factors in physiology and medicine, Cell, 148, 399-408, https://
doi.org/10.1016/j.cell.2012.01.021.
36. Chen,H., Denton, T. T., Xu, H., Calingasan, N., Beal, M. F., and Gibson, G.E. (2016) Reductions in the mitochondri-
al enzyme alpha-ketoglutarate dehydrogenase complex in neurodegenerative disease - beneficial or detrimental?
J. Neurochem., 139, 823-838, https://doi.org/10.1111/jnc.13836.
37. Mastrogiacomo, F., LaMarche, J., Dozic, S., Lindsay, G., Bettendorff, L., Robitaille, Y., Schut, L., and Kish, S. J.
(1996) Immunoreactive levels of alpha-ketoglutarate dehydrogenase subunits in Friedreich’s ataxia and spinoc-
erebellar ataxia type 1, Neurodegeneration, 5, 27-33, https://doi.org/10.1006/neur.1996.0004.
38. Mizuno, Y., Matuda, S., Yoshino, H., Mori, H., Hattori, N., and Ikebe, S. (1994) An immunohistochemical study
on alpha-ketoglutarate dehydrogenase complex in Parkinson’s disease, Ann. Neurol., 35, 204-210, https://
doi.org/10.1002/ana.410350212.
39. Gibson, G. E., Kingsbury, A. E., Xu, H., Lindsay, J. G., Daniel, S., Foster, O. J., Lees, A. J., and Blass, J. P. (2003)
Deficits in a tricarboxylic acid cycle enzyme in brains from patients with Parkinson’s disease, Neurochem. Int.,
43, 129-135, https://doi.org/10.1016/s0197-0186(02)00225-5.
40. Johnson,M., Yang, H. S., Johanning, G. L., and Patel, M. S. (1997) Characterization of the mouse dihydrolipoamide
dehydrogenase (Dld) gene: genomic structure, promoter sequence, and chromosomal localization, Genomics, 41,
320-326, https://doi.org/10.1006/geno.1997.4670.
41. Starkov, A. A., Fiskum, G., Chinopoulos, C., Lorenzo, B. J., Browne, S. E., Patel, M. S., and Beal, M. F. (2004)
Mitochondrial alpha-ketoglutarate dehydrogenase complex generates reactive oxygen species, J. Neurosci., 24,
7779-7788, https://doi.org/10.1523/JNEUROSCI.1899-04.2004.
42. Horvath, G., Svab, G., Komlodi, T., Ravasz, D., Kacso, G., Doczi, J., Chinopoulos, C., Ambrus, A., and Tretter, L.
(2022) Reverse and forward electron flow-induced H2O2 formation is decreased in alpha-ketoglutarate dehydro-
genase (alpha-KGDH) subunit (E2 or E3) heterozygote knock out animals, Antioxidants (Basel), 11, 1487, https://
doi.org/10.3390/antiox11081487.
43. Kokas, M., Budai, A., Kadar, A., Mozaffaritabar, S., Zhou, L., Teglas, T., Orova, R. S., Gaspar, D., Nemeth, K.,
Toth, D. M., Sayour, N. V., Kovacshazi, C., Xue, A., Szatmari, R. Z., Torocsik, B., Mathe, D., Kovacs, N., Szigeti, K.,
Nagy, P., Szatmari, I., et al. (2025) Microgliosis, neuronal death, minor behavioral abnormalities and reduced
endurance performance in alpha-ketoglutarate dehydrogenase complex deficient mice, Redox Biol., 85, 103743,
https://doi.org/10.1016/j.redox.2025.103743.
44. Broxton, C. N., Kaur, P., Lavorato, M., Ganesh, S., Xiao, R., Mathew, N. D., Nakamaru-Ogiso, E., Anderson, V. E.,
and Falk, M. J. (2022) Dichloroacetate and thiamine improve survival and mitochondrial stress in a C. ele-
gans model of dihydrolipoamide dehydrogenase deficiency, JCI Insight, 7, e156222, https://doi.org/10.1172/jci.
insight.156222.
45. Lavorato, M., Iadarola, D., Remes, C., Kaur, P., Broxton, C., Mathew, N. D., Xiao, R., Seiler, C., Nakamaru-Ogiso, E.,
Anderson, V. E., and Falk, M.J. (2024) dldh
cri3
zebrafish exhibit altered mitochondrial ultrastructure, morphology,
KOKAS et al.904
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
and dysfunction partially rescued by probucol or thiamine, JCI Insight, 9, e178973, https://doi.org/10.1172/jci.
insight.178973.
46. Bunik, V. I., Artiukhov, A. V., Kazantsev, A. V., Aleshin, V. A., Boyko, A. I., Ksenofontov, A. L., Lukashev, N.V., and
Graf, A. V. (2022) Administration of phosphonate inhibitors of dehydrogenases of 2-oxoglutarate and 2-oxoad-
ipate to rats elicits target-specific metabolic and physiological responses, Front. Chem., 10, 892284, https://
doi.org/10.3389/fchem.2022.892284.
47. Graf, A., Trofimova, L., Loshinskaja, A., Mkrtchyan, G., Strokina, A., Lovat, M., Tylicky, A., Strumilo, S.,
Bettendorff, L., and Bunik, V. I. (2013) Up-regulation of 2-oxoglutarate dehydrogenase as a stress response, Int.
J. Biochem. Cell Biol., 45, 175-189, https://doi.org/10.1016/j.biocel.2012.07.002.
48. Van de Vondel, L., Lee, G. S., De Winter, J., Matsuzaki, S., Sandoval, A., Ramirez, J. F., Monticelli, A., Lee, S.,
Horvath, R., de Bleecker, J., Zuchner, S., Humphries, K. M., Baets, J., and Yoon, W. H. (2026) Heterozygous OGDH
variants are involved in peripheral neuropathy with ataxia and optical atrophy, JIMD Rep., 67, e70103, https://
doi.org/10.1002/jmd2.70103.
49. Berndt, N., Holzhutter, H. G., and Bulik, S. (2013) Implications of enzyme deficiencies on mitochondrial ener-
gy metabolism and reactive oxygen species formation of neurons involved in rotenone-induced Parkinson’s
disease: a model-based analysis, FEBS J., 280, 5080-5093, https://doi.org/10.1111/febs.12480.
50. Du, H., Guo, L., and Yan, S. S. (2012) Synaptic mitochondrial pathology in Alzheimer’s disease, Antioxid Redox
Signal, 16, 1467-1475, https://doi.org/10.1089/ars.2011.4277.
51. Nilsen, L. H., Shi, Q., Gibson, G. E., and Sonnewald, U. (2011) Brain [U-13 C]glucose metabolism in mice
with decreased alpha-ketoglutarate dehydrogenase complex activity, J.Neurosci. Res., 89, 1997-2007, https://
doi.org/10.1002/jnr.22606.
52. Remacha, L., Pirman, D., Mahoney, C. E., Coloma, J., Calsina, B., Curras-Freixes, M., Leton, R., Torres-Perez, R.,
Richter, S., Pita, G., Herraez, B., Cianchetta, G., Honrado, E., Maestre, L., Urioste, M., Aller, J., Garcia-Uriarte, O.,
Galvez, M. A., Luque, R. M., Lahera, M., et al. (2019) Recurrent germline DLST mutations in individuals
with multiple pheochromocytomas and paragangliomas, Am. J. Hum. Genet., 104, 651-664, https://doi.org/
10.1016/j.ajhg.2019.02.017.
53. Chinopoulos, C. (2011) The “B space” of mitochondrial phosphorylation, J.Neurosci. Res., 89, 1897-1904, https://
doi.org/10.1002/jnr.22659.
54. Lin, S. C., and Hardie, D. G. (2018) AMPK: sensing glucose as well as cellular energy status, Cell Metab, 27,
299-313, https://doi.org/10.1016/j.cmet.2017.10.009.
55. Steinberg, G. R., and Hardie, D. G. (2023) New insights into activation and function of the AMPK, Nat. Rev. Mol.
Cell Biol., 24, 255-272, https://doi.org/10.1038/s41580-022-00547-x.
56. Hawley, S. A., Ross, F. A., Chevtzoff, C., Green, K. A., Evans, A., Fogarty, S., Towler, M. C., Brown, L. J.,
Ogunbayo, O. A., Evans, A. M., and Hardie, D. G. (2010) Use of cells expressing gamma subunit variants
to identify diverse mechanisms of AMPK activation, Cell Metab., 11, 554-565, https://doi.org/10.1016/j.cmet.
2010.04.001.
57. Klivenyi, P., Starkov, A. A., Calingasan, N. Y., Gardian, G., Browne, S. E., Yang, L., Bubber, P., Gibson, G. E.,
Patel, M. S., and Beal, M. F. (2004) Mice deficient in dihydrolipoamide dehydrogenase show increased vul-
nerability to MPTP, malonate and 3-nitropropionic acid neurotoxicity, J. Neurochem., 88, 1352-1360, https://
doi.org/10.1046/j.1471-4159.2003.02263.x.
58. Dumont, M., Ho, D. J., Calingasan, N. Y., Xu, H., Gibson, G., and Beal, M. F. (2009) Mitochondrial dihydroli-
poyl succinyltransferase deficiency accelerates amyloid pathology and memory deficit in a transgenic mouse
model of amyloid deposition, Free Radic. Biol. Med., 47, 1019-1027, https://doi.org/10.1016/j.freeradbiomed.
2009.07.008.
59. Kanamori, T., Nishimaki, K., Asoh, S., Ishibashi, Y., Takata, I., Kuwabara, T., Taira,K., Yamaguchi, H., Sugihara, S.,
Yamazaki, T., Ihara, Y., Nakano, K., Matuda, S., and Ohta, S. (2003) Truncated product of the bifunctional DLST
gene involved in biogenesis of the respiratory chain, EMBOJ., 22, 2913-2923, https://doi.org/10.1093/emboj/cdg299.
60. Sies, H. (2021) Oxidative eustress: on constant alert for redox homeostasis, Redox Biol., 41, 101867, https://
doi.org/10.1016/j.redox.2021.101867.
61. Sies, H., and Jones, D. P. (2020) Reactive oxygen species (ROS) as pleiotropic physiological signaling agents, Nat.
Rev. Mol. Cell Biol., 21, 363-383, https://doi.org/10.1038/s41580-020-0230-3.
62. Sies, H., Belousov, V. V., Chandel, N. S., Davies, M. J., Jones, D. P., Mann, G. E., Murphy, M. P., Yamamoto, M., and
Winterbourn, C. (2022) Defining roles of specific reactive oxygen species (ROS) in cell biology and physiology,
Nat. Rev. Mol. Cell Biol., 23, 499-515, https://doi.org/10.1038/s41580-022-00456-z.
63. Reczek, C. R., and Chandel, N. S. (2015) ROS-dependent signal transduction, Curr. Opin. Cell Biol., 33, 8-13, https://
doi.org/10.1016/j.ceb.2014.09.010.
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 905
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
64. Brand, M. D. (2016) Mitochondrial generation of superoxide and hydrogen peroxide as the source of mitochon-
drial redox signaling, Free Radic. Biol. Med., 100, 14-31, https://doi.org/10.1016/j.freeradbiomed.2016.04.001.
65. Andreyev, A. Y., Kushnareva, Y. E., Murphy, A. N., and Starkov, A. A. (2015) Mitochondrial ROS metabolism:
10 years later, Biochemistry (Moscow), 80, 517-531, https://doi.org/10.1134/S0006297915050028.
66. Andreyev, A. Y., Kushnareva, Y. E., and Starkov, A. A. (2005) Mitochondrial metabolism of reactive oxygen spe-
cies, Biochemistry (Moscow), 70, 200-214, https://doi.org/10.1007/s10541-005-0102-7.
67. Gorlach, A., Dimova, E. Y., Petry, A., Martinez-Ruiz, A., Hernansanz-Agustin, P., Rolo, A. P., Palmeira, C. M., and
Kietzmann, T. (2015) Reactive oxygen species, nutrition, hypoxia and diseases: problems solved? Redox Biol., 6,
372-385, https://doi.org/10.1016/j.redox.2015.08.016.
68. Tretter, L., and Adam-Vizi, V. (2004) Generation of reactive oxygen species in the reaction catalyzed by alpha-
ketoglutarate dehydrogenase, J. Neurosci., 24, 7771-7778, https://doi.org/10.1523/JNEUROSCI.1842-04.2004.
69. Quinlan, C. L., Goncalves, R. L., Hey-Mogensen, M., Yadava, N., Bunik, V. I., and Brand, M. D. (2014) The 2-oxoac-
id dehydrogenase complexes in mitochondria can produce superoxide/hydrogen peroxide at much higher rates
than complex I, J.Biol. Chem., 289, 8312-8325, https://doi.org/10.1074/jbc.M113.545301.
70. Ambrus, A., Tretter, L., and Adam-Vizi, V. (2009) Inhibition of the alpha-ketoglutarate dehydrogenase-medi-
ated reactive oxygen species generation by lipoic acid, J. Neurochem., 109, 222-229, https://doi.org/10.1111/
j.1471-4159.2009.05942.x.
71. Tretter, L., and Adam-Vizi, V. (2005) Alpha-ketoglutarate dehydrogenase: a target and generator of oxidative
stress, Philos. Trans. R Soc. Lond. B Biol. Sci., 360, 2335-2345, https://doi.org/10.1098/rstb.2005.1764.
72. Bunik, V. I., and Sievers, C. (2002) Inactivation of the 2-oxo acid dehydrogenase complexes upon generation of
intrinsic radical species, Eur. J. Biochem., 269, 5004-5015, https://doi.org/10.1046/j.1432-1033.2002.03204.x.
73. Humphries, K. M., and Szweda, L. I. (1998) Selective inactivation of alpha-ketoglutarate dehydrogenase and
pyruvate dehydrogenase: reaction of lipoic acid with 4-hydroxy-2-nonenal, Biochemistry, 37, 15835-15841,
https://doi.org/10.1021/bi981512h.
74. McLain, A. L., Szweda, P. A., and Szweda, L. I. (2011) alpha-Ketoglutarate dehydrogenase: a mitochondrial redox
sensor, Free Radic. Res., 45, 29-36, https://doi.org/10.3109/10715762.2010.534163.
75. Seim, G. L., John, S. V., Arp, N. L., Fang, Z., Pagliarini, D. J., and Fan, J. (2023) Nitric oxide-driven modifica-
tions of lipoic arm inhibit alpha-ketoacid dehydrogenases, Nat. Chem. Biol., 19, 265-274, https://doi.org/10.1038/
s41589-022-01153-w.
76. McLain, A. L., Cormier, P. J., Kinter, M., and Szweda, L. I. (2013) Glutathionylation of alpha-ketoglutarate dehy-
drogenase: the chemical nature and relative susceptibility of the cofactor lipoic acid to modification, Free Radic.
Biol. Med., 61, 161-169, https://doi.org/10.1016/j.freeradbiomed.2013.03.020.
77. Shi, Q., Xu, H., Kleinman, W. A., and Gibson, G. E. (2008) Novel functions of the alpha-ketoglutarate dehy-
drogenase complex may mediate diverse oxidant-induced changes in mitochondrial enzymes associated
with Alzheimer’s disease, Biochim. Biophys. Acta, 1782, 229-238, https://doi.org/10.1016/j.bbadis.2007.12.008.
78. Mailloux, R. J. (2024) The emerging importance of the alpha-keto acid dehydrogenase complexes in serv-
ing as intracellular and intercellular signaling platforms for the regulation of metabolism, Redox Biol., 72,
103155, https://doi.org/10.1016/j.redox.2024.103155.
79. Grayson, C., Chalifoux, O., Russo, M. S. T., Avizonis, D. Z., Sterman, S., Faerman, B., Koufos, O., Agellon, L. B.,
and Mailloux, R. J. (2024) Ablating the glutaredoxin-2 (Glrx2) gene protects male mice against non-alcoholic
fatty liver disease (NAFLD) by limiting oxidative distress, Free Radic. Biol. Med., 224, 660-677, https://doi.org/
10.1016/j.freeradbiomed.2024.09.016.
80. Applegate, M. A., Humphries, K. M., and Szweda, L. I. (2008) Reversible inhibition of alpha-ketoglutarate de-
hydrogenase by hydrogen peroxide: glutathionylation and protection of lipoic acid, Biochemistry, 47, 473-478,
https://doi.org/10.1021/bi7017464.
81. Shi, Q., Xu, H., Yu, H., Zhang, N., Ye, Y., Estevez, A. G., Deng, H., and Gibson, G. E. (2011) Inactivation and
reactivation of the mitochondrial alpha-ketoglutarate dehydrogenase complex, J. Biol. Chem., 286, 17640-17648,
https://doi.org/10.1074/jbc.M110.203018.
82. Nulton-Persson, A. C., Starke, D. W., Mieyal, J. J., and Szweda, L. I. (2003) Reversible inactivation of alpha-ke-
toglutarate dehydrogenase in response to alterations in the mitochondrial glutathione status, Biochemistry, 42,
4235-4242, https://doi.org/10.1021/bi027370f.
83. Zundorf, G., Kahlert, S., Bunik, V. I., and Reiser, G. (2009) alpha-Ketoglutarate dehydrogenase contributes to
production of reactive oxygen species in glutamate-stimulated hippocampal neurons in situ, Neuroscience, 158,
610-616, https://doi.org/10.1016/j.neuroscience.2008.10.015.
84. Chouchani, E. T., Pell, V. R., Gaude, E., Aksentijevic, D., Sundier, S. Y., Robb, E. L., Logan, A., Nadtochiy, S. M.,
Ord, E. N. J., Smith, A. C., Eyassu, F., Shirley, R., Hu, C. H., Dare, A. J., James, A. M., Rogatti, S., Hartley, R. C.,
KOKAS et al.906
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
Eaton, S., Costa, A. S. H., Brookes, P. S., et al. (2014) Ischaemic accumulation of succinate controls reperfusion
injury through mitochondrial ROS, Nature, 515, 431-435, https://doi.org/10.1038/nature13909.
85. Zhang, J., Wang, Y. T., Miller, J. H., Day, M. M., Munger, J. C., and Brookes, P. S. (2018) Accumulation of succi-
nate in cardiac ischemia primarily occurs via canonical krebs cycle activity, Cell Rep., 23, 2617-2628, https://
doi.org/10.1016/j.celrep.2018.04.104.
86. Calingasan, N. Y., Ho, D. J., Wille, E. J., Campagna, M. V., Ruan, J., Dumont, M., Yang, L., Shi, Q., Gibson, G. E.,
and Beal, M. F. (2008) Influence of mitochondrial enzyme deficiency on adult neurogenesis in mouse models
of neurodegenerative diseases, Neuroscience, 153, 986-996, https://doi.org/10.1016/j.neuroscience.2008.02.071.
87. Bindokas, V. P., Jordan, J., Lee, C. C., and Miller, R. J. (1996) Superoxide production in rat hippocampal neu-
rons: selective imaging with hydroethidine, J. Neurosci., 16, 1324-1336, https://doi.org/10.1523/JNEUROSCI.
16-04-01324.1996.
88. Katsuki, H., Nakanishi, C., Saito, H., and Matsuki, N. (1997) Biphasic effect of hydrogen peroxide on
field potentials in rat hippocampal slices, Eur. J. Pharmacol., 337, 213-218, https://doi.org/10.1016/s0014-
2999(97)01323-x.
89. Klann, E., Roberson, E. D., Knapp, L. T., and Sweatt, J. D. (1998) A role for superoxide in protein kinase C
activation and induction of long-term potentiation, J. Biol. Chem., 273, 4516-4522, https://doi.org/10.1074/jbc.
273.8.4516.
90. Klann, E. (1998) Cell-permeable scavengers of superoxide prevent long-term potentiation in hippocampal area
CA1, J. Neurophysiol., 80, 452-457, https://doi.org/10.1152/jn.1998.80.1.452.
91. Thiels, E., Urban, N. N., Gonzalez-Burgos, G. R., Kanterewicz, B. I., Barrionuevo, G., Chu, C. T., Oury, T. D., and
Klann, E. (2000) Impairment of long-term potentiation and associative memory in mice that overexpress extra-
cellular superoxide dismutase, J. Neurosci., 20, 7631-7639, https://doi.org/10.1523/JNEUROSCI.20-20-07631.2000.
92. Viggiano, A., Seru, R., Damiano, S., De Luca, B., Santillo, M., and Mondola, P. (2012) Inhibition of long-term po-
tentiation by CuZn superoxide dismutase injection in rat dentate gyrus: involvement of muscarinic M1 receptor,
J. Cell Physiol., 227, 3111-3115, https://doi.org/10.1002/jcp.23062.
93. Aquilano, K., Baldelli, S., Pagliei, B., Cannata, S. M., Rotilio, G., and Ciriolo, M. R. (2013) p53 orchestrates the
PGC-1alpha-mediated antioxidant response upon mild redox and metabolic imbalance, Antioxid. Redox Signal.,
18, 386-399, https://doi.org/10.1089/ars.2012.4615.
94. Aquilano, K., Vigilanza, P., Rotilio, G., and Ciriolo, M. R. (2006) Mitochondrial damage due to SOD1 deficien-
cy in SH-SY5Y neuroblastoma cells: a rationale for the redundancy of SOD1, FASEB J., 20, 1683-1685, https://
doi.org/10.1096/fj.05-5225fje.
95. Rojo, A. I., Pajares, M., Rada, P., Nunez, A., Nevado-Holgado, A. J., Killik, R., Van Leuven, F., Ribe, E.,
Lovestone, S., Yamamoto, M., and Cuadrado, A. (2017) NRF2 deficiency replicates transcriptomic changes in
Alzheimer’s patients and worsens APP and TAU pathology, Redox Biol., 13, 444-451, https://doi.org/10.1016/j.redox.
2017.07.006.
96. Singulani, M. P., Pereira, C. P. M., Ferreira, A. F. F., Garcia, P. C., Ferrari, G. D., Alberici, L. C., and Britto, L. R.
(2020) Impairment of PGC-1alpha-mediated mitochondrial biogenesis precedes mitochondrial dysfunction and
Alzheimer’s pathology in the 3xTg mouse model of Alzheimer’s disease, Exp. Gerontol., 133, 110882, https://
doi.org/10.1016/j.exger.2020.110882.
97. Banerjee, K., Munshi, S., Xu, H., Frank, D. E., Chen, H. L., Chu, C. T., Yang, J., Cho, S., Kagan, V. E., Denton, T. T.,
Tyurina, Y. Y., Jiang, J. F., and Gibson, G. E. (2016) Mild mitochondrial metabolic deficits by alpha-ketoglutarate
dehydrogenase inhibition cause prominent changes in intracellular autophagic signaling: potential role in the
pathobiology of Alzheimer’s disease, Neurochem. Int., 96, 32-45, https://doi.org/10.1016/j.neuint.2016.02.011.
98. Manczak, M., Calkins, M. J., and Reddy, P. H. (2011) Impaired mitochondrial dynamics and abnormal interaction
of amyloid beta with mitochondrial protein Drp1 in neurons from patients with Alzheimer’s disease: implications
for neuronal damage, Hum. Mol. Genet., 20, 2495-2509, https://doi.org/10.1093/hmg/ddr139.
99. Frank, M., Duvezin-Caubet, S., Koob, S., Occhipinti, A., Jagasia, R., Petcherski, A., Ruonala, M. O., Priault, M.,
Salin, B., and Reichert, A. S. (2012) Mitophagy is triggered by mild oxidative stress in a mitochondrial fission
dependent manner, Biochim. Biophys. Acta, 1823, 2297-2310, https://doi.org/10.1016/j.bbamcr.2012.08.007.
100. Youle, R. J., and van der Bliek, A. M. (2012) Mitochondrial fission, fusion, and stress, Science, 337, 1062-1065,
https://doi.org/10.1126/science.1219855.
101. Toyama, E.Q., Herzig, S., Courchet, J., Lewis, T. L., Jr., Loson, O.C., Hellberg,K., Young, N. P., Chen, H., Polleux, F.,
Chan, D. C., and Shaw, R. J. (2016) Metabolism. AMP-activated protein kinase mediates mitochondrial fission in
response to energy stress, Science, 351, 275-281, https://doi.org/10.1126/science.aab4138.
102. Zhang, Z., Tan, M., Xie, Z., Dai, L., Chen, Y., and Zhao, Y. (2011) Identification of lysine succinylation as a new
post-translational modification, Nat. Chem. Biol., 7, 58-63, https://doi.org/10.1038/nchembio.495.
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 907
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
103. Chen, H., Xu, H., Potash, S., Starkov, A., Belousov, V. V., Bilan, D. S., Denton, T. T., and Gibson, G. E. (2017) Mild
metabolic perturbations alter succinylation of mitochondrial proteins, J. Neurosci. Res., 95, 2244-2252, https://
doi.org/10.1002/jnr.24103.
104. Hou, X., Chen, Y., Li, X., Gu, X., Dong, W., Shi, J., and Ji, S. (2024) Protein succinylation: regulating metabolism
and beyond, Front. Nutr., 11, 1336057, https://doi.org/10.3389/fnut.2024.1336057.
105. Gibson, G. E., Xu, H., Chen, H. L., Chen, W., Denton, T. T., and Zhang, S. (2015) Alpha-ketoglutarate dehy-
drogenase complex-dependent succinylation of proteins in neurons and neuronal cell lines, J. Neurochem.,
134, 86-96, https://doi.org/10.1111/jnc.13096.
106. Yang, Y., Tapias, V., Acosta, D., Xu, H., Chen, H., Bhawal, R., Anderson, E. T., Ivanova, E., Lin, H., Sagdullaev, B. T.,
Chen, J., Klein, W. L., Viola, K. L., Gandy, S., Haroutunian, V., Beal, M. F., Eliezer, D., Zhang, S., and Gibson, G. E.
(2022) Altered succinylation of mitochondrial proteins, APP and tau in Alzheimer’s disease, Nat. Commun., 13,
159, https://doi.org/10.1038/s41467-021-27572-2.
107. Piroli, G. G., Manuel, A. M., McCain, R. S., Smith, H. H., Ozohanics, O., Mellid, S., Cox, J. H., Cotham, W. E.,
Walla, M. D., Cascon, A., Ambrus, A., and Frizzell, N. (2023) Defective function of alpha-ketoglutarate dehy-
drogenase exacerbates mitochondrial ATP deficits during complex I deficiency, Redox Biol., 67, 102932, https://
doi.org/10.1016/j.redox.2023.102932.
108. Alderson, N. L., Wang, Y., Blatnik, M., Frizzell, N., Walla, M. D., Lyons, T. J., Alt, N., Carson, J. A., Nagai, R.,
Thorpe, S. R., and Baynes, J. W. (2006) S-(2-Succinyl)cysteine: a novel chemical modification of tissue proteins
by a Krebs cycle intermediate, Arch. Biochem. Biophys., 450, 1-8, https://doi.org/10.1016/j.abb.2006.03.005.
109. Andreassen, O. A., Dedeoglu, A., Friedlich, A., Ferrante, K. L., Hughes, D., Szabo, C., and Beal, M. F. (2001) Effects
of an inhibitor of poly(ADP-ribose) polymerase, desmethylselegiline, trientine, and lipoic acid in transgenic ALS
mice, Exp. Neurol., 168, 419-424, https://doi.org/10.1006/exnr.2001.7633.
110. Andreassen, O. A., Ferrante, R. J., Dedeoglu, A., and Beal, M. F. (2001) Lipoic acid improves survival in trans-
genic mouse models of Huntington’s disease, Neuroreport, 12, 3371-3373, https://doi.org/10.1097/00001756-
200110290-00044.
111. Rodriguez, M. C., MacDonald, J. R., Mahoney, D. J., Parise, G., Beal, M. F., and Tarnopolsky, M. A. (2007) Bene-
ficial effects of creatine, CoQ10, and lipoic acid in mitochondrial disorders, Muscle Nerve, 35, 235-242, https://
doi.org/10.1002/mus.20688.
112. Fan, P., Liu, Y., Liu, C., Yao, H., Xu, S., Jiang, Y., Wu, Y., Liu, Y., and Guo, X. (2025) NEDD4L induces mi-
tochondrial dysfunction and neurodegeneration by promoting LIPT2 degradation in Huntington’s disease,
Proc. Natl. Acad. Sci. USA, 122, e2503342122, https://doi.org/10.1073/pnas.2503342122.
113. Biryukov, A. I., Bunik, V. I., Zhukov, Y. N., Khurs, E. N., and Khomutov, R. M. (1996) Succinyl phosphonate in-
hibits alpha-ketoglutarate oxidative decarboxylation, catalyzed by alpha-ketoglutarate dehydrogenase complexes
from E. coli and pigeon breast muscle, FEBS Lett., 382, 167-170, https://doi.org/10.1016/0014-5793(96)00166-4.
114. Bunik, V. I., Denton, T. T., Xu, H., Thompson, C. M., Cooper, A. J., and Gibson, G. E. (2005) Phosphonate analogues
of alpha-ketoglutarate inhibit the activity of the alpha-ketoglutarate dehydrogenase complex isolated from brain
and in cultured cells, Biochemistry, 44, 10552-10561, https://doi.org/10.1021/bi0503100.
115. Bunik, V. I., Mkrtchyan, G., Grabarska, A., Oppermann, H., Daloso, D., Araujo, W. L., Juszczak, M., Rzeski, W.,
Bettendorff, L., Fernie, A. R., Meixensberger, J., Stepulak, A., and Gaunitz, F. (2016) Inhibition of mitochondrial
2-oxoglutarate dehydrogenase impairs viability of cancer cells in a cell-specific metabolism-dependent manner,
Oncotarget, 7, 26400-26421, https://doi.org/10.18632/oncotarget.8387.
116. Bunik, V. I., Kabysheva, M. S., Klimuk, E. I., Storozhevykh, T. P., and Pinelis, V. G. (2009) Phosphono ana-
logues of 2-oxoglutarate protect cerebellar granule neurons upon glutamate excitotoxicity, Ann. N Y Acad. Sci.,
1171, 521-529, https://doi.org/10.1111/j.1749-6632.2009.04709.x.
117. Kabysheva, M. S., Storozhevykh, T. P., Pinelis, V. G., and Bunik, V. I. (2009) Synthetic regulators of the 2-oxoglu-
tarate oxidative decarboxylation alleviate the glutamate excitotoxicity in cerebellar granule neurons, Biochem.
Pharmacol., 77, 1531-1540, https://doi.org/10.1016/j.bcp.2009.02.001.
118. Diaz-Munoz, M. D., Bell, S. E., Fairfax, K., Monzon-Casanova, E., Cunningham, A. F., Gonzalez-Porta, M.,
Andrews,S.R., Bunik, V.I., Zarnack,K., Curk,T., Heggermont, W.A., Heymans,S., Gibson, G.E., Kontoyiannis, D.L.,
Ule, J., and Turner, M. (2015) The RNA-binding protein HuR is essential for the B cell antibody response, Nat.
Immunol., 16, 415-425, https://doi.org/10.1038/ni.3115.
119. Bunik, V. I., Tylicki, A., and Lukashev, N. V. (2013) Thiamin diphosphate-dependent enzymes: from enzymol-
ogy to metabolic regulation, drug design and disease models, FEBS J., 280, 6412-6442, https://doi.org/10.1111/
febs.12512.
120. Barclay, L. L., Gibson, G. E., and Blass, J. P. (1982) Cholinergic therapy of abnormal open-field behavior in thi-
amin-deficient rats, J. Nutr., 112, 1906-1913, https://doi.org/10.1093/jn/112.10.1906.
KOKAS et al.908
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
121. Barclay, L. L., Gibson, G. E., and Blass, J. P. (1981) The string test: an early behavioral change in thiamine de-
ficiency, Pharmacol. Biochem. Behav., 14, 153-157, https://doi.org/10.1016/0091-3057(81)90236-7.
122. Butterworth, R. F. (1993) Pathophysiology of cerebellar dysfunction in the Wernicke-Korsakoff syndrome, Can.
J. Neurol. Sci., 20, S123-S126, https://doi.org/10.1017/S0317167100048630.
123. Butterworth, R. F., Kril, J. J., and Harper, C. G. (1993) Thiamine-dependent enzyme changes in the brains of
alcoholics: relationship to the Wernicke-Korsakoff syndrome, Alcohol. Clin. Exp. Res., 17, 1084-1088, https://
doi.org/10.1111/j.1530-0277.1993.tb05668.x.
124. Butterworth, R. F. (1989) Effects of thiamine deficiency on brain metabolism: implications for the pathogen-
esis of the Wernicke-Korsakoff syndrome, Alcohol Alcohol., 24, 271-279, https://doi.org/10.1093/oxfordjournals.
alcalc.a044913.
125. Bubber, P., Ke, Z. J., and Gibson, G. E. (2004) Tricarboxylic acid cycle enzymes following thiamine deficiency,
Neurochem. Int., 45, 1021-1028, https://doi.org/10.1016/j.neuint.2004.05.007.
126. Park, L. C., Calingasan, N. Y., Uchida, K., Zhang, H., and Gibson, G. E. (2000) Metabolic impairment elicits brain
cell type-selective changes in oxidative stress and cell death in culture, J. Neurochem., 74, 114-124, https://
doi.org/10.1046/j.1471-4159.2000.0740114.x.
127. Gibson, G. E., Hirsch, J. A., Fonzetti, P., Jordan, B. D., Cirio, R. T., and Elder, J. (2016) Vitamin B1 (thiamine) and
dementia, Ann. N Y Acad. Sci., 1367, 21-30, https://doi.org/10.1111/nyas.13031.
128. Shi, Q., Karuppagounder, S. S., Xu, H., Pechman, D., Chen, H., and Gibson, G. E. (2007) Responses of the mito-
chondrial alpha-ketoglutarate dehydrogenase complex to thiamine deficiency may contribute to regional selective
vulnerability, Neurochem. Int., 50, 921-931, https://doi.org/10.1016/j.neuint.2007.03.010.
129. Hazell, A. S., Todd, K. G., and Butterworth, R. F. (1998) Mechanisms of neuronal cell death in Wernicke’s en-
cephalopathy, Metab. Brain Dis., 13, 97-122, https://doi.org/10.1023/a:1020657129593.
130. Hazell, A. S., and Butterworth, R. F. (2009) Update of cell damage mechanisms in thiamine deficiency:
focus on oxidative stress, excitotoxicity and inflammation, Alcohol Alcohol., 44, 141-147, https://doi.org/10.1093/
alcalc/agn120.
131. Calingasan, N. Y., Baker, H., Sheu, K. F., and Gibson, G. E. (1995) Blood-brain barrier abnormalities in vulnerable
brain regions during thiamine deficiency, Exp. Neurol., 134, 64-72, https://doi.org/10.1006/exnr.1995.1037.
132. Smith, T. J., Johnson, C. R., Koshy, R., Hess, S. Y., Qureshi, U. A., Mynak, M. L., and Fischer, P. R. (2021) Thi-
amine deficiency disorders: a clinical perspective, Ann. N Y Acad. Sci., 1498, 9-28, https://doi.org/10.1111/
nyas.14536.
133. Calingasan, N. Y., Gandy, S. E., Baker, H., Sheu, K. F., Kim, K. S., Wisniewski, H. M., and Gibson, G. E. (1995)
Accumulation of amyloid precursor protein-like immunoreactivity in rat brain in response to thiamine deficien-
cy, Brain Res., 677, 50-60, https://doi.org/10.1016/0006-8993(95)00136-e.
134. Bingham, P. M., Stuart, S. D., and Zachar, Z. (2014) Lipoic acid and lipoic acid analogs in cancer metabolism
and chemotherapy, Exp. Rev. Clin. Pharmacol., 7, 837-846, https://doi.org/10.1586/17512433.2014.966816.
135. Dorsam, B., and Fahrer, J. (2016) The disulfide compound alpha-lipoic acid and its derivatives: A novel
class of anticancer agents targeting mitochondria, Cancer Lett., 371, 12-19, https://doi.org/10.1016/j.canlet.
2015.11.019.
136. Reddy, V. B., Boteju, L., Boteju, A., Shen, L., Kassahun, K., Reddy, N., Sheldon, A., Luther, S., and Hu, K. (2022)
In vitro and in vivo metabolism of a novel antimitochondrial cancer metabolism agent, CPI-613, in rat and
human, Drug Metab. Dispos., 50, 361-373, https://doi.org/10.1124/dmd.121.000726.
137. Nguyen, T. T., Torrini, C., Shang, E., Shu, C., Mun, J. Y., Gao, Q., Humala, N., Akman, H. O., Zhang, G.,
Westhoff, M. A., Karpel-Massler, G., Bruce, J. N., Canoll, P., and Siegelin, M. D. (2024) OGDH and Bcl-xL loss
causes synthetic lethality in glioblastoma, JCI Insight, 9, e172565, https://doi.org/10.1172/jci.insight.172565.
138. Allen, E. L., Ulanet, D. B., Pirman, D., Mahoney, C. E., Coco, J., Si, Y., Chen, Y., Huang, L., Ren, J., Choe, S.,
Clasquin, M. F., Artin, E., Fan, Z. P., Cianchetta, G., Murtie, J., Dorsch, M., Jin, S., and Smolen, G. A. (2016)
Differential aspartate usage identifies a subset of cancer cells particularly dependent on OGDH, Cell Rep, 17,
876-890, https://doi.org/10.1016/j.celrep.2016.09.052.
139. Hanan, M., Soreq, H., and Kadener, S. (2017) CircRNAs in the brain, RNA Biol., 14, 1028-1034, https://doi.org/10
.1080/15476286.2016.1255398.
140. Zheng, R., Huang, Z., Zhou, H., and Zhang, L. (2026) CircOGDH promotes oxygen-glucose deprivation/reoxygen-
ation-induced neuron injury by targeting the miR-195-5p/brain-specific angiogenesis inhibitor 1-associated protein
2 axis, Neuroreport, 37, 277-285, https://doi.org/10.1097/WNR.0000000000002261.
141. Liu, Y., Li, Y., Zang, J., Zhang, T., Li, Y., Tan, Z., Ma, D., Zhang, T., Wang, S., Zhang, Y., Huang, L., Wu, Y., Su, X.,
Weng, Z., Deng, D., Tsang, C. K., Xu, A., and Lu, D. (2022) CircOGDH is a penumbra biomarker and therapeutic
target in acute ischemic stroke, Circ. Res., 130, 907-924, https://doi.org/10.1161/CIRCRESAHA.121.319412.
INTERPLAY BETWEEN NEURONAL METABOLISM AND SIGNALING 909
BIOCHEMISTRY (MOSCOW) Vol. 91 No. 6 2026
142. Liu, Y., Li, Y., Zang, J., Zhang, T., Li, Y., Tan, Z., Ma, D., Zhang, T., Wang, S., Zhang, Y., Huang, L., Wu, Y., Su, X.,
Weng, Z., Deng, D., Tsang, C. K., Xu, A., and Lu, D. (2022) Correction to: CircOGDH Is a penumbra biomarker
and therapeutic target in acute ischemic stroke, Circ. Res., 130, e24, https://doi.org/10.1161/RES.0000000000000548.
143. Deng, D., Wu, Y., Wu, P., Tan, N., Li, X., Su, X., Chen, W., Zhang, X., Mai, H., Xu, A., and Lu, D. (2026) Penum-
bra-derived small extracellular vesicles carry pathogenic RNA driving remote damage after cortical infarction,
Brain, https://doi.org/10.1093/brain/awag049.
144. Chen, W., Wu, Y., Liang, Y., Su, X., Ke, M., Deng, D., Zang, J., Zhu, J., Mai, H., Xu, A., and Lu, D. (2025) Small
extracellular vesicles from hypoxia-neuron maintain blood-brain barrier integrity, Stroke, 56, 1569-1580, https://
doi.org/10.1161/STROKEAHA.124.048446.
145. Liu, Y., Zhang, T., Zou, X., Yuan, Z., Li, Y., Zang, J., He, N., He, L., Xu, A., and Lu, D. (2024) Penumbra-targeted
CircOGDH siRNA-loaded nanoparticles alleviate neuronal apoptosis in focal brain ischaemia, Stroke Vasc. Neurol.,
9, 134-144, https://doi.org/10.1136/svn-2022-002009.
Publisher’s Note. Pleiades Publishing remains neutral with regard to jurisdictional claims in published
maps and institutional affiliations. AI tools may have been used in the translation or editing of this article.
