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biology
Review
From Powerhouse to Perpetrator—Mitochondria in
Health and Disease
Nima B. Fakouri 1, Thomas Lau Hansen 2, Claus Desler 2, Sharath Anugula 2
and Lene Juel Rasmussen 2, *
1
Laboratory of Molecular Gerontology, National Institute on Aging, National Institutes of Health, Baltimore,
MD 21224, USA; nima.borhanfakouri@nih.gov
2Center for Healthy Aging, Department of Cellular and Molecular Medicine, University of Copenhagen,
2200 Copenhagen, Denmark; tlhansen@sund.ku.dk (T.L.H.); cdesler@sund.ku.dk (C.D.);
sharath@sund.ku.dk (S.A.)
*Correspondence: lenera@sund.ku.dk
Received: 2 January 2019; Accepted: 5 March 2019; Published: 11 May 2019
Abstract:
In this review we discuss the interaction between metabolic stress, mitochondrial
dysfunction, and genomic instability. Unrepaired DNA damage in the nucleus resulting from
excess accumulation of DNA damages and stalled replication can initiate cellular signaling responses
that negatively affect metabolism and mitochondrial function. On the other hand, mitochondrial
pathologies can also lead to stress in the nucleus, and cause sensitivity to DNA-damaging agents.
These are examples of how hallmarks of cancer and aging are connected and influenced by each other
to protect humans from disease.
Keywords: mitochondria; cancer; nucleotide metabolism; DNA damage; NAD+
1. Introduction
It has been almost two decades since Hanahan and Weinberg for the first time classified the
hallmarks of cancer [
1
]. Ten years later, they updated that list and introduced genomic instability and
dysregulation of cellular energetics and mitochondrial function as emerging hallmarks [
2
]. Recently,
both genomic instability and mitochondrial dysfunction are considered as two of the key hallmarks of
aging [
3
]. Both of these have been implicated in several pathologies, as reviewed in References [
4
–
6
].
There are other hallmarks that are common between cancer and aging, such as epigenetic changes and
altered cellular communication. Moreover, other hallmarks are in opposition to each other in cancer
and aging. These include the dysregulation of apoptosis and senescence, which are stimulated in aging
cells and suppressed in cancer cells [
2
,
3
]. Whether we can consider cancer as the disease of aging is a
topic that is beyond the scope of this review, but age is the largest risk factor in the development of
cancer [7].
One of the key questions that remain to be answered is, how are these hallmarks are connected
and influenced by each other [
3
]? As these pathologic cellular changes occur gradually, understanding
the connection between them would help to develop more effective therapeutic strategies to treat
cancer or rather prevent it.
It has long been known that byproducts of cellular metabolism such as reactive oxygen and
nitrogen species (ROS and RNS) can damage cellular components and macromolecules including
DNA [
8
,
9
]. Damage to DNA can have severe effects on cells by blocking replication, transcription,
generation of DNA double and single strand breaks, as well as chromosome rearrangements [
10
,
11
].
Activation of the DNA damage response (DDR) following DNA damage is an energy-demanding
Biology 2019,8, 35; doi:10.3390/biology8020035 www.mdpi.com/journal/biology
Biology 2019,8, 35 2 of 15
process [
12
], and can deplete cells of substrates such as NAD
+
and ATP, which can in turn lead to
additional metabolic stress and mitochondrial dysfunction (Figure 1) [13–15].
Biology 2019, 8, x FOR PEER REVIEW 2 of 14
process [12], and can deplete cells of substrates such as NAD+ and ATP, which can in turn lead to
additional metabolic stress and mitochondrial dysfunction (Figure 1) [13–15].
Figure 1. Illustration of the mitochondrial-nuclear interactions in aging or cancer. Abnormal
metabolism and/or metabolic defects lead to metabolic stress and mitochondrial dysfunction. This is
followed by the increased generation of reactive oxygen and nitrogen species (ROS and RNS) as well
as reactive aldehydes. These reactive species can react and damage macromolecules such as proteins
and DNA. Damage to DNA causes genomic instability via stalled replication and transcription, and
the generation of double- and single-strand breaks (DSBs and SSBs, respectively) within the genome.
Increased activities of the DNA damage response (DDR) deplete cells of key cellular substrates and
cofactors, mainly ATP and NAD+. This generates a positive feedback that enhances metabolic stress
and mitochondrial dysfunction.
In this review, we aim to discuss the interaction between metabolic stress and mitochondrial
dysfunction with genomic instability. Stress in the nucleus, such as the accumulation of DNA damage
and stalled replication, negatively affects metabolism and mitochondrial function [13–17], while
mitochondrial pathologies lead to stress in the nucleus [18] and cause sensitivity to DNA-damaging
agents [19], as reviewed by Desler et al. in 2012 [20].
First, we explain how the accumulation of DNA damages and the activation of the DDR leads
to mitochondrial dysfunction. Next, we explain how the dysregulation of mitochondrial function and
metabolism contributes to the epigenetic changes, imbalanced dNTP pools, and genomic instability.
2. From DNA Damage to Mitochondrial Dysfunction
Activation of the main components of the DDR, including poly (ADP-ribose) polymerase
(PARP) enzymes (mainly PARP1 and PARP2) as well as ataxia telangiectasia mutated (ATM) [21],
DNA-dependent protein kinase (DNA-PK) [22,23] and P53 [24–27], is able to influence mitochondrial
function and cellular metabolism. Chronic activation of PARP1 negatively affects cellular physiology
and mitochondrial function [28]. Activation of ATM, DNA-PK, and P53 can influence mitochondrial
and cellular metabolism to promote either cell survival or death (Figure 2). Here, we briefly describe
how each of these enzymes are able to influence mitochondrial function.
Figure 1.
Illustration of the mitochondrial-nuclear interactions in aging or cancer. Abnormal metabolism
and/or metabolic defects lead to metabolic stress and mitochondrial dysfunction. This is followed by
the increased generation of reactive oxygen and nitrogen species (ROS and RNS) as well as reactive
aldehydes. These reactive species can react and damage macromolecules such as proteins and DNA.
Damage to DNA causes genomic instability via stalled replication and transcription, and the generation
of double- and single-strand breaks (DSBs and SSBs, respectively) within the genome. Increased
activities of the DNA damage response (DDR) deplete cells of key cellular substrates and cofactors,
mainly ATP and NAD
+
. This generates a positive feedback that enhances metabolic stress and
mitochondrial dysfunction.
In this review, we aim to discuss the interaction between metabolic stress and mitochondrial
dysfunction with genomic instability. Stress in the nucleus, such as the accumulation of DNA damage
and stalled replication, negatively affects metabolism and mitochondrial function [
13
–
17
], while
mitochondrial pathologies lead to stress in the nucleus [
18
] and cause sensitivity to DNA-damaging
agents [19], as reviewed by Desler et al. in 2012 [20].
First, we explain how the accumulation of DNA damages and the activation of the DDR leads to
mitochondrial dysfunction. Next, we explain how the dysregulation of mitochondrial function and
metabolism contributes to the epigenetic changes, imbalanced dNTP pools, and genomic instability.
2. From DNA Damage to Mitochondrial Dysfunction
Activation of the main components of the DDR, including poly (ADP-ribose) polymerase
(PARP) enzymes (mainly PARP1 and PARP2) as well as ataxia telangiectasia mutated (ATM) [
21
],
DNA-dependent protein kinase (DNA-PK) [
22
,
23
] and P53 [
24
–
27
], is able to influence mitochondrial
function and cellular metabolism. Chronic activation of PARP1 negatively affects cellular physiology
and mitochondrial function [
28
]. Activation of ATM, DNA-PK, and P53 can influence mitochondrial
and cellular metabolism to promote either cell survival or death (Figure 2). Here, we briefly describe
how each of these enzymes are able to influence mitochondrial function.
Biology 2019,8, 35 3 of 15
Biology 2019, 8, x FOR PEER REVIEW 3 of 14
Figure 2. Core DNA damage response proteins are able to influence mitochondrial activity and
quality control via multiple pathways. Abnormal DNA structure and certain genomic lesions, such
as strand breaks, activate poly (ADP-ribose) polymerase (PARP) enzymes, mainly PARP1. Chronic
activation of PARP1 can deplete the cell of NAD
+
, which is a rate-limiting substrate for SIRT1. SIRT1
together with AMP-activated kinase (AMPK) can enhance metabolism and mitochondrial function by
enhancing mitochondrial biogenesis and mitophagy. Activation of ataxia telangiectasia mutated
(ATM), Rad3-related (ATR) and DNA-dependent protein kinase (DNA-PK) following DNA damage
can promote the activation AKT and P53. Activation of AKT rewires cellular metabolism by inhibiting
Forkhead box (FOXO) enzymes. Deacetylation of P53 by SIRT1 targets P53 for degradation. Decrease
in SIRT1 activity stabilizes P53. P53 decreases mitophagy via inhibition of PTEN-induced kinase 1
(PINK1) and PARKIN transcription. SIRT1 promotes AMPK activity indirectly. Decrease in SIRT1
activity is followed by the decrease in activated AMPK.
3. PARP Modulates Mitochondrial Function and Cellular Metabolism
PARPs are a group of enzymes (16 in mice and 17 in humans) that are the main constituent of
the cellular stress response [29]. PARPs cleave NAD
+
to nicotinamide (NAM) and ADP-ribose
(ADPR), and the ADPR is subsequently transferred to certain amino acids within the target protein.
The attachment of poly ADP-ribose (PAR) to the target proteins is referred to as PARylation, and it
can affect protein–protein and protein–DNA interaction as well as protein localization [30]. PAR has
a short half-life and is degraded almost directly after its formation by the activity of the PAR-
degrading enzyme, poly(ADP-ribose) glycohydrolase (PARG) [31]. PARylation modulates several
key cellular processes such as chromatin structure, transcription, translation, cell cycle, DNA repair,
mitochondrial homeostasis, apoptosis, and metabolism [29,32]. PARP1 is activated by several
mechanisms, including mono(ADP-ribosyl)ation, phosphorylation, and acetylation [29]. PARP1
possesses a DNA-binding domain that recognizes abnormal DNA structures such as gapped DNA,
single- and double strand breaks, cruciform structures, and nucleosome linker DNA [33,34]. In the
initial steps of the repair, PARP1 PARylates histones and facilitates the chromatin relaxation that
provides more space for the recruitment of DNA repair proteins. Subsequent PAR generation recruits
the DNA repair proteins via their PAR-binding domains [35,36]. Under mild genotoxic stress, PARP
activation results in repair and survival. However, in response to DNA damage, PARP hyper-
activation results in decrease in NAD
+
and ATP levels, mitochondrial dysfunction, and eventually
cell death [32,35,37].
Figure 2.
Core DNA damage response proteins are able to influence mitochondrial activity and
quality control via multiple pathways. Abnormal DNA structure and certain genomic lesions, such
as strand breaks, activate poly (ADP-ribose) polymerase (PARP) enzymes, mainly PARP1. Chronic
activation of PARP1 can deplete the cell of NAD
+
, which is a rate-limiting substrate for SIRT1. SIRT1
together with AMP-activated kinase (AMPK) can enhance metabolism and mitochondrial function
by enhancing mitochondrial biogenesis and mitophagy. Activation of ataxia telangiectasia mutated
(ATM), Rad3-related (ATR) and DNA-dependent protein kinase (DNA-PK) following DNA damage
can promote the activation AKT and P53. Activation of AKT rewires cellular metabolism by inhibiting
Forkhead box (FOXO) enzymes. Deacetylation of P53 by SIRT1 targets P53 for degradation. Decrease
in SIRT1 activity stabilizes P53. P53 decreases mitophagy via inhibition of PTEN-induced kinase 1
(PINK1) and PARKIN transcription. SIRT1 promotes AMPK activity indirectly. Decrease in SIRT1
activity is followed by the decrease in activated AMPK.
3. PARP Modulates Mitochondrial Function and Cellular Metabolism
PARPs are a group of enzymes (16 in mice and 17 in humans) that are the main constituent
of the cellular stress response [
29
]. PARPs cleave NAD
+
to nicotinamide (NAM) and ADP-ribose
(ADPR), and the ADPR is subsequently transferred to certain amino acids within the target protein.
The attachment of poly ADP-ribose (PAR) to the target proteins is referred to as PARylation, and it
can affect protein–protein and protein–DNA interaction as well as protein localization [
30
]. PAR has a
short half-life and is degraded almost directly after its formation by the activity of the PAR-degrading
enzyme, poly(ADP-ribose) glycohydrolase (PARG) [
31
]. PARylation modulates several key cellular
processes such as chromatin structure, transcription, translation, cell cycle, DNA repair, mitochondrial
homeostasis, apoptosis, and metabolism [
29
,
32
]. PARP1 is activated by several mechanisms, including
mono(ADP-ribosyl)ation, phosphorylation, and acetylation [
29
]. PARP1 possesses a DNA-binding
domain that recognizes abnormal DNA structures such as gapped DNA, single- and double strand
breaks, cruciform structures, and nucleosome linker DNA [
33
,
34
]. In the initial steps of the repair,
PARP1 PARylates histones and facilitates the chromatin relaxation that provides more space for the
recruitment of DNA repair proteins. Subsequent PAR generation recruits the DNA repair proteins via
their PAR-binding domains [
35
,
36
]. Under mild genotoxic stress, PARP activation results in repair and
survival. However, in response to DNA damage, PARP hyper-activation results in decrease in NAD
+
and ATP levels, mitochondrial dysfunction, and eventually cell death [32,35,37].
Biology 2019,8, 35 4 of 15
4. DNA Damage can Activate Both Pro-Survival and Pro-Death Pathways That Involve the
Mitochondria
DNA damage and DDR can activate pathways that promote cell survival or death depending on
the extent and type of DNA damages [
38
]. In addition to PARP1, other immediate sensors of DDR
are enzymes belonging to the superfamily of phosphatidylinositol 3-kinase-related kinases (PIKKs),
including ATM, ataxia telangiectasia and Rad3-related (ATR), and DNA-PK. Activated ATR and ATM, in
turn, activate P53 through CHK1 and CHK2, respectively. Apart from preventing cell cycle progression
and the recruitment of DNA repair proteins, these enzymes can modulate mitochondrial function and
survival [
23
,
39
]. ATM, ATR, and DNA-PK are able to promote survival via the direct phosphorylation
of AKT (also known as protein kinase B, PKB) independently of growth factor signaling [
22
,
40
–
43
].
However, the mechanism of this interaction is not fully understood [
44
]. This is particularly interesting,
as in many cancer cells, the AKT is activated independently of growth factors [
45
]. Activated AKT
stimulates glucose uptake and ATP production through glycolysis, one of the main hallmarks of cancer,
also known as the Warburg effect [
46
]. In addition, activated AKT inhibits Forkhead box (FOXO)
transcription factors [
47
,
48
]. FOXO proteins regulate the expression of key genes that are involved in
mitochondrial biogenesis and homeostasis, such as peroxisome proliferator-activated receptor gamma
co-activator 1alpha (PGC1a) and PTEN-induced kinase 1 (PINK1). Decrease in FOXO activity results
in the decrease of mitochondrial biogenesis, mitophagy, autophagy, and lipolysis [49].
5. Mito-Nuclear Signaling in Aging and Cancer
For a long time, it was believed that mitochondria are regulated from the nucleus by the nuclear
genome, and changes in mitochondria follow changes in the nucleus [
50
,
51
]. However, during
recent years, accumulative evidence suggest that mitochondria and mitochondrial metabolites can
influence nuclear processes and gene expression in response to various stimuli and environmental
cues [
52
–
54
]. Mitochondrially generated ROS and intermediate metabolites are essential for several
processes, including proliferation, epigenetic modifications, and post-translational modifications [
54
,
55
].
In addition, mitochondria contribute to genomic stability by replenishing dNTP pools for replication
and repair of the genome (Figure 3) [
56
,
57
]. In this section we will discuss nuclear processes that are
dependent on mitochondrial function and intermediate metabolites.
Biology 2019,8, 35 5 of 15
Biology 2019, 8, x FOR PEER REVIEW 4 of 14
4. DNA Damage can Activate Both Pro-Survival and Pro-Death Pathways That Involve the
Mitochondria
DNA damage and DDR can activate pathways that promote cell survival or death depending
on the extent and type of DNA damages [38]. In addition to PARP1, other immediate sensors of DDR
are enzymes belonging to the superfamily of phosphatidylinositol 3-kinase-related kinases (PIKKs),
including ATM, ataxia telangiectasia and Rad3-related (ATR), and DNA-PK. Activated ATR and
ATM, in turn, activate P53 through CHK1 and CHK2, respectively. Apart from preventing cell cycle
progression and the recruitment of DNA repair proteins, these enzymes can modulate mitochondrial
function and survival [23,39]. ATM, ATR, and DNA-PK are able to promote survival via the direct
phosphorylation of AKT (also known as protein kinase B, PKB) independently of growth factor
signaling [22,40–43]. However, the mechanism of this interaction is not fully understood [44]. This is
particularly interesting, as in many cancer cells, the AKT is activated independently of growth factors
[45]. Activated AKT stimulates glucose uptake and ATP production through glycolysis, one of the
main hallmarks of cancer, also known as the Warburg effect [46]. In addition, activated AKT inhibits
Forkhead box (FOXO) transcription factors [47,48]. FOXO proteins regulate the expression of key
genes that are involved in mitochondrial biogenesis and homeostasis, such as peroxisome
proliferator-activated receptor gamma co-activator 1alpha (PGC1a) and PTEN-induced kinase 1
(PINK1). Decrease in FOXO activity results in the decrease of mitochondrial biogenesis, mitophagy,
autophagy, and lipolysis [49].
5. Mito-Nuclear Signaling in Aging and Cancer
For a long time, it was believed that mitochondria are regulated from the nucleus by the nuclear
genome, and changes in mitochondria follow changes in the nucleus [50,51]. However, during recent
years, accumulative evidence suggest that mitochondria and mitochondrial metabolites can influence
nuclear processes and gene expression in response to various stimuli and environmental cues [52–
54]. Mitochondrially generated ROS and intermediate metabolites are essential for several processes,
including proliferation, epigenetic modifications, and post-translational modifications [54,55]. In
addition, mitochondria contribute to genomic stability by replenishing dNTP pools for replication
and repair of the genome (Figure 3) [56,57]. In this section we will discuss nuclear processes that are
dependent on mitochondrial function and intermediate metabolites.
Figure 3.
Mitochondrial function and metabolites influence nuclear processes. Mitochondrial ROS
and secondary metabolites act as signaling molecules and cofactors that regulate fundamental nuclear
processes. Mitochondrial ROS that are released into the cytosol stabilize hypoxia-inducible factor 1-
α
(HIF1
α
) and activate the transcription of genes that are involved in proliferation. Citrate generated
through the TCA cycle is released into the cytosol and in the nucleus. It is further converted into
acetyl-coenzyme A (acetyl CoA) that is used for the acetylation of target proteins. Through a reverse
reaction, SIRT1 uses NAD
+
to deacetylate the target proteins. S-adenosylmethionine (SAM) is a methyl
donor that is generated from methionine and ATP in the cytosol. Demethylases such as Jumonji
C (JMJC) family members and the ten-eleven translocation (TET) methylcytosine hydroxylases use
α
-ketoglutarate (
α
-KG) as cofactor to remove methyl groups from proteins and DNA. AMPK stimulates
the activity of TET enzymes, and thus the inhibition of AMPK by glucose impairs the function TET
enzymes. The accumulation of fumarate and succinate due to impaired fumarate hydratase (FH) and
succinate dehydrogenase (SDH) can inhibit
α
-KG-dependent demethylases and even cause defects in
homologous recombination (HR) DNA repair.
6. Mitochondrial ROS Are Involved in Signaling and Determine Cell Fate
The mitochondrial electron transport chain (ETC) generates reactive oxygen species (ROS) as
the byproduct of oxidative phosphorylation (OXPHOS) from different complexes, though mainly
complexes I, II, and III. While complexes I and II exclusively create O
2·
in the mitochondrial matrix,
complex III produces O
2·
in both the matrix and intermembrane space [
58
]. However, the ROS that are
released outside of the matrix are converted into H
2
O
2
by cytosolic superoxide dismutase 1 (SOD1)
and participate in mitochondrial signaling through reversible cysteine oxidation [
59
]. Mitochondrially
produced ROS can serve as second-messenger molecules. Mitochondrial ROS (mtROS) are required for
the stabilization of HIF
α
(hypoxia-inducible factor 1-
α
) and the activation of downstream pathways
that promote proliferation [60].
It is possible that the type of cell and energy demand determine the effect of ROS and mtDNA
mutation over the cell. Cells that mostly rely on glycolysis will probably not be affected by mutation
in mtDNA under physiological conditions. During the exposure to stress and stimuli, however,
these cells might not be able to trigger an adaptive response to an increase in demand for ATP and
NAD
+
[
61
]. During stress and increased energy demand, cells respond by boosting cellular respiration
Biology 2019,8, 35 6 of 15
and mitochondrial activity to provide the cells with ATP and NAD
+
[
12
,
62
]. This is accompanied by
increased mitochondrial membrane potential and ROS above the physiological level [
61
,
63
]. Increased
ROS cause damage to macromolecules such as DNA, proteins, and lipids, which can cause cell death
or malignancy, as reviewed by Sies et al. in 2017 [
64
]. The inability to enhance ATP production in
response to stress and increased energy demand probably contributes to cellular deterioration during
aging [65].
In contrast to increased ROS generation, a decrease in ROS in metabolically active or proliferative
tissues interferes with cellular metabolism or proliferation that can promote senescence, as reviewed
by Diebold and Chandel in 2016 [66].
Hematopoietic stem cells (HSCs) represent an excellent example for both situations [
67
]. HSCs
are mainly quiescent, and they rely on glycolysis for ATP production [
68
]. While low levels of ROS
prevent the proliferation of HSCs and maintain their quiescent state, stress and increased energy
demand promote a shift toward ATP production by mitochondria and OXPHOS. This is accompanied
by increased ROS and proliferation [67]. Chronic stress followed by enhanced mitochondrial activity
and ROS generation leads to the depletion of HSCs, which is one of the hallmarks of aging [69–71].
7. Mitochondria Influence Post-Translational Modifications (PTMs) and Epigenetic Marks
Reversible acetylation and methylation are two frequently employed post-translational
modifications that regulate a variety of protein functions, protein stability, gene expression [
72
],
as well as DNA repair [
73
]. Epigenetic changes are also one of the main hallmarks of both aging and
cancer [
2
,
3
]. The modifications include alterations in DNA methylation, modifications of histones,
and chromatin remodeling. While DNA methylations show similar patterns in aging and cancer, the
histone modifications show distinct patterns, as reviewed by Zane et al. in 2014 [74].
The multiple enzymatic systems assuring the generation and maintenance of epigenetic patterns
include DNA methyltransferases, histone acetylases, deacetylases, methylases, and demethylases, as
well as protein complexes implicated in chromatin remodeling. Most PTMs, such as phosphorylation,
acetylation, methylation, and O-linked N-acetylglucosamine modification (O-GlcNAcylation), require
metabolites as substrates [
75
]. If not all, the majority of substrates required for PTMs are intermediate
metabolites generated by mitochondria [
54
]. Chromatin modifiers use metabolic intermediates as
cofactors or substrates, but are also regulated by their availability. These metabolites include NAD
+
for deacetylation, acetyl-CoA for histone acetylation, S-adenosylmethionine (SAM) for histone as well
as DNA methylation, and α-ketoglutarate (α-KG) for demethylation [54].
Lysine acetyltransferases (KATs) add acetyl groups to proteins while lysine deacetylases (KDACs)
remove acetyl groups from proteins [
76
,
77
]. KATs such as GCN5, CBP/p300, and MYST use
acetyl-coenzyme A (acetyl CoA) as an acetyl group donor for protein acetylation [
78
]. KDACs are
classified into two groups with different catalytic mechanisms: Zn
2+
-dependent histone deacetylases
(HDAC1-11) and NAD
+
-dependent deacetylases (SIRT1-7) [
76
,
77
]. Acetyl CoA is generated in
mitochondria and converted to citrate through the TCA cycle. Citrate can then be exported from
mitochondria. In the cytoplasm and nucleus, citrate is converted back into acetyl CoA via the function
of ATP-citrate lyase (ACLY). Citrate is the major source of acetyl CoA in the cytoplasm and the nucleus.
Depletion of mtDNA and the subsequent decrease in NAD negatively affect the TCA cycle and lead
to a decrease in histone acetylation. This can be rescued by the restoration of electron flow and TCA
cycle [
60
,
79
]. The degree of acetylation directly correlates with the availability of cofactors such as
acetyl CoA and NAD
+
. While increased nuclear acetyl CoA promotes increased acetylation and
the formation of euchromatin, an increase in NAD
+
promotes deacetylation and the formation of
heterochromatin, resulting in a decrease of gene expression [
79
]. MYC and AKT stimulate nutrient
uptake and promote acetyl CoA production via ACLY. AKT can directly phosphorylate and activate
ACLY to maintain the acetyl CoA levels, regardless of glucose concentrations [80].
Another key chromatin modification that is strongly interconnected with metabolism is
methylation [
54
,
81
]. Methylation is regulated by S-adenosylmethionine (SAM) abundance, whereby
Biology 2019,8, 35 7 of 15
SAM serves as a universal methyl donor, synthesized from methionine and ATP by methionine
adenosyltransferases (MATs) [
54
,
81
]. Histone and DNA methylation is removed by demethylases
such as Jumonji C (JMJC) family members and the ten-eleven translocation (TET) methylcytosine
hydroxylases, which use a dioxygenation reaction that requires Fe2+, O2, and α-ketoglutarate (α-KG)
as cofactors [
82
].
α
-KG is generated in the TCA cycle via the catabolism of glucose and glutamine.
The function of lysine-specific histone demethylase 1A (LSD1; KDM1A) and LSD2 (KDM1B), which
catalyze an amine oxidation reaction, is dependent on flavin adenine dinucleotide (FAD) [83,84].
The dysregulation of the TCA cycle and cellular metabolism affect PTMs, epigenetic changes, and
choice of DNA repair pathway. Accumulation of succinate or fumarate, which occurs respectively in
tumors deficient for succinate dehydrogenase (SDH) or fumarate hydratase (FH), similarly inhibits
α
-KG-dependent enzymes, leading to defects in homologous recombination (HR) DNA repair [
53
].
Changes in nutrient availability can directly affect chromatin modifications. The tumor suppressor
ten-eleven translocation (TET) protein family of dioxygenases (TET1, TET2, and TET3) converts a 5mC
DNA methylation to a hydroxymethylation, 5hmC [
85
]. AMP-activated kinase (AMPK) phosphorylates
TET2 and stabilizes this tumor suppressor protein. However, increase in blood glucose levels impairs
AMPK activity, which leads to destabilization of TET2 and the subsequent dysregulation of 5mCs and
5hmCs [86].
8. Regulation of dNTP Pools
In humans, nucleotide levels are maintained by the nucleotide salvage and/or de novo synthesis
of ribo- and deoxyribonucleotide triphosphates (rNTPs and dNTPs).
It is generally accepted that the levels and especially the relative balance of the cytosolic dNTP pools
have great influence on the replication, repair, and stability of the nuclear genome [
87
–
92
]. The efficiency
and fidelity of most, if not all, polymerases and many repair enzymes are affected by the level of substrate
nucleotides [
87
]. Too low a concentration of a nucleotide results in poor incorporation frequency,
and a level which is too high risks the misincorporation of the high-concentration nucleotide [
93
–
95
].
Therefore, it is of no surprise that the process of synthesizing dNTPs in the right concentrations is
governed by a generous amount of regulation, feedback loops, and redundancy [96,97].
Mitochondrial dysfunction, due to mutations in the mitochondrial genome or a decrease in the
mitochondrial DNA (mtDNA) copy number, is associated with a poor prognosis of many types of
cancer [
98
–
102
], also reviewed by Chatterjee et al. in 2006 [
103
]. The accumulation of mutations
in mtDNA or impeded ETC have even been associated with tumor aggressiveness [
99
,
101
,
104
–
106
].
We have previously shown a relationship between mitochondrial respiration and the regulation of
cytosolic dNTP pools, and demonstrated a co-occurring decrease of chromosomal stability [51].
The de novo synthesis of nucleotides is split up into purine and pyrimidine synthesis, which go
through two distinct pathways.
Despite having separate synthesis pathways, both purine and pyrimidine-based ribonucleotides
occupy a central role in cellular metabolism. In addition to being basal components of DNA and RNA,
they function as phosphate donors in the transport of cellular energy and participate in enzymatic
reactions as well as intracellular and extracellular signaling [107].
Both the salvage and the de novo synthesis pathways utilize an activated sugar intermediate:
5-phosphoribosyl-1-pyrophosphate (PRPP). PRPP is generated by the action of PRPP synthetase and is
utilized in both purine and pyrimidine synthesis [
108
]. The purine nucleotides are synthesized from
PRPP, through inosine 5-monophosphate (IMP), and further into AMP and GMP, through two separate
but allosterically regulated pathways. Pyrimidines, on the other hand, originate from the precursor
pyrimidine nucleotide, UMP, which is used to synthesize all the cellular ribosyl and deoxyribosyl
pyrimidines including UTP, CTP, dUMP, and dTTP [
108
]. A key catalytic enzyme in this process is the
dihydroorotate dehydrogenase (DHODH) [
109
], located in the inner mitochondrial membrane and
functionally codependent with the OXPHOS [
110
]. The oxidation of dihydroorotate by DHODH, to
form orotate, is a bottleneck reaction of the de novo synthesis of pyrimidines and is electrochemically
Biology 2019,8, 35 8 of 15
coupled with the reduction of ubiquinone to ubiquinol [
56
,
111
–
113
]. Mitochondrial respiration can
modulate the nucleotide synthesis at two separate steps. A decrease of the mitochondrial respiration is
therefore linked to an inhibition of the DHODH enzyme, which in turn modulates the synthesis of
pyrimidines [
51
,
114
]. Furthermore, the activity of the RNR complex is regulated by the binding of
ATP to its active site and inhibited by dATP. By affecting the levels of cytosolic ATP, the mitochondria
can influence the activity of RNR, and hence the levels of dNTPs. Furthermore, RNR is allosterically
regulated by the relative levels of individual NTPs and dNTPs, and so the de novo synthesis of
dNTPs and their relative balance is highly dependent on the mitochondrial supply of pyrimidines and
ATP [114–116].
The cytosolic pool of dNTPs, which supplies the replication of the nuclear genome, is cell
cycle-regulated. Synthesis is initiated at the beginning of the S-phase, and stopped upon reaching
G2. In the de novo pathway, this phase-dependent synthesis of dNTPs is mediated through the
RNR—specifically, through the cell cycle regulation of expression and degradation of the RNR
subunit RNR-R2 [
117
]. In the salvage pathway, the phase-dependent synthesis is controlled by the
translational regulation of constituents of the nucleotide salvage pathway [
118
]. Outside of S-phase, a
dNTP hydrolase called SAMHD1 (SAM and HD domain-containing deoxynucleoside triphosphate
triphosphohydrolase 1) depletes the dNTP pools by its hydrolase activity to block viral replication,
amongst other things [
119
]. In response to DNA damage, however, dNTP levels can increase by up to
4-fold [
96
]. In this case, both the RNR and SAMHD1 are then recruited to the site of damage to tightly
regulate the amount of dNTPs supplied to the DNA repair machinery [96,120].
Not only replication of the nuclear genome requires a balanced dNTP pool. Unlike the replication
of the nuclear genome, the replication of mitochondrial DNA is not regulated by the cell cycle,
but is carried out continuously in both mitotic and post-mitotic cells and tissue. Imbalance of the
mitochondrial dNTP pools affects the replication of mtDNA, resulting in the accumulation of point
mutations and deletions [121,122].
The dNTP pool of the mitochondrial compartment is somewhat separate from the much larger
pool supplying the nuclear genome [
123
,
124
]. However, cytosolic de novo synthesis of dNTP is
essential for mtDNA maintenance, even in post-mitotic cells and tissue [125–127].
P53R2 is a protein that substitutes RNR-R2 in post-mitotic cells in response to DNA damage.
P53R2 is transcribed by P53 and results, when forming the complex, in the activation of RNR and the
de novo synthesis of dNTPs intended as substrates for DNA repair mechanisms [
128
]. Mutations of the
RRM2B gene encoding P53R2 have been shown to induce mtDNA replication and repair deficiency in
post-mitotic, but not dividing, human fibroblasts [
125
]. In humans, mutations in the gene encoding the
RRM2B subunit have been correlated with severe mtDNA depletion of muscle tissue, and RRM2b
−/−
mice further display a severe decrease of mtDNA content in liver, kidney, and muscle [126].
It is important to realize that balanced dNTP pools for mtDNA maintenance do not need to be of
mitochondrial origin. Imbalances of the nuclear dNTP pools resulting from genetic predisposition, age,
or even just diet [
129
] have the potential to start a vicious cycle, whereby failure to maintain mtDNA
integrity results in the decreased synthesis of pyrimidines and further dNTP pool imbalance.
Mitochondrial dysfunction is therefore not only a risk to the mitotic cell, but also fully differentiated
post-mitotic cells [
130
], and is involved in the etiology of a wide array of pathologies, including cancer
and Alzheimer’s disease [131–133].
9. Conclusions
According to recent advancements, hallmarks of cancer include genomic instability, dysregulation
of cellular energetics, and mitochondrial dysfunction, which also are common pathways important
for cellular aging. Mitochondrial dysfunction is associated with a poor prognosis of many types
of cancer, which could very well be linked to an imbalance of the cytosolic dNTP pools, as both of
these conditions are related to one of the hallmarks of cancer—chromosomal instability. A better
Biology 2019,8, 35 9 of 15
understanding of these pathological cellular processes would advance the development of therapeutic
modalities in the prevention of cancer and at the same time help the understanding of biological aging.
Author Contributions: N.B.F., T.L.H., C.D., S.A., L.J.R. wrote the review.
Funding: This research was funded by Nordea-fonden and Olav Thon Foundation.
Conflicts of Interest: The authors declare no conflict of interest.
References
1. Hanahan, D.; Weinberg, R.A. The hallmarks of cancer. Cell 2000,100, 57–70. [CrossRef]
2.
Hanahan, D.; Weinberg, R.A. Hallmarks of Cancer: The Next Generation. Cell
2011
,144, 646–674. [CrossRef]
[PubMed]
3.
L
ó
pez-Ot
í
n, C.; Blasco, M.A.; Partridge, L.; Serrano, M.; Kroemer, G. The hallmarks of aging. Cell
2013
,153,
1194–1217. [CrossRef]
4.
Tubbs, A.; Nussenzweig, A. Endogenous DNA Damage as a Source of Genomic Instability in Cancer. Cell
2017,168, 644–656. [CrossRef]
5.
Sharma, P.; Sampath, H. Mitochondrial DNA Integrity: Role in Health and Disease. Cells
2019
,8, 100.
[CrossRef] [PubMed]
6.
Fan, P.; Xie, X.-H.; Chen, C.-H.; Peng, X.; Zhang, P.; Yang, C.; Wang, Y.-T. Molecular Regulation Mechanisms
and Interactions Between Reactive Oxygen Species and Mitophagy. DNA Cell Biol.
2019
,38, 10–22. [CrossRef]
[PubMed]
7.
Aunan, J.R.; Cho, W.C.; Søreide, K. The Biology of Aging and Cancer: A Brief Overview of Shared and
Divergent Molecular Hallmarks. Aging Dis. 2017,8, 628. [CrossRef] [PubMed]
8.
Burcham, P.C. Internal hazards: Baseline DNA damage by endogenous products of normal metabolism.
Mutat. Res./Genet. Toxicol. Environ. Mutagenesis 1999,443, 11–36. [CrossRef]
9.
De Bont, R.; van Larebeke, N. Endogenous DNA damage in humans: A review of quantitative data.
Mutagenesis 2004,19, 169–185. [CrossRef]
10.
Zeman, M.K.; Cimprich, K.A. Causes and consequences of replication stress. Nat. Cell Biol.
2014
,16, 2–9.
[CrossRef] [PubMed]
11.
White, R.R.; Vijg, J. Do DNA Double-Strand Breaks Drive Aging? Mol. Cell
2016
,63, 729–738. [CrossRef]
[PubMed]
12. Qin, L.; Fan, M.; Candas, D.; Jiang, G.; Papadopoulos, S.; Tian, L.; Woloschak, G.; Grdina, D.J.; Li, J.J. CDK1
Enhances Mitochondrial Bioenergetics for Radiation-Induced DNA Repair. Cell Rep.
2015
,13, 2056–2063.
[CrossRef]
13.
Scheibye-Knudsen, M.; Mitchell, S.J.; Fang, E.F.; Iyama, T.; Ward, T.; Wang, J.; Dunn, C.A.; Singh, N.; Veith, S.;
Hasan-Olive, M.M.; et al. A high-fat diet and NAD(+) activate Sirt1 to rescue premature aging in cockayne
syndrome. Cell Metab. 2014,20, 840–855. [CrossRef]
14.
Fang, E.F.; Kassahun, H.; Croteau, D.L.; Scheibye-Knudsen, M.; Marosi, K.; Lu, H.; Shamanna, R.A.;
Kalyanasundaram, S.; Bollineni, R.C.; Wilson, M.A.; et al. NAD(+) Replenishment Improves Lifespan and
Healthspan in Ataxia Telangiectasia Models via Mitophagy and DNA Repair. Cell Metab.
2016
,24, 566–581.
[CrossRef]
15.
Fakouri, N.B.; Durhuus, J.A.; Regnell, C.E.; Angleys, M.; Desler, C.; Olive, M.H.; Mart
í
n-Pardillos, A.;
Tsaalbi-Shtylik, A.; Thomsen, K.; Lauritzen, M.; et al. Rev1 contributes to proper mitochondrial function via
the PARP-NAD+-SIRT1-PGC1αaxis. Sci. Rep. 2017,7, 12480. [CrossRef]
16.
Rivera-Torres, J.; Ac
í
n-Perez, R.; Cabezas-S
á
nchez, P.; Osorio, F.G.; Gonzalez-G
ó
mez, C.; Megias, D.;
C
á
mara, C.; L
ó
pez-Ot
í
n, C.; Enr
í
quez, J.A.; Luque-Garc
í
a, J.L.; et al. Identification of mitochondrial
dysfunction in Hutchinson–Gilford progeria syndrome through use of stable isotope labeling with amino
acids in cell culture. J. Proteom. 2013,91, 466–477. [CrossRef] [PubMed]
17.
Fang, E.F.; Scheibye-Knudsen, M.; Brace, L.E.; Kassahun, H.; SenGupta, T.; Nilsen, H.; Mitchell, J.R.;
Croteau, D.L.; Bohr, V.A. Defective mitophagy in XPA via PARP-1 hyperactivation and NAD(+)/SIRT1
reduction. Cell 2014,157, 882–896. [CrossRef]
Biology 2019,8, 35 10 of 15
18.
Qian, W.; Choi, S.; Gibson, G.A.; Watkins, S.C.; Bakkenist, C.J.; Van Houten, B. Mitochondrial hyperfusion
induced by loss of the fission protein Drp1 causes ATM-dependent G2/M arrest and aneuploidy through
DNA replication stress. J. Cell Sci. 2012,125, 5745–5757. [CrossRef] [PubMed]
19.
Temelie, M.; Savu, D.I.; Moisoi, N. Intracellular and Intercellular Signalling Mechanisms following DNA
Damage Are Modulated By PINK1. Oxid. Med. Cell. Longev. 2018,2018, 1–15. [CrossRef]
20.
Desler, C.; Hansen, T.L.; Frederiksen, J.B.; Marcker, M.L.; Singh, K.K.; Juel Rasmussen, L. Is There a Link
between Mitochondrial Reserve Respiratory Capacity and Aging? J. Aging Res.
2012
,2012, 192503. [CrossRef]
21.
Cosentino, C.; Grieco, D.; Costanzo, V. ATM activates the pentose phosphate pathway promoting anti-oxidant
defence and DNA repair. EMBO J. 2011,30, 546–555. [CrossRef] [PubMed]
22.
Bozulic, L.; Surucu, B.; Hynx, D.; Hemmings, B.A. PKB
α
/Akt1 Acts Downstream of DNA-PK in the DNA
Double-Strand Break Response and Promotes Survival. Mol. Cell 2008,30, 203–213. [CrossRef]
23.
Park, S.-J.; Gavrilova, O.; Brown,A.L.; Soto, J.E.; Bremner, S.; Kim, J.; Xu, X.; Yang, S.; Um, J.-H.; Koch, L.G.;et al.
DNA-PK Promotes the Mitochondrial, Metabolic, and Physical Decline that Occurs During Aging. Cell Metab.
2017,25, 1135–1146.e7. [CrossRef] [PubMed]
24.
Matoba, S.; Kang, J.-G.; Patino, W.D.; Wragg, A.; Boehm, M.; Gavrilova, O.; Hurley, P.J.; Bunz, F.; Hwang, P.M.
p53 regulates mitochondrial respiration. Science 2006,312, 1650–1653. [CrossRef]
25. Bensaad, K.; Vousden, K.H. p53: New roles in metabolism. Trends Cell Biol. 2007,17, 286–291. [CrossRef]
26.
Hoshino, A.; Mita, Y.; Okawa, Y.; Ariyoshi, M.; Iwai-Kanai, E.; Ueyama, T.; Ikeda, K.; Ogata, T.; Matoba, S.
Cytosolic p53 inhibits Parkin-mediated mitophagy and promotes mitochondrial dysfunction in the mouse
heart. Nat. Commun. 2013,4, 2308. [CrossRef] [PubMed]
27.
Wang, D.B.; Kinoshita, C.; Kinoshita, Y.; Morrison, R.S. p53 and mitochondrial function in neurons. Biochim.
Biophys. Acta Mol. Basis Dis. 2014,1842, 1186–1197. [CrossRef]
28.
Bai, P.; Nagy, L.; Fodor, T.; Liaudet, L.; Pacher, P. Poly(ADP-ribose) polymerases as modulators of
mitochondrial activity. Trends Endocrinol. Metab. 2015,26, 75–83. [CrossRef]
29.
Luo, X.; Kraus, W.L. On PAR with PARP: Cellular stress signaling through poly(ADP-ribose) and PARP-1.
Genes Dev. 2012,26, 417–432. [CrossRef]
30.
Krietsch, J.; Rouleau, M.; Pic,
É
.; Ethier, C.; Dawson, T.M.; Dawson, V.L.; Masson, J.-Y.; Poirier, G.G.;
Gagn
é
, J.-P. Reprogramming cellular events by poly(ADP-ribose)-binding proteins. Mol. Asp. Med.
2013
,34,
1066–1087. [CrossRef]
31.
Davidovic, L.; Vodenicharov, M.; Affar, E.B.; Poirier, G.G. Importance of poly(ADP-ribose) glycohydrolase in
the control of poly(ADP-ribose) metabolism. Exp. Cell Res. 2001,268, 7–13. [CrossRef]
32.
Bürkle, A.; Virag, L. Poly(ADP-ribose): PARadigms and PARadoxes. Mol. Asp. Med.
2013
,34, 1046–1065.
[CrossRef]
33.
Langelier, M.-F.; Planck, J.L.; Roy, S.; Pascal, J.M. Crystal structures of poly(ADP-ribose) polymerase-1
(PARP-1) zinc fingers bound to DNA: Structural and functional insights into DNA-dependent PARP-1
activity. J. Biol. Chem. 2011,286, 10690–10701. [CrossRef] [PubMed]
34.
Ali, A.A.E.; Timinszky, G.; Arribas-Bosacoma, R.; Kozlowski, M.; Hassa, P.O.; Hassler, M.; Ladurner, A.G.;
Pearl, L.H.; Oliver, A.W. The zinc-finger domains of PARP1 cooperate to recognize DNA strand breaks.
Nat. Struct. Mol. Biol. 2012,19, 685–692. [CrossRef] [PubMed]
35.
Gottschalk, A.J.; Timinszky, G.; Kong, S.E.; Jin, J.; Cai, Y.; Swanson, S.K.; Washburn, M.P.; Florens, L.;
Ladurner, A.G.; Conaway, J.W.; et al. Poly(ADP-ribosyl)ation directs recruitment and activation of an
ATP-dependent chromatin remodeler. Proc. Natl. Acad. Sci. USA
2009
,106, 13770–13774. [CrossRef]
[PubMed]
36.
Krishnakumar, R.; Kraus, W.L. The PARP side of the nucleus: Molecular actions, physiological outcomes,
and clinical targets. Mol. Cell 2010,39, 8–24. [CrossRef] [PubMed]
37.
Fang, E.F.; Scheibye-Knudsen, M.; Chua, K.F.; Mattson, M.P.; Croteau, D.L.; Bohr, V.A. Nuclear DNA damage
signalling to mitochondria in ageing. Nat. Rev. Mol. Cell Biol. 2016,17, 308–321. [CrossRef] [PubMed]
38.
Roos, W.P.; Thomas, A.D.; Kaina, B. DNA damage and the balance between survival and death in cancer
biology. Nat. Rev. Cancer 2016,16, 20–33. [CrossRef]
39.
Maryanovich, M.; Zaltsman, Y.; Ruggiero, A.; Goldman, A.; Shachnai, L.; Zaidman, S.L.; Porat, Z.; Golan, K.;
Lapidot, T.; Gross, A. An MTCH2 pathway repressing mitochondria metabolism regulates haematopoietic
stem cell fate. Nat. Commun. 2015,6, 7901. [CrossRef]
Biology 2019,8, 35 11 of 15
40.
Feng, J.; Park, J.; Cron, P.; Hess, D.; Hemmings, B.A. Identification of a PKB/Akt hydrophobic motif Ser-473
kinase as DNA-dependent protein kinase. J. Biol. Chem. 2004,279, 41189–41196. [CrossRef]
41.
Matsuoka, S.; Ballif, B.A.; Smogorzewska, A.; McDonald, E.R.; Hurov, K.E.; Luo, J.; Bakalarski, C.E.; Zhao, Z.;
Solimini, N.; Lerenthal, Y.; et al. ATM and ATR substrate analysis reveals extensive protein networks
responsive to DNA damage. Science 2007,316, 1160–1166. [CrossRef]
42.
Park, J.; Feng, J.; Li, Y.; Hammarsten, O.; Brazil, D.P.; Hemmings, B.A. DNA-dependent protein
kinase-mediated phosphorylation of protein kinase B requires a specific recognition sequence in the
C-terminal hydrophobic motif. J. Biol. Chem. 2009,284, 6169–6174. [CrossRef]
43.
Toulany, M.; Maier, J.; Iida, M.; Rebholz, S.; Holler, M.; Grottke, A.; Jüker, M.; Wheeler, D.L.; Rothbauer, U.;
Rodemann, H.P. Akt1 and Akt3 but not Akt2 through interaction with DNA-PKcs stimulate proliferation
and post-irradiation cell survival of K-RAS-mutated cancer cells. Cell Death Discov.
2017
,3, 17072. [CrossRef]
[PubMed]
44.
Liu, Q.; Turner, K.M.; Yung, W.K.A.; Chen, K.; Zhang, W. Role of AKT signaling in DNA repair and clinical
response to cancer therapy. Neuro-Oncology 2014,16, 1313–1323. [CrossRef]
45.
Stronach, E.A.; Chen, M.; Maginn, E.N.; Agarwal, R.; Mills, G.B.; Wasan, H.; Gabra, H. DNA-PK mediates
AKT activation and apoptosis inhibition in clinically acquired platinum resistance. Neoplasia
2011
,13,
1069–1080. [CrossRef]
46.
Robey, R.B.; Hay, N. Is Akt the “Warburg kinase”-Akt-energy metabolism interactions and oncogenesis.
Semin. Cancer Biol. 2009,19, 25–31. [CrossRef]
47.
Brunet, A.; Bonni, A.; Zigmond, M.J.; Lin, M.Z.; Juo, P.; Hu, L.S.; Anderson, M.J.; Arden, K.C.; Blenis, J.;
Greenberg, M.E. Akt promotes cell survival by phosphorylating and inhibiting a Forkhead transcription
factor. Cell 1999,96, 857–868. [CrossRef]
48.
Kops, G.J.P.L.; de Ruiter, N.D.; De Vries-Smits, A.M.M.; Powell, D.R.; Bos, J.L.; Burgering, B.M.T. Direct
control of the Forkhead transcription factor AFX by protein kinase B. Nature
1999
,398, 630–634. [CrossRef]
49.
Webb, A.E.; Brunet, A. FOXO transcription factors: Key regulators of cellular quality control. Trends Biochem.
Sci. 2014,39, 159–169. [CrossRef] [PubMed]
50.
Gray, M.W. Mitochondrial evolution. Cold Spring Harb. Perspect. Biol.
2012
,4, a011403. [CrossRef] [PubMed]
51.
Lee, C.; Yen, K.; Cohen, P. Humanin: A harbinger of mitochondrial-derived peptides? Trends Endocrinol.
Metab. 2013,24, 222–228. [CrossRef] [PubMed]
52.
Kim, K.H.; Son, J.M.; Benayoun, B.A.; Lee, C. The Mitochondrial-Encoded Peptide MOTS-c Translocates
to the Nucleus to Regulate Nuclear Gene Expression in Response to Metabolic Stress. Cell Metab.
2018
,28,
516–524. [CrossRef]
53.
Sulkowski, P.L.; Sundaram, R.K.; Oeck, S.; Corso, C.D.; Liu, Y.; Noorbakhsh, S.; Niger, M.; Boeke, M.;
Ueno, D.; Kalathil, A.N.; et al. Krebs-cycle-deficient hereditary cancer syndromes are defined by defects in
homologous-recombination DNA repair. Nat. Genet. 2018,50, 1086–1092. [CrossRef]
54.
Li, X.; Egervari, G.; Wang, Y.; Berger, S.L.; Lu, Z. Regulation of chromatin and gene expression by metabolic
enzymes and metabolites. Nat. Rev. Mol. Cell Biol. 2018,19, 563–578. [CrossRef] [PubMed]
55.
Sivanand, S.; Rhoades, S.; Jiang, Q.; Lee, J.V.; Benci, J.; Zhang, J.; Yuan, S.; Viney, I.; Zhao, S.; Carrer, A.;
et al. Nuclear Acetyl-CoA Production by ACLY Promotes Homologous Recombination. Mol. Cell
2017
,
67, 252–265. [CrossRef] [PubMed]
56.
Desler, C.; Munch-Petersen, B.; Stevnsner, T.; Matsui, S.-I.; Kulawiec, M.; Singh, K.K.; Rasmussen, L.J.
Mitochondria as determinant of nucleotide pools and chromosomal stability. Mutat. Res.
2007
,625, 112–124.
[CrossRef]
57.
Rasmussen, A.K.; Chatterjee, A.; Rasmussen, L.J.; Singh, K.K. Mitochondria-mediated nuclear mutator
phenotype in Saccharomyces cerevisiae. Nucleic Acids Res. 2003,31, 3909–3917. [CrossRef]
58.
Murphy, M.P. How mitochondria produce reactive oxygen species. Biochem. J.
2009
,417, 1–13. [CrossRef]
[PubMed]
59.
Sena, L.A.; Chandel, N.S. Physiological roles of mitochondrial reactive oxygen species. Mol. Cell
2012
,
48, 158–167. [CrossRef] [PubMed]
60.
Mart
í
nez-Reyes, I.; Diebold, L.P.; Kong, H.; Schieber, M.; Huang, H.; Hensley, C.T.; Mehta, M.M.; Wang, T.;
Santos, J.H.; Woychik, R.; et al. TCA Cycle and Mitochondrial Membrane Potential Are Necessary for Diverse
Biological Functions. Mol. Cell 2016,61, 199–209. [CrossRef]
Biology 2019,8, 35 12 of 15
61.
Westermann, B. Bioenergetic role of mitochondrial fusion and fission. Biochim. Biophys. Acta Bioenerget.
2012
,
1817, 1833–1838. [CrossRef]
62.
Brace, L.E.; Vose, S.C.; Stanya, K.; Gathungu, R.M.; Marur, V.R.; Longchamp, A.; Treviño-Villarreal, H.;
Mejia, P.; Vargas, D.; Inouye, K.; et al. Increased oxidative phosphorylation in response to acute and chronic
DNA damage. NPJ Aging Mech. Dis. 2016,2, 16022. [CrossRef] [PubMed]
63.
Youle, R.J.; van der Bliek, A.M. Mitochondrial fission, fusion, and stress. Science
2012
,337, 1062–1065.
[CrossRef]
64. Sies, H.; Berndt, C.; Jones, D.P. Oxidative Stress. Annu. Rev. Biochem. 2017,86, 715–748. [CrossRef]
65.
Trifunovic, A.; Hansson, A.; Wredenberg, A.; Rovio, A.T.; Dufour, E.; Khvorostov, I.; Spelbrink, J.N.;
Wibom, R.; Jacobs, H.T.; Larsson, N.-G. Somatic mtDNA mutations cause aging phenotypes without affecting
reactive oxygen species production. Proc. Natl. Acad. Sci. USA 2005,102, 17993–17998. [CrossRef]
66.
Diebold, L.; Chandel, N.S. Mitochondrial ROS regulation of proliferating cells. Free Radic. Biol. Med.
2016
,
100, 86–93. [CrossRef] [PubMed]
67.
Maryanovich, M.; Gross, A. A ROS rheostat for cell fate regulation. Trends Cell Biol.
2013
,23, 129–134.
[CrossRef] [PubMed]
68.
Folmes, C.D.L.; Dzeja, P.P.; Nelson, T.J.; Terzic, A. Metabolic Plasticity in Stem Cell Homeostasis and
Differentiation. Cell Stem Cell 2012,11, 596–606. [CrossRef] [PubMed]
69.
Nijnik, A.; Woodbine, L.; Marchetti, C.; Dawson, S.; Lambe, T.; Liu, C.; Rodrigues, N.P.; Crockford, T.L.;
Cabuy, E.; Vindigni, A.; et al. DNA repair is limiting for haematopoietic stem cells during ageing. Nature
2007,447, 686–690. [CrossRef]
70.
Niedernhofer, L.J. DNA repair is crucial for maintaining hematopoietic stem cell function. DNA Repair
2008
,
7, 523–529. [CrossRef] [PubMed]
71.
Flach, J.; Bakker, S.T.; Mohrin, M.; Conroy, P.C.; Pietras, E.M.; Reynaud, D.; Alvarez, S.; Diolaiti, M.E.;
Ugarte, F.; Forsberg, E.C.; et al. Replication stress is a potent driver of functional decline in ageing
haematopoietic stem cells. Nature 2014,512, 198–202. [CrossRef]
72.
Drazic, A.; Myklebust, L.M.; Ree, R.; Arnesen, T. The world of protein acetylation. Biochim. Biophys. Acta
Proteins Proteom. 2016,1864, 1372–1401. [CrossRef] [PubMed]
73.
Hauer, M.H.; Gasser, S.M. Chromatin and nucleosome dynamics in DNA damage and repair. Genes Dev.
2017,31, 2204–2221. [CrossRef]
74. Zane, L.; Sharma, V.; Misteli, T. Common features of chromatin in aging and cancer: Cause or coincidence?
Trends Cell Biol. 2014,24, 686–694. [CrossRef] [PubMed]
75.
Su, X.; Wellen, K.E.; Rabinowitz, J.D. Metabolic control of methylation and acetylation. Curr. Opin. Chem.
Biol. 2016,30, 52–60. [CrossRef] [PubMed]
76.
Choudhary, C.; Weinert, B.T.; Nishida, Y.; Verdin, E.; Mann, M. The growing landscape of lysine acetylation
links metabolism and cell signalling. Nat. Rev. Mol. Cell Biol. 2014,15, 536–550. [CrossRef]
77.
Ali, I.; Conrad, R.J.; Verdin, E.; Ott, M. Lysine Acetylation Goes Global: From Epigenetics to Metabolism and
Therapeutics. Chem. Rev. 2018,118, 1216–1252. [CrossRef] [PubMed]
78.
Montgomery, D.C.; Sorum, A.W.; Guasch, L.; Nicklaus, M.C.; Meier, J.L. Metabolic Regulation of Histone
Acetyltransferases by Endogenous Acyl-CoA Cofactors. Chem. Biol. 2015,22, 1030–1039. [CrossRef]
79.
Katada, S.; Imhof, A.; Sassone-Corsi, P. Connecting Threads: Epigenetics and Metabolism. Cell
2012
,148,
24–28. [CrossRef]
80.
Lee, J.V.; Carrer, A.; Shah, S.; Snyder, N.W.; Wei, S.; Venneti, S.; Worth, A.J.; Yuan, Z.-F.; Lim, H.-W.; Liu, S.;
et al. Akt-Dependent Metabolic Reprogramming Regulates Tumor Cell Histone Acetylation. Cell Metab.
2014,20, 306–319. [CrossRef]
81.
Kinnaird, A.; Zhao, S.; Wellen, K.E.; Michelakis, E.D. Metabolic control of epigenetics in cancer. Nat. Rev.
Cancer 2016,16, 694–707. [CrossRef]
82.
Metzger, E.; Schüle, R. The expanding world of histone lysine demethylases. Nat. Struct. Mol. Biol.
2007
,14,
252–254. [CrossRef] [PubMed]
83.
Forneris, F.; Binda, C.; Vanoni, M.A.; Mattevi, A.; Battaglioli, E. Histone demethylation catalysed by LSD1 is
a flavin-dependent oxidative process. FEBS Lett. 2005,579, 2203–2207. [CrossRef]
84.
Metzger, E.; Imhof, A.; Patel, D.; Kahl, P.; Hoffmeyer, K.; Friedrichs, N.; Müller, J.M.; Greschik, H.; Kirfel, J.;
Ji, S.; et al. Phosphorylation of histone H3T6 by PKC
β
I controls demethylation at histone H3K4. Nature
2010
,
464, 792–796. [CrossRef]
Biology 2019,8, 35 13 of 15
85.
He, Y.-F.; Li, B.-Z.; Li, Z.; Liu, P.; Wang, Y.; Tang, Q.; Ding, J.; Jia, Y.; Chen, Z.; Li, L.; et al. Tet-Mediated
Formation of 5-Carboxylcytosine and Its Excision by TDG in Mammalian DNA. Science
2011
,333, 1303–1307.
[CrossRef] [PubMed]
86.
Wu, D.; Hu, D.; Chen, H.; Shi, G.; Fetahu, I.S.; Wu, F.; Rabidou, K.; Fang, R.; Tan, L.; Xu, S.; et al.
Glucose-regulated phosphorylation of TET2 by AMPK reveals a pathway linking diabetes to cancer. Nature
2018,559, 637–641. [CrossRef]
87.
Pai, C.-C.; Kearsey, S.E. A Critical Balance: dNTPs and the Maintenance of Genome Stability. Genes
2017
,
8, 57. [CrossRef]
88.
Kunz, B.A. Mutagenesis and deoxyribonucleotide pool imbalance. Mutat. Res.
1988
,200, 133–147. [CrossRef]
89.
Kunz, B.A.; Kohalmi, S.E.; Kunkel, T.A.; Mathews, C.K.; Mclntosh, E.M.; Reidy, J.A. Deoxyribonucleoside
triphosphate levels: A critical factor in the maintenance of genetic stability. Mutat. Res. Toxicol.
1994
,318,
1–64. [CrossRef]
90.
Mathews, C.K. DNA precursor metabolism and genomic stability. FASEB J.
2006
,20, 1300–1314. [CrossRef]
[PubMed]
91.
Kumar, D.; Abdulovic, A.L.; Viberg, J.; Nilsson, A.K.; Kunkel, T.A.; Chabes, A. Mechanisms of mutagenesis
in vivo due to imbalanced dNTP pools. Nucleic Acids Res. 2011,39, 1360–1371. [CrossRef] [PubMed]
92.
Reichard, P. Interactions between deoxyribonucleotide and DNA synthesis. Annu. Rev. Biochem.
1988
,57,
349–374. [CrossRef]
93.
Meuth, M. The molecular basis of mutations induced by deoxyribonucleoside triphosphate pool imbalances
in mammalian cells. Exp. Cell Res. 1989,181, 305–316. [CrossRef]
94.
Meuth, M. The genetic consequences of nucleotide precursor pool imbalance in mammalian cells. Mutat. Res.
Mol. Mech. Mutagen. 1984,126, 107–112. [CrossRef]
95.
Haracska, L.; Prakash, S.; Prakash, L. Replication pastO6-Methylguanine by Yeast and Human DNA
Polymerase η.Mol. Cell. Biol. 2000,20, 8001–8007. [CrossRef]
96.
Niida, H.; Shimada, M.; Murakami, H.; Nakanishi, M. Mechanisms of dNTP supply that play an essential role
in maintaining genome integrity in eukaryotic cells. Cancer Sci.
2010
,101, 2505–2509. [CrossRef] [PubMed]
97.
Niida, H.; Katsuno, Y.; Sengoku, M.; Shimada, M.; Yukawa, M.; Ikura, M.; Ikura, T.; Kohno, K.; Shima, H.;
Suzuki, H.; et al. Essential role of Tip60-dependent recruitment of ribonucleotide reductase at DNA damage
sites in DNA repair during G1 phase. Genes Dev. 2010,24, 333–338. [CrossRef]
98.
Tseng, L.-M.; Yin, P.-H.; Chi, C.-W.; Hsu, C.-Y.; Wu, C.-W.; Lee, L.-M.; Wei, Y.-H.; Lee, H.-C. Mitochondrial
DNA mutations and mitochondrial DNA depletion in breast cancer. Genes Chromosomes Cancer
2006
,45,
629–638. [CrossRef]
99.
Yu, M.; Zhou, Y.; Shi, Y.; Ning, L.; Yang, Y.; Wei, X.; Zhang, N.; Hao, X.; Niu, R. Reduced mitochondrial DNA
copy number is correlated with tumor progression and prognosis in Chinese breast cancer patients. IUBMB
Life 2007,59, 450–457. [CrossRef] [PubMed]
100.
Yamada, S.; Nomoto, S.; Fujii, T.; Kaneko, T.; Takeda, S.; Inoue, S.; Kanazumi, N.; Nakao, A. Correlation
between copy number of mitochondrial DNA and clinico-pathologic parameters of hepatocellular carcinoma.
Eur. J. Surg. Oncol. 2006,32, 303–307. [CrossRef]
101.
Matsuyama, W.; Nakagawa, M.; Wakimoto, J.; Hirotsu, Y.; Kawabata, M.; Osame, M. Mitochondrial DNA
mutation correlates with stage progression and prognosis in non-small cell lung cancer. Hum. Mutat.
2003
,
21, 441–443. [CrossRef] [PubMed]
102.
Li
è
vre, A.; Chapusot, C.; Bouvier, A.-M.; Zinzindohou
é
, F.; Piard, F.; Roignot, P.; Arnould, L.; Beaune, P.;
Faivre, J.; Laurent-Puig, P. Clinical value of mitochondrial mutations in colorectal cancer. J. Clin. Oncol.
2005
,
23, 3517–3525. [CrossRef] [PubMed]
103.
Chatterjee, A.; Mambo, E.; Sidransky, D. Mitochondrial DNA mutations in human cancer. Oncogene
2006
,25,
4663–4674. [CrossRef] [PubMed]
104.
Simonnet, H. Low mitochondrial respiratory chain content correlates with tumor aggressiveness in renal cell
carcinoma. Carcinogenesis 2002,23, 759–768. [CrossRef]
105.
Petros, J.A.; Baumann, A.K.; Ruiz-Pesini, E.; Amin,M.B.; Sun, C.Q.; Hall, J.; Lim, S.; Issa, M.M.; Flanders,W.D.;
Hosseini, S.H.; et al. mtDNA mutations increase tumorigenicity in prostate cancer. Proc. Natl. Acad. Sci. USA
2005,102, 719–724. [CrossRef] [PubMed]
Biology 2019,8, 35 14 of 15
106.
Shidara, Y.; Yamagata, K.; Kanamori, T.; Nakano, K.; Kwong, J.Q.; Manfredi, G.; Oda, H.; Ohta, S. Positive
contribution of pathogenic mutations in the mitochondrial genome to the promotion of cancer by prevention
from apoptosis. Cancer Res. 2005,65, 1655–1663. [CrossRef]
107.
Ralevic, V.; Burnstock, G. Receptors for purines and pyrimidines. Pharmacol. Rev.
1998
,50, 413–492.
[PubMed]
108. Smith, J.L. Enzymes of nucleotide synthesis. Curr. Opin. Struct. Biol. 1995,5, 752–757. [CrossRef]
109.
Fang, J.; Uchiumi, T.; Yagi, M.; Matsumoto, S.; Amamoto, R.; Takazaki, S.; Yamaza, H.; Nonaka, K.; Kang, D.
Dihydro-orotate dehydrogenase is physically associated with the respiratory complex and its loss leads to
mitochondrial dysfunction. Biosci. Rep. 2013,33, e00021. [CrossRef]
110.
Bader, B.; Knecht, W.; Fries, M.; Löffler, M. Expression, purification, and characterization of histidine-tagged
rat and human flavoenzyme dihydroorotate dehydrogenase. Protein Expr. Purif.
1998
,13, 414–422. [CrossRef]
111.
Löffler, M.; Jöckel, J.; Schuster, G.; Becker, C. Dihydroorotat-ubiquinone oxidoreductase links mitochondria
in the biosynthesis of pyrimidine nucleotides. Mol. Cell. Biochem. 1997,174, 125–129. [CrossRef]
112.
Beuneu, C.; Auger, R.; Löffler, M.; Guissani, A.; Lemaire, G.; Lepoivre, M. Indirect inhibition of mitochondrial
dihydroorotate dehydrogenase activity by nitric oxide. Free Radic. Biol. Med.
2000
,28, 1206–1213. [CrossRef]
113.
Rawls, J.; Knecht, W.; Diekert, K.; Lill, R.; Löffler, M. Requirements for the mitochondrial import and
localization of dihydroorotate dehydrogenase. Eur. J. Biochem. 2000,267, 2079–2087. [CrossRef]
114.
Uhlin, U.; Eklund, H. Structure of ribonucleotide reductase protein R1. Nature
1994
,370, 533–539. [CrossRef]
115.
Jordan, A.; Reichard, P. Ribonucleotide reductases. Annu. Rev. Biochem.
1998
,67, 71–98. [CrossRef] [PubMed]
116.
Kashlan, O.B.; Scott, C.P.; Lear, J.D.; Cooperman, B.S. A comprehensive model for the allosteric regulation of
mammalian ribonucleotide reductase. Functional consequences of ATP- and dATP-induced oligomerization
of the large subunit. Biochemistry 2002,41, 462–474. [CrossRef]
117.
Chabes, A.L.; Pfleger, C.M.; Kirschner, M.W.; Thelander, L. Mouse ribonucleotide reductase R2 protein:
A new target for anaphase-promoting complex-Cdh1-mediated proteolysis. Proc. Natl. Acad. Sci. USA
2003
,
100, 3925–3929. [CrossRef] [PubMed]
118.
Munch-Petersen, B.; Cloos, L.; Jensen, H.K.; Tyrsted, G. Human thymidine kinase 1. Regulation in normal
and malignant cells. Adv. Enzyme Regul. 1995,35, 69–89. [CrossRef]
119.
Goldstone, D.C.; Ennis-Adeniran, V.; Hedden, J.J.; Groom, H.C.T.; Rice, G.I.; Christodoulou, E.; Walker, P.A.;
Kelly, G.; Haire, L.F.; Yap, M.W.; et al. HIV-1 restriction factor SAMHD1 is a deoxynucleoside triphosphate
triphosphohydrolase. Nature 2011,480, 379–382. [CrossRef]
120.
Clifford, R.; Louis, T.; Robbe, P.; Ackroyd, S.; Burns, A.; Timbs, A.T.; Wright Colopy, G.; Dreau, H.; Sigaux, F.;
Judde, J.G.; et al. SAMHD1 is mutated recurrently in chronic lymphocytic leukemia and is involved in
response to DNA damage. Blood 2014,123, 1021–1031. [CrossRef]
121.
Song, S.; Wheeler, L.J.; Mathews, C.K. Deoxyribonucleotide pool imbalance stimulates deletions in HeLa cell
mitochondrial DNA. J. Biol. Chem. 2003,278, 43893–43896. [CrossRef]
122.
L
ó
pez, L.C.; Akman, H.O.; Garc
í
a-Cazorla, A.; Dorado, B.; Mart
í
, R.; Nishino, I.; Tadesse, S.; Pizzorno, G.;
Shungu, D.; Bonilla, E.; et al. Unbalanced deoxynucleotide pools cause mitochondrial DNA instability in
thymidine phosphorylase-deficient mice. Hum. Mol. Genet. 2009,18, 714–722. [CrossRef] [PubMed]
123.
Pontarin, G.; Gallinaro, L.; Ferraro, P.; Reichard, P.; Bianchi, V. Origins of mitochondrial thymidine
triphosphate: Dynamic relations to cytosolic pools. Proc. Natl. Acad. Sci. USA
2003
,100, 12159–12164.
[CrossRef] [PubMed]
124.
Desler, C.; Munch-Petersen, B.; Rasmussen, L.J. The Role of Mitochondrial dNTP Levels in Cells with
Reduced TK2 Activity. Nucleosides Nucleotides Nucleic Acids 2006,25, 1171–1175. [CrossRef] [PubMed]
125.
Pontarin, G.; Ferraro, P.; Bee, L.; Reichard, P.; Bianchi, V. Mammalian ribonucleotide reductase subunit p53R2
is required for mitochondrial DNA replication and DNA repair in quiescent cells. Proc. Natl. Acad. Sci. USA
2012,109, 13302–13307. [CrossRef]
126.
Bourdon, A.; Minai, L.; Serre, V.; Jais, J.-P.; Sarzi, E.; Aubert, S.; Chr
é
tien, D.; de Lonlay,P.; Paquis-Flucklinger, V.;
Arakawa, H.; et al. Mutation of RRM2B, encoding p53-controlled ribonucleotide reductase (p53R2), causes
severe mitochondrial DNA depletion. Nat. Genet. 2007,39, 776–780. [CrossRef] [PubMed]
127.
Pontarin, G.; Ferraro, P.; Rampazzo, C.; Kollberg, G.; Holme, E.; Reichard, P.; Bianchi, V. Deoxyribonucleotide
metabolism in cycling and resting human fibroblasts with a missense mutation in p53R2, a subunit of
ribonucleotide reductase. J. Biol. Chem. 2011,286, 11132–11140. [CrossRef] [PubMed]
Biology 2019,8, 35 15 of 15
128.
Tanaka, H.; Arakawa, H.; Yamaguchi, T.; Shiraishi, K.; Fukuda, S.; Matsui, K.; Takei, Y.; Nakamura, Y. A
ribonucleotide reductase gene involved in a p53-dependent cell-cycle checkpoint for DNA damage. Nature
2000,404, 42–49. [CrossRef]
129. James, S.J.; Miller, B.J.; McGarrity, L.J.; Morris, S.M. The effect of folic acid and/or methionine deficiency on
deoxyribonucleotide pools and cell cycle distribution in mitogen-stimulated rat lymphocytes. Cell Prolif.
1994,27, 395–406. [CrossRef]
130.
Micheli, V.; Camici, M.; G Tozzi, M.; L Ipata, P.; Sestini, S.; Bertelli, M.; Pompucci, G. Neurological Disorders
of Purine and Pyrimidine Metabolism. Curr. Top. Med. Chem. 2011,11, 923–947. [CrossRef]
131.
Desler, C.; Marcker, M.L.; Singh, K.K.; Rasmussen, L.J. The importance of mitochondrial DNA in aging and
cancer. J. Aging Res. 2011,2011, 407536. [CrossRef] [PubMed]
132.
Desler, C.; Frederiksen, J.H.; Angleys, M.; Maynard, S.; Keijzers, G.; Fagerlund, B.; Mortensen, E.L.; Osler, M.;
Lauritzen, M.; Bohr, V.A.; et al. Increased deoxythymidine triphosphate levels is a feature of relative cognitive
decline. Mitochondrion 2015,25, 34–37. [CrossRef] [PubMed]
133.
Maynard, S.; Hejl, A.-M.; Dinh, T.-S.T.; Keijzers, G.; Hansen, Å.M.; Desler, C.; Moreno-Villanueva, M.;
Bürkle, A.; Rasmussen, L.J.; Waldemar, G.; et al. Defective mitochondrial respiration, altered dNTP pools and
reduced AP endonuclease 1 activity in peripheral blood mononuclear cells of Alzheimer’s disease patients.
Aging 2015,7, 793–815. [CrossRef] [PubMed]
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