Open access peer-reviewed chapter

Role of Subcellular ROS in Providing Resilience to Vascular Endothelium

Written By

Sarah R. Aldosari, Maan A. Awad, Frank W. Sellke and Md. Ruhul Abid

Submitted: August 4th, 2020 Reviewed: August 10th, 2020 Published: September 21st, 2020

DOI: 10.5772/intechopen.93568

Chapter metrics overview

482 Chapter Downloads

View Full Metrics


For decades, elevated levels of reactive oxygen species (ROS) have been associated with the pathogenesis of cardiovascular diseases (CVD), including myocardial ischemia and infarction (MI). However, several large clinical trials failed to demonstrate beneficial outcomes in response to the global reduction of ROS in patients with underlying CVD. Recent studies from our and other labs showed that it is rather a critical balance between mitochondrial and cytosolic ROS than total ROS levels which determines resilience of coronary endothelial cells (EC). Here, we will discuss published and unpublished work that has helped elucidate the molecular mechanisms by which subcellular ROS levels, duration and localization modulate metabolic pathways, including glycolysis and oxidative phosphorylation, energy production and utilization, and dNTP synthesis in EC. These redox-regulated processes play critical roles in providing resilience to EC which in turn help protect existing coronary vessels and induce coronary angiogenesis to improve post-MI recovery of cardiac function.


  • endothelial cell metabolism
  • angiogenesis
  • vascular endothelial growth factor (VEGF)
  • nitric oxide
  • reactive oxygen species (ROS)
  • glycolysis
  • dNTP
  • fatty acid oxidation

1. Introduction

A single layer of endothelial cells (ECs) that covers the vascular lumen and plexus exhibits great plasticity to adapt to environmental cues [1, 2]. It is fascinating how the vascular system, the largest organ system of the body, connects all organs to secure adequate nutrients and blood supply. For that reason, maintaining vascular homeostasis is crucial for the health of the cardiovascular system. In a healthy body, although the ECs are an intricate, dynamic system, they appear to be in a quiescent state [1]. In pathological conditions such as ischemia and infarction, ECs rapidly switch phenotype to form new vessels in a process known as sprouting angiogenesis [3]. Reactive oxygen species (ROS) are believed to play crucial roles in determining the phenotype and fate of EC in both physiological and pathological conditions. Recent work has shown that a critical balance between mitochondrial and cytosolic ROS levels, but not global ROS levels, modulates endothelial function, EC metabolism and angiogenesis, and thus determines resilience of coronary EC [4, 5, 6, 7, 8]. In this chapter, we will discuss the molecular mechanisms by which subcellular ROS modulate various metabolic pathways, regulate EC function and angiogenesis.


2. Reactive oxygen species in coronary endothelium

Previously, the pathology of cardiovascular diseases (CVD), including myocardial ischemia and infarction (MI), was believed to be associated with increased levels of ROS [4, 5, 6, 7, 8]. Recent studies show that it is rather a critical balance between cytosolic and mitochondrial ROS levels than total ROS levels which determine the resilience of coronary ECs in physiological as well as adverse conditions [6, 7, 9, 10].

ROS are produced in higher levels as a response to injuries by the cellular enzymes and mitochondria [8, 11]. ROS have been reported to contribute to the underlying pathology in almost all organs, and thus the notion that antioxidants would ameliorate pathological effects of ROS came into being. However, clinical trials failed to show beneficial effects of antioxidants in the treatment of CVD [12]. Other studies showed that decreased ROS levels had rather deleterious effects on CVD [6, 7]. Also, decreased ROS levels resulted in the inactivation of endothelial nitric oxide synthase (eNOS) and reduction in NO (nitric oxide) levels [11, 13]. Taken together, global reduction of ROS appeared to reduce endothelial resilience. It is crucial to note that ROS have paradoxical effects on ECs, and thus careful study of the levels, durations and sources of ROS while studying effects of ROS on EC will help advance our understanding of EC resilience during oxidative stress.

2.1 Source of reactive oxygen species in ECs

ROS are produced from different oxidoreductase enzymes and locations including NADPH oxidase, mitochondrial, xanthine oxidase, cytochrome P450 monooxygenase, and uncoupling of NOS [8, 11, 14]. In the vasculature, ECs rely on glycolytic pathways as their source of energy, thus NADPH oxidase enzymes appear to be the major source of ROS in both physiological and pathological conditions [14]. NADPH enzymes have different isoforms, and the major contributors are NOX1, NOX2, NOX4, and NOX5 [15, 16]. Recent studies showed importance of NOX2- and NOX4-derived ROS in endothelial survival or dysfunction, depending on their subcellular location and duration [8].

2.2 Endothelial NADPH oxidase as a major source of ROS in ECs

NADPH oxidase is an intracellular complex enzyme containing membrane-bound and cytosolic regulator subunits [14, 17, 18]. This enzyme produces ROS by transferring electrons from NAD(P)H to an oxygen molecule and is considered the major source of ROS in coronary endothelium. Distinct isoforms of NADPH enzymes have been shown to exhibit different physiological and pathological responses in vascular homeostasis.

NOX1 enzyme is primarily expressed in the vascular smooth muscle cells (VSMC) and it contributes to VSMC proliferation and migration [11, 19, 20, 21]. In disease conditions, NOX1 contributes to the impairment of endothelium-dependent vasorelaxation, as well as the augmentation of angiotensin II vasomotor response [11, 22, 23]. A study showed that NOX1-deficient mice attenuated the levels of ROS, neointimal growth, and migration. These findings suggest that the downregulation of NOX1 enzyme can prevent the formation of atherosclerotic plaque [15, 24]. Yet, further studies are warranted to explore the exact role of NOX1 in endothelial signaling.

In contrast, NOX2 enzyme has exhibited positive effects on coronary ECs. NOX2 enzyme stimulates the production of NO by the activation of AMPK-eNOS axis through Ca2+−/calmodulin-dependent protein kinase kinase β (CaMKKβ) [6] resulting in coronary vasodilation, EC proliferation and migration. Although several studies support the beneficial effects of NOX2, they also exert detrimental effects on coronary EC depending on the duration of exposure. Short exposure of elevated ROS levels was associated with the previously mentioned pathway (i.e. CaMKKβ pathway). On the contrary, prolonged exposure of high ROS levels resulted in decrease bioavailability of NO, inactivation of mitochondrial antioxidant MnSOD [7, 8], and decreased EC proliferation and coronary vasodilatation.

NOX4 enzyme is abundant in human ECs [8] and produces H2O2 molecules rather than O2 [9, 11, 25, 26, 27, 28]. NOX4 enzyme stimulates vascular angiogenesis through the activation of transforming growth factor β1 (TGF β1), and increases hemoglobin content [29]. NOX4-derived ROS cause vasodilation through endothelium hyperpolarization [30, 31, 32]. This occurs via the stimulation of endothelium Ca2+-activated K+ channel that causes the release of Ca2+ from the endoplasmic reticulum [29]. Additionally, NOX4 enzyme activates heme oxygenase-1 (HO-1), which confers a vascular protective response via different mechanisms [29]. Thus, therapeutic modalities that advocate for antioxidants in CVD needs careful consideration of the source and location of ROS.

Calcium-dependent NADPH oxidase, NOX5, is implicated in angiogenic response [33, 34]. It gets its name from its structure because it has an additional N-terminal region that binds to calcium [33]. This unique structure allows the enzyme activation through increased intracellular calcium. Similar to NOX4, NOX5 enzyme seems to produce predominantly H2O2 in ECs [24]. H2O2 has been implicated in the development of atherosclerotic plaque plausibly by increasing Ca2+ levels to promote eNOS-mediated NO synthesis and increasing nitroxide radicals [24]. One mechanism may include increased consumption of NO by ROS. Thus, it has been hypothesized that inhibition of NOX5 enzyme may show beneficial results by precluding oxidant injury to vascular EC.

NADPH enzyme isoforms have distinct locations and EC phenotypes. They have been shown to employ different physiological and pathological responses in vascular homeostasis. As discussed above, NOX1, NOX2, NOX4, and NOX5 are found in the vascular system and they contribute to endothelial resilience through several mechanisms. The roles of NADPH enzymes in physiological and pathological conditions have undergone a considerable evolution in recent years. However, further studies are necessary to deepen our understanding of their roles and contributions to EC resilience.

2.3 Endothelial mitochondrial ROS

Although oxidative phosphorylation in mitochondria play a major role in synthesizing energy in most tissues, EC primarily depends on anaerobic glycolysis for 85% of its ATP generation. ECs have fewer mitochondria and consume lower amounts of O2 than other cell types, and thus mitochondrial ROS are believed to be a minor source of ROS in EC in physiological conditions. However, recent studies demonstrated that sustained increase in NADPH oxidase-derived cytosolic ROS may affect the levels of mitochondrial ROS and thus mitochondrial function in EC [6, 7, 8].


3. Metabolic pathways in ECs

EC metabolism plays an important role in facilitating cellular proliferation and migration during the process of angiogenesis. Alterations in metabolic pathways are necessary to provide energy supplies in the most efficient way under certain circumstances that induces blood vessel sprouting such as hypoxia. In addition, these alterations mediate the formation of important molecules that are essential for cytoskeletal remodeling during the process. This section highlights some of these metabolic pathways and their role in angiogenesis.

3.1 Glycolysis

Glycolysis is a major metabolic pathway that is utilized for energy production through the anaerobic oxidation of glucose molecules [35, 36]. It is the major source of ATP in ECs. Glycolysis involves consumption of 2 ATP molecules, and the end products include 4 ATP, 2 NADH and 2 pyruvate molecules (Figures 1 and 2). Subsequently, pyruvate can be shifted to the mitochondria and metabolized into acetyl-CoA to be used in the tricarboxylic acid cycle (TCA). The substrates and products of this process are as follows:

Figure 1.

The investment phase of glycolysis and regulation of the rate limiting PFK1 enzyme by fructose-2,6-bisphosphate and AMP.

Figure 2.

The payoff phase of glycolysis.


Glycolysis occurs in the cytosol, and the process does not require oxygen (anaerobic), therefore it constitutes the primary source of energy in cells that lack mitochondria (e.g. red blood cells). In addition, glycolysis is the main source of pyruvate, which is converted to acetyl-CoA to be utilized in the TCA cycle in cells that use oxidative phosphorylation (aerobic respiration) as a primary source of ATP. Also, glycolysis is a more efficient source of energy in periods of hypoxia and ischemia when oxygen supply becomes scarce.

3.1.1 Mechanisms of glycolysis

The first step in glycolysis constitutes the investment phase of glycolysis, in which 2 ATP molecules are consumed as shown in Figure 1. It involves trapping of the glucose molecule inside the cell via phosphorylation into glucose-6-phosphate [35, 36]. This reaction is catalyzed by glucokinase in the liver and pancreatic β cells, or a hexokinase enzyme in the rest of body cells. It also involves the transfer of a phosphate group from an ATP molecule. Next, glucose-6-phosphate is converted to fructose-6-phosphate by an isomerase. This is followed by the rate-limiting step of glycolysis, which involves the phosphorylation of fructose-6-phosphate into fructose-1-6-bisphosphate by phosphofructokinase 1 (PFK1). This step is critical in the glycolytic pathway and the PFK1 enzyme is highly regulated by multiple factors that determine the direction of the reaction. Fructose-1-6-bisphosphate is subsequently converted by an aldolase into dihydroxyacetone phosphate (DHAP) and glyceraldehyde-3-phosphate (G3P).

The following reactions constitute what can be referred to as the payoff phase of glycolysis. It is also important to remember that at this stage, we have two 3-carbon molecules per 1 glucose molecule as shown in Figure 2. G3P is converted to 1-3-diphosphoglycerate, generating NADH in the process. 1-3-diphosphoglycerate then loses a phosphate group to 3-phosphoglycerate via phosphoglycerate kinase, which generates an ATP molecule. 3-phosphoglycerate is subsequently converted in a two-step reaction into phosphoenolpyruvate (PEP). Finally, pyruvate kinase converts PEP into pyruvate, generating ATP in the process. Thus, the end product of glycolysis includes 4 ATP molecules, but because of the initial consumption of 2 ATP, the return on investment includes 2 ATP molecules per glucose [35, 36].

3.1.2 Regulation of glycolysis

The availability of glucose regulates the rate of glycolysis and is determined by two main mechanisms: glucose uptake from the blood, and breakdown of glycogen [35, 37]. In addition, the amount of oxygen can also regulate glycolysis through what is called the “Pasteur Effect”, which describes how increased oxygen levels inhibit glycolysis, and decreased availability results in acceleration of glycolysis [35]. Within the glycolytic pathway, PFK1, which catalyzes the rate limiting step is considered the main player in terms of glycolysis regulation, and its activity can be affected in a number of ways.

Fructose 2–6 bisphosphate is an allosteric regulator of PFK1, which increases the enzyme activity [35, 37]. It is produced by phosphofructokinase 2 (PFK2), an enzyme that has both kinase and phosphorylase activity and can transform fructose 6 phosphates to fructose 2,6 bisphosphate and vice versa. Insulin dephosphorylates PFK2 activating its kinase activity, and increasing fructose 2,6 bisphosphate production, which subsequently activates PFK1 (Figure 1). Moreover, Glucagon phosphorylates PFK2, activating its phosphatase, which transforms fructose 2,6 bisphosphate back to fructose 6 phosphate. This decreases fructose 2,6 bisphosphate levels and decreases PFK1 activity. Low energy levels within the cell which result in increased AMP and low ATP/AMP ratio, induce allosteric activation of PFK1.

3.1.3 Glycolysis and EC angiogenesis

The endothelium is one of the most diverse tissues in the human body, which displays significant organ-specific heterogeneity. This diversity determines the function of the endothelium according to the organ being supplied [38]. Since ECs lining blood vessels are responsible for supplying oxygen and nutrient to body tissues, the ability to expand this network of blood vessels via angiogenesis is critical for organ growth and function in health and disease [39]. Low oxygen levels serve as a primary stimulus for angiogenesis, which in its classic meaning refers to the sprouting of branches from the existing vessels. Angiogenesis

ECs are essential for the normal functioning of the vascular system. They drive the vascular system expansion during physiologic organ growth to supply sufficient nutrients, as well as under pathologic conditions through a process known as angiogenesis (Figure 3). Angiogenesis depends highly on the coordinated orchestra of several regulatory steps [1]. Briefly, this process is guided by the migratory non-proliferative “tip” cells at the forefront from an existing vessel, while the “stalk” cells trail the proliferative and elongation part of the sprout. “Tip” and “stalk” cells continuously switch their phenotype between being either tip or stalk cells. For example, the “tip” cell becomes a “stalk” cell when it loses its migratory behavior, and the “stalk” cell will compete for the position [1]. Several studies found that the vascular endothelial growth factor (VEGF) controls the “tip” cells induction, filopodia formation, and expression of the Notch ligand Delta–like 4 (NLD4) [40, 41]. NLD4, subsequently, suppresses VEGF receptor 2 (VEGFR2/kdr/Flk1) and thus modulates the tip cell behavior. While many genetic and molecular signaling pathways were recognized to be part of this process, the role of ECs metabolism has not been studied and explored until recently.

Figure 3.

Angiogenesis is mainly regulated via VEGF. Tip cells require increasing amounts of ATP necessary for migration into hypoxic tissues while proliferating stalk cells generate building blocks (dNTP, protein) to maintain their growth and cellular division.

Switching on the angiogenic machinery of ECs has significant consequences on EC metabolism. This is because angiogenic ECs require nutrients and energy not only for motility but also for the synthesis of building blocks (proteins, nucleotides, and lipids) for cellular proliferation. Hence, during angiogenesis, ECs must increase their metabolic activity to generate energy quickly, while at the same time meeting the challenge of scarce resources as they proliferate in harsh hypoxic environments. Therefore, EC metabolism has to be flexible to support vessel formation under different conditions [39, 42].

Upon switching from quiescence state to vessel branching, the rate of glycolysis is increased in order to fuel subcellular processes required for migration such as cytoskeleton remodeling. Notably, the pro-angiogenic VEGF increases expression of the glycolysis activator phosphofructokinase-2/fructose-2,6-bisphosphatase 3 (PFKFB3) [43]. PFKFB3 generates higher levels of fructose-2,6-bisphosphate, which activates phosphofructokinase 1, the rate limiting enzyme in glycolysis [43, 44]. In fact, studies have shown that genetic and pharmacologic inhibition of the phosphofructokinase 2 reduced EC sprouting and branching capacity [44, 45]. Another regulator enzyme is the hexokinase-2 (HK2) which phosphorylate glucose to glucose-6-phosphate [44, 46]. Several transcription factors such as KLF2 and forkhead box 1 (FOXO1) were found to suppress these key glycolytic enzymes in the quiescent phalanx cells [47, 48]. However, the rate of glycolysis increases in the actively sprouting tip and stalk cells due to VEGF-mediated activation of PFKFB3 and the decreased levels of KLF2 and FOXO1. Interestingly, PFKFB3 and other glycolytic enzymes are highly concentrated in filopodia to generate ATP at the so-called ‘ATP hot-spots’. And several studies showed that pharmacologic or genetic inhibition PFKFB3 impairs new vessel formation [43, 44]. Endothelial cell metabolism

Despite the fact that oxygen is readily available for EC consumption, the glycolytic pathway remains primary source of energy for EC [38, 44, 49]. In fact, 85% of EC energy production in the form of ATP is generated through glycolysis, even though oxidative phosphorylation (OXPHOS) can generate significantly larger amounts of ATP molecules at much faster rate [43, 49, 50]. However, ECs have fewer mitochondria and consume lower amounts of O2 than other cell types, especially in the presence of abundant supplies of glucose, where only a small fraction of pyruvate is shifted to the TCA cycle [39, 43, 50]. Nonetheless, ECs retain their capacity for oxidative metabolism when glycolysis is compromised or in conditions of stress. Surprisingly, even though the amount of energy generated per glucose molecule via oxidative phosphorylation is significantly greater, higher rates of glycolysis can provide more ATP in a shorter period of time when glucose supply is unlimited. In fact, the rate of glycolysis is high in EC compared to other normal cells, and their glucose consumption is comparable with that of some cancer cells [43].

A logical question that can be asked here is, why do ECs depend on glycolysis for their energy production when they have direct supply of oxygen from blood? There are several explanations for this observation. First, despite the fact that the energy yield via glycolysis is significantly low compared to aerobic respiration, glycolysis can generate ATP molecules at a much faster rate [39, 49]. This is especially important when considering the energy requirements of ECs during angiogenesis. In addition, anaerobic glycolysis facilitates ECs sprouting and proliferation in hypoxic tissues and makes them resistant to hypoxic insults [39]. Also, it limits ROS generation and produces larger amounts of lactic acid, which acts as a pro-angiogenic factor [38, 3944]. Moreover, oxygen can be spared to be utilized by the underlying tissue cells. The low oxygen dependence allows sprouting cells to explore and reach distant hypoxic tissues [44]. Also, low oxygen consumption by ECs facilitates oxygen delivery to vital organs. Furthermore, glycolysis provides essential metabolites that are used in multiple cellular pathways such as pentose phosphate pathway (PPP), hexosamine biosynthesis pathway (HBP) and 3-phosphoglycerate (G3P) which generate important molecules and compounds that are used in different cellular processes [39, 49]. Thus, glycolysis provides a metabolic platform that allows ECs to perform diverse roles in the growing and resting vasculature with minimal ROS generation. Alternative metabolism of glucose

ECs engage in several other pathways that can potentially affect angiogenesis, but their exact roles are understudied. Once phosphorylated by hexokinase (HK), glucose-6-phosphate (G6P) can be used to form glycogen, which could serve as an endogenous source of glucose when ECs sprout into glucose-deprived milieu. In fact, inhibition of glycogen phosphorylase (PYG), was found to impair EC migration [51].

G6P can also enter the pentose phosphate pathway (PPP) to generate NADPH [44]. NADPH is essential for restoring the reduced form of glutathione (GSH) from its oxidized form (GSSG), which serves as an antioxidant [38, 52]. PPP provides two intermediates of glycolysis, fructose-6-phosphate (F6P) and glyceraldehyde-3-phosphate (G3P). Interestingly, inhibition of G6P dehydrogenase (G6PD) or Transketolase (TKT) in the PPP was found to impair EC viability and migration [44]. Pathways regulation

ECs react to environmental conditions and energy requirements through several mechanisms that involve cellular molecules sensing changes in energy levels. One of these molecules is the AMP-kinase (AMPK), which gets activated by the rising levels of AMP as energy levels dwindle. Activation of AMPK-mediated phosphorylation of metabolic targets promotes catabolic pathways and ATP production, while inhibiting anabolic pathways that consume ATP [39, 53]. This allows ECs to balance their energy level according to environmental changes. For instance, AMPK increases energy production via fatty acid oxidation (FAO) in EC mitochondria and help maintain ATP levels when glucose supplies are low [39, 54]. In addition, AMPK is activated by EC-specific stimuli such as hypoxia and shear stress generated by blood flow [38]. Interestingly, inhibition of AMPK was found to hinder EC angiogenesis in response to hypoxia [55].

3.2 Oxidative phosphorylation

The mechanism by which ATP is produced in the mitochondria via oxidative phosphorylation (OxPhos) was first discovered in the second half of the twentieth century [56, 57]. OxPhos is a process that involves the use of high-energy intermediates for energy transduction between the electron transport chain of the mitochondria and the chemical synthesis of ATP from ADP and phosphate. OxPhos generates 15 times the amount of ATP produced by glycolysis during anaerobic conditions. The reaction involves oxygen consumption, and energy is released from the high energy molecules (NADH, FADH2) and stored in the form of an electrochemical proton gradient across the inner mitochondrial membrane. This energy extraction occurs in three steps each catalyzed by a specific membrane complex including Complex I (NADH dehydrogenase), Complex III (Cytochrome bc1) and Complex IV (Cytochrome oxidase/COX). Complex II (Succinate dehydrogenase) converts succinate to fumarate, a TCA cycle intermediate, and in the process H+ is produced from FADH2, which is then shunted by Complex III across the inner mitochondrial membrane. COX is also considered the rate-limiting step of this aerobic respiration. Eventually, the electrochemical proton gradient is utilized by Complex V (ATP Synthase) to produce ATP, or it can be dissipated in the form of heat by passive proton leakage [56, 58, 59].

The electron transport chain is regulated through different mechanisms. Allosteric effectors such as ADP and ATP regulate the process by binding to their specific binding sites on the different mitochondrial complexes. Regulation of the enzyme activity by ATP or ADP binding to the same site on the complex subunit depends on the ATP/ADP ratio. For instance, the exchange of bound ADP by ATP on COX results in an allosteric ATP synthesis inhibition at an ATP/ADP ratio of 28 [60]. In addition, phosphorylation and dephosphorylation of the enzyme complexes is considered another mean of regulating the electron transport chain. For example, phosphorylation of COX was found to inhibit the enzyme activity [56].

3.3 Fatty acid oxidation contribute to dNTP synthesis

Deoxyribonucleoside Triphosphate (dNTP) is a molecule consisting of a deoxyribose sugar attached to three phosphate groups and one of the nucleotide bases, adenine, guanine, cytosine, or thymine as shown in Figure 4 [61]. Apart from DNA replication, dNTPs may also function as a source of energy for different cellular reactions and signaling pathways [62].

Figure 4.

Structure of Deoxyribonucleoside Triphosphate (dNTP).

3.3.1 dNTP formation

There are two biosynthetic pathways for nucleotides formation: de novo and salvage [62]. The de novo pathways require high energy and the use of raw material like glucose, glutamine, aspartate, and HCO3 to form nucleotides [62, 63]. However, salvage pathways exist as an alternative energy-efficient route to form nucleotides [63].

The enzyme ribonucleotide reductase (RR), which is NADPH-dependent, is responsible for catalyzing the rate-limiting reaction in which ribonucleotides are converted to their respective deoxyribonucleotides [62, 63]. This reaction is regulated by the number of RR enzymes and allosteric control mechanism [62, 63]. RR consists of two nonidentical subunits, α and β. α subunit has the catalytic site, substrate-specificity site, and activity site; whereas the β subunit contains a stable tyrosyl free radical [63]. The activity of RR enzymes is tightly controlled by allosteric mechanism [63, 64]. The reduction of ribonucleotides requires a specific positive effector, however, the product dNTP can also serve as a negative effector on the enzyme (Table 1) [61, 63].

Positive EffectordGTPdTTPATPATP
Negative EffectordATPdATPdATP

Table 1.

Ribonucleotide reductase enzyme regulators.

dNTPs levels and RR enzyme activity are important to control the fidelity of nuclear and mitochondrial DNA replication and repair. It has been reported that increased levels of dNTP, in vitro, decreased the length of ‘S phase’ of the cell cycle during DNA replication, which implies that under physiological conditions, nucleotides are used mainly for DNA synthesis [65, 66]. Interestingly, whereas elevated levels of dNTP resulted in delay in the S phase entry through unclear mechanisms [67, 68], depletion of dNTP pool also resulted in inhibition of DNA replication, and fork stalling [69]. In fact, when the enzyme RR was blocked, DNA synthesis was arrested, preserving the dNTP for DNA damage repair under suboptimal conditions [69, 70].

3.3.2 Mitochondrial dNTP

Mitochondria are one of the major endomembrane organelles in eukaryotic cells [14, 71, 72] owing to their ability to produce ATP through oxidative phosphorylation as discussed in Section 3.2. Yet they participate in cellular function and dysfunction, including calcium regulation, activation of cellular death, ROS formation, and cellular building block synthesis [73, 74]. In ECs, the mitochondria compromise only 6% of cell volume, implicating that EC rely on anaerobic glycolysis rather than mitochondria-derived energy [7, 71]. However, mitochondria act primarily as major signaling organelles in the ECs and maintain mitochondrial dNTP pools for proper EC functions. Additionally, alternation in the levels of mitochondrial ROS has been shown to be associated with impaired one-carbon metabolism, which is essential for purines and pyrimidines nucleotides [75, 76].

In response to mild oxidative stress, the mitochondria attempt to re-establish homeostasis by ROS-buffering capacity of mitochondria. For example, the activity of adenine nucleotide translocase is impaired under mitochondrial oxidation, leading to shortage of adenine diphosphate (ADP) [77]. On the other hand, up-regulation of mitochondrial anti-oxidant systems and other molecules counteract ROS-induced protein unfolding [78, 79]. If oxidative stress is persistent, mitochondria may translate the adaptive response into activation of cellular death [74]. These responses and deregulation of ROS levels contribute to the pathogenesis of cardiovascular system, including coronary artery diseases.

3.3.3 Fatty acid oxidation (FAO)

Long chain fatty acids are a major source of energy productions, primarily in mitochondria [61, 80, 81]. Fatty acids are broken up into acetyl CoA, NADH and FADH2 in the mitochondria [80]. These three products are used by the mitochondrial matrix for energy production through TCA and oxidative phosphorylation [80]. Fatty acid oxidation as a major energy-producing pathway

Fatty acid oxidation (FAO) is an important catabolic and anabolic process. On the outer membrane of mitochondria, FAO transfers the acyl group from CoA to carnitine by carnitine palmitoyltransferase I (CPT1). Acyl-carnitine is then exchanged across the inner membrane of mitochondria. The acyl group is transferred back again to CoA by carnitine palmitoyltransferase II (CPT II) as shown in Figure 5 [82]. CPT1 is an important enzyme for FAO and is a rate limiting factor for FAO in the mitochondria. Malonyl CoA, an intermediate product of fatty acid synthesis, is an inhibitor of CPT1.

Figure 5.

Long-chain fatty acid transportation in the mitochondra. Fatty acids are transported through the mitochondrial membrane as acyl CoA for subsequent oxidation. Malonyl CoA acts as a key inhibitor molecule for CPT I, and thus regulating the rate of fatty acid oxidation (FAO).

β-oxidation is a four steps process carried by enzymatic oxidation, hydration, and oxidation that act on acyl CoA to yield a shorter acyl CoA and acetyl CoA [83]. The four-step process is shown in the schematics of Figure 6.

Figure 6.

Fatty acid β-oxidation pathway. Role of fatty acid oxidation in vessel sprouting

Recent studies have shown the critical role of FAO for vessel sprouting [42, 84]. In a study, the levels of FAO and dNTP synthesis were reduced when mitochondrial CPT1A was silenced. This resulted in impaired vascular sprouting due to reduction ECs proliferation but not migration. Additionally, silencing long-chain acyl-CoA dehydrogenase (ACADVL) has yielded similar results, supporting the role of FAO in vessel sprouting. Overexpression of CPT1A obtained opposite results, further supporting a crucial role of CPT1A in angiogenesis. Fatty acid oxidation for de novo synthesis of nucleotides

As noted above, silencing CPT1A in ECs showed impaired de novo synthesis of dNTPs. This impaired de novo dNTP synthesis contributed to reduced vessel sprouting [82]. Nonetheless, the levels of glucose oxidation were increased to compensate for the FAO loss, yet it was not sufficient to help in the proliferative defects of ECs with knockdown CPT1A. This reflects the irreplaceable role of FAO for de novo dNTP synthesis in ECs [82]. Fatty acid β-oxidation in quiescent vs. proliferating endothelial cells

Depending on the cellular status, the FAO are directed either toward DNA synthesis or redox homeostasis. FAO are involved in regenerating NADP+ to NADPH, they also upregulate the expression of NADP+ producing genes, which are critical for redox homeostasis [43, 82]. Quiescent ECs upregulate FAO, but do not rely on them for ATP production or nucleotide synthesis, rather utilize it for redox homeostasis [85]. Unlike quiescent ECs, proliferating ECs utilize FAO for DNA synthesis, as previously discussed [82].

CPT1A, the rate limiting enzyme for FAO in mitochondria, has been shown to be critical for redox homeostasis in EC. In quiescent ECs, CPT1A inhibition caused the levels of ROS to elevate, leading to decreased anti-fibrinolytic gene expression, endothelial leakage, and increased leukocytes adhesion and/or infiltration [80, 85]. Thus, it is believed that quiescent ECs require more redox buffering capacity compared to proliferating ECs due to higher levels of ROS.

Besides the involvement of FAO in redox balance in quiescent ECs, they are also involved in other vasculo-protective NADPH-regenerating pathways such as oxidative PPP and nicotine nucleotide transhydrogenase [85].

4. Endothelial metabolism in atherosclerosis

The generation of increased amounts of NO in atherosclerosis is critical for its anti-atherogenic effects, including vasodilation, inhibition of platelet aggregation, smooth muscle proliferation as well as leukocyte migration and oxidative stress [38]. Endothelial cells produce NO through enzymatic oxidation of arginine to citrulline via eNOS enzyme. eNOS requires several co-factors including NADPH, flavin adenine dineucleotide (FAD), flavin mononeucleotide (FMN), Calcium/Calmodulin and tetrahydrobiopterin (BH4). Decreased availability of Arginine or deficiency of BH4 results in the paradoxical generation of ROS instead of NO by eNOS, a process known as eNOS uncoupling [38, 86]. Arginine in particular, has been found to be rate-limiting for NO synthesis in patients with atherosclerosis [87]. It was demonstrated that an arginine analog asymmetric dimethyl arginine (ADMA), that acts as a competitor for eNOS, impaired NO production. ADMA levels are markedly increased in atherosclerosis and therefore it is recognized as a major cardiovascular risk factor [88]. Moreover, Dimethyl arginine dimethyl aminohydrolase (DDAH), an enzyme that metabolizes ADMA into citrulline and dimethylamine is impaired by the oxidative stress in atherosclerosis [38]. Interestingly, because of this competition, Arginine supplements have been found to be of great benefit in atherosclerotic patients with high ADMA levels, by enhancing endothelial-dependent vasodilation and inhibition of leukocyte adhesion and migration to the atherosclerotic plaque [89].

Furthermore, endothelial NADPH oxidase is induced by certain atherosclerotic plaque components such as the oxidized LDL (oxLDL). The NADPH oxidase-derived ROS were found to have detrimental effects in promoting plaque progression. These include oxidation of LDLs, inducing vascular smooth muscle proliferation and migration and EC apoptosis as well as promoting the expression of vascular adhesion adhesion molecules [38].


5. Conclusions

The endothelium is one of the most diverse tissues in the human body. It maintains the integrity of the vascular system and provides nutrition to underlying tissues. In addition, EC drives the growth and proliferation of blood vessels under physiologic and pathologic conditions. ECs exhibit significant flexibility in response to various environmental changes such as hypoxia and ischemia. Careful analysis of the process of sprouting angiogenesis explains how ECs function in such an orchestrated way to reach their end goal of providing nutrients and oxygen supply to the affected tissues. ECs display phenomenal resilience in the process through various mechanisms, one of which is their metabolic adaptation and the other is critical balance between subcellular levels of ROS (cytosolic versus mitochondrial). ECs limit their oxygen consumption in order to preserve it for the tissues that they supply to and also to maintain a balanced intracellular redox state. Although ECs do not utilize mitochondrial OxPhos for ATP synthesis and thus generate very little mitochondrial ROS, NADPH oxidase-derived ROS appear to regulate many critical EC functions in health and disease. However, EC has intricated intracellular mechanisms by which subcellular oxidants may communicate at the subcellular levels [7]. Unlike most cells in the body (except tumor cells), ECs upregulate and accelerate their glycolytic pathways in order to generate energy (ATP production) and certain molecules that act as building blocks (dNTPs) and are essential for supporting EC proliferation and migration. CPT1A-mediated FAO appears to play a significant role in synthesizing dNTPs and NADP+, NADPH in EC mitochondria. Depending on the metabolic states of ECs (quiescent versus proliferative), FAO-generated NADPH is utilized for quiescent EC’s redox homeostasis or dNTPS for cell proliferation in angiogenic endothelium. Further studies aimed at understanding the molecular mechanisms by which subcellular ROS modulate EC metabolism in health and disease will help develop therapeutics modalities for CVD.



This work was supported by the National Heart, Lung, and Blood Institute (NHLBI) 1R01HL133624 (M.R.A.); R01HL46716 and R01HL128831-01A1 (F.W.S).


Conflict of interest

The authors declare no conflict of interest.


  1. 1. Vandekeere S, Dewerchin M, Carmeliet P. Angiogenesis revisited: An overlooked role of endothelial cell metabolism in vessel sprouting. Microcirculation. 2015;22(7):509-517. DOI: 10.1111/micc.12229
  2. 2. Cantelmo AR, Brajic A, Carmeliet P. Endothelial metabolism driving angiogenesis: Emerging concepts and principles. The Cancer Journal. 2015;21(4):244-249. DOI: 10.1097/PPO.0000000000000133
  3. 3. Eelen G, Cruys B, Welti J, De Bock K, Carmeliet P. Control of vessel sprouting by genetic and metabolic determinants. Trends in Endocrinology and Metabolism: TEM. 2013;24(12):589-596. DOI: 10.1016/j.tem.2013.08.006
  4. 4. Ruiz-Lozano P, Rajan P. Stem cells as in vitro models of disease. Current Stem Cell Research & Therapy. 2007;2(4):280-292. DOI: 10.2174/157488807782793772
  5. 5. Fearon IM, Gaça MD, Nordskog BK. In vitro models for assessing the potential cardiovascular disease risk associated with cigarette smoking. oxicology in Vitro: An International Journal Published in Association with BIBRA. 2013;27(1):513-522. DOI: 10.1016/j.tiv.2012.08.018
  6. 6. Shafique E, Choy WC, Liu Y, Feng J, Cordeiro B, Lyra A, et al. Oxidative stress improves coronary endothelial function through activation of the pro-survival kinase AMPK. Aging. 2013;5(7):515-530
  7. 7. Shafique E, Torina A, Reichert K, Colantuono B, Nur N, Zeeshan K, et al. Mitochondrial redox plays a critical role in the paradoxical effects of NAPDH oxidase-derived ROS on coronary endothelium. Cardiovascular Research. 2017;113(2):234-246. DOI: 10.1093/cvr/cvw249
  8. 8. Aldosari S, Awad M, Harrington EO, Sellke FW, Abid MR. Subcellular reactive oxygen species (ROS) in cardiovascular pathophysiology. Antioxidants. 2018;7(1):14. DOI: 10.3390/antiox7010014
  9. 9. Kim YM, Kim SJ, Tatsunami R, Yamamura H, Fukai T, Ushio-Fukai M. ROS-induced ROS release orchestrated by Nox4, Nox2, and mitochondria in VEGF signaling and angiogenesis. American Journal of Physiology. Cell Physiology. 2017;312(6):C749-C764. DOI: 10.1152/ajpcell.00346.2016
  10. 10. Schröder K, Zhang M, Benkhoff S, Mieth A, Pliquett R, Kosowski J, et al. Nox4 is a protective reactive oxygen species generating vascular NADPH oxidase. Circulation Research. 2012;110(9):1217-1225. DOI: 10.1161/CIRCRESAHA.112.267054
  11. 11. Awad MA, Aldosari SR, Abid MR. Genetic alterations in oxidant and anti-oxidant enzymes in the vascular system. Frontiers in Cardiovascular Medicine. 2018;5:107. DOI: 10.3389/fcvm.2018.00107
  12. 12. Sleight P. The HOPE study (heart outcomes prevention evaluation). Journal of the Renin-Angiotensin-Aldosterone System. 2000;1(1):18-20. DOI: 10.3317/jraas.2000.002
  13. 13. Ponnuswamy P, Schröttle A, Ostermeier E, Grüner S, Huang PL, Ertl G, et al. eNOS protects from atherosclerosis despite relevant superoxide production by the enzyme in apoE mice. PLoS One. 2012;7(1):e30193. DOI: 10.1371/journal.pone.0030193
  14. 14. Aldosari S, Awad M, Gao MZ, McCormack IG, Sellke FW, Abid MR. Oxidant-dependent and oxidant-independent proangiogenic and vasomotor signaling in coronary vascular endothelium. In: Chakraborti S, Dhalla N, Dikshit M, Ganguly N, editors. Modulation of Oxidative Stress in Heart Disease. Singapore: Springer; 2019
  15. 15. Frey RS, Ushio-Fukai M, Malik AB. NADPH oxidase-dependent signaling in endothelial cells: Role in physiology and pathophysiology. Antioxidants & Redox Signaling. 2009;11(4):791-810. DOI: 10.1089/ars.2008.2220
  16. 16. Manuneedhi Cholan P, Cartland SP, Kavurma MM. NADPH oxidases, angiogenesis, and peripheral artery disease. Antioxidants. 2017;6(3):56. DOI: 10.3390/antiox6030056
  17. 17. Bedard K, Krause KH. The NOX family of ROS-generating NADPH oxidases: Physiology and pathophysiology. Physiological Reviews. 2007;87(1):245-313. DOI: 10.1152/physrev.00044.2005
  18. 18. Feng J, Damrauer SM, Lee M, Sellke FW, Ferran C, Abid MR. Endothelium-dependent coronary vasodilatation requires NADPH oxidase-derived reactive oxygen species. Arteriosclerosis, Thrombosis, and Vascular Biology. 2010;30(9):1703-1710. DOI: 10.1161/ATVBAHA.110.209726
  19. 19. Szöcs K, Lassègue B, Sorescu D, Hilenski LL, Valppu L, Couse TL, et al. Upregulation of Nox-based NAD(P)H oxidases in restenosis after carotid injury. Arteriosclerosis, Thrombosis, and Vascular Biology. 2002;22(1):21-27. DOI: 10.1161/hq0102.102189
  20. 20. Schröder K, Helmcke I, Palfi K, Krause KH, Busse R, Brandes RP. Nox1 mediates basic fibroblast growth factor-induced migration of vascular smooth muscle cells. Arteriosclerosis, Thrombosis, and Vascular Biology. 2007;27(8):1736-1743. DOI: 10.1161/ATVBAHA.107.142117
  21. 21. Sheehan AL, Carrell S, Johnson B, Stanic B, Banfi B, Miller FJ Jr. Role for Nox1 NADPH oxidase in atherosclerosis. Atherosclerosis. 2011;216(2):321-326. DOI: 10.1016/j.atherosclerosis.2011.02.028
  22. 22. Mollnau H, Wendt M, Szöcs K, Lassègue B, Schulz E, Oelze M, et al. Effects of angiotensin II infusion on the expression and function of NAD(P)H oxidase and components of nitric oxide/cGMP signaling. Circulation Research. 2002;90(4):E58-E65. DOI: 10.1161/01.res.0000012569.55432.02
  23. 23. Gavazzi G, Banfi B, Deffert C, Fiette L, Schappi M, Herrmann F, et al. Decreased blood pressure in NOX1-deficient mice. FEBS Letters. 2006;580(2):497-504. DOI: 10.1016/j.febslet.2005.12.049
  24. 24. Guzik TJ, Chen W, Gongora MC, Guzik B, Lob HE, Mangalat D, et al. Calcium-dependent NOX5 nicotinamide adenine dinucleotide phosphate oxidase contributes to vascular oxidative stress in human coronary artery disease. Journal of the American College of Cardiology. 2008;52(22):1803-1809. DOI: 10.1016/j.jacc.2008.07.063
  25. 25. Bendall JK, Rinze R, Adlam D, Tatham AL, de Bono J, Wilson N, et al. Endothelial Nox2 overexpression potentiates vascular oxidative stress and hemodynamic response to angiotensin II: Studies in endothelial-targeted Nox2 transgenic mice. Circulation Research. 2007;100(7):1016-1025. DOI: 10.1161/01.RES.0000263381.83835.7b
  26. 26. Chen L, Hou X, Xiao J, Kuroda J, Ago T, Sadoshima J, et al. Both hydrogen peroxide and transforming growth factor beta 1 contribute to endothelial Nox4 mediated angiogenesis in endothelial Nox4 transgenic mouse lines. Biochimica et Biophysica Acta (BBA) - Molecular Basis of Disease. 2014;1842(12):2489-2499. DOI: 10.1016/j.bbadis.2014.10.007
  27. 27. Datla SR, Peshavariya H, Dusting GJ, Mahadev K, Goldstein BJ, Jiang F. Important role of Nox4 type NADPH oxidase in angiogenic responses in human microvascular endothelial cells in vitro. Arteriosclerosis, Thrombosis, and Vascular Biology. 2007;27(11):2319-2324. DOI: 10.1161/ATVBAHA.107.149450
  28. 28. Ray R, Murdoch CE, Wang M, Santos CX, Zhang M, Alom-Ruiz S, et al. Endothelial Nox4 NADPH oxidase enhances vasodilatation and reduces blood pressure in vivo. Arteriosclerosis, Thrombosis, and Vascular Biology. 2011;31(6):1368-1376. DOI: 10.1161/ATVBAHA.110.219238
  29. 29. Craige SM, Chen K, Pei Y, Li C, Huang X, Chen C, et al. NADPH oxidase 4 promotes endothelial angiogenesis through endothelial nitric oxide synthase activation. Circulation. 2011;124(6):731-740. DOI: 10.1161/CIRCULATIONAHA.111.030775
  30. 30. Shimokawa H, Morikawa K. Hydrogen peroxide is an endothelium-derived hyperpolarizing factor in animals and humans. Journal of Molecular and Cellular Cardiology. 2005;39(5):725-732. DOI: 10.1016/j.yjmcc.2005.07.007
  31. 31. Matoba T, Shimokawa H, Nakashima M, Hirakawa Y, Mukai Y, Hirano K, et al. Hydrogen peroxide is an endothelium-derived hyperpolarizing factor in mice. The Journal of Clinical Investigation. 2000;106(12):1521-1530. DOI: 10.1172/JCI10506
  32. 32. Miura H, Bosnjak JJ, Ning G, Saito T, Miura M, Gutterman DD. Role for hydrogen peroxide in flow-induced dilation of human coronary arterioles. Circulation Research. 2003;92(2):e31-e40. DOI: 10.1161/01.res.0000054200.44505.ab
  33. 33. BelAiba RS, Djordjevic T, Petry A, Diemer K, Bonello S, Banfi B, et al. NOX5 variants are functionally active in endothelial cells. Free Radical Biology & Medicine. 2007;42(4):446-459. DOI: 10.1016/j.freeradbiomed.2006.10.054
  34. 34. Herkert O, Djordjevic T, BelAiba RS, Görlach A. Insights into the redox control of blood coagulation: Role of vascular NADPH oxidase-derived reactive oxygen species in the thrombogenic cycle. Antioxidants and Redox Signaling. 2004;6(4):765-776. DOI: 10.1089/1523086041361695
  35. 35. Chaudhry R, Varacallo M. Biochemistry, Glycolysis. StatPearls. 2018. Available from:
  36. 36. Naifeh J, Varacallo M. Biochemistry, Aerobic Glycolysis. StatPearls. 2019. Available from:
  37. 37. Jois T, Sleeman MW. The regulation and role of carbohydrate response element-binding protein in metabolic homeostasis and disease. Journal of Neuroendocrinology. 2017;29(10). DOI: 10.1111/jne.12473
  38. 38. Eelen G, de Zeeuw P, Simons M, Carmeliet P. Endothelial cell metabolism in normal and diseased vasculature. Circulation Research. 2015;116(7):1231-1244. DOI: 10.1161/CIRCRESAHA.116.302855
  39. 39. Potente M, Carmeliet P. The link between angiogenesis and endothelial metabolism. Annual Review of Physiology. 2017;79(1):43-66. DOI: 10.1146/annurev-physiol-021115-105134
  40. 40. Potente M, Gerhardt H, Carmeliet P. Basic and therapeutic aspects of angiogenesis. Cell. 2011;146(6):873-887. DOI: 10.1016/j.cell.2011.08.039
  41. 41. Jakobsson L, Franco CA, Bentley K, Collins RT, Ponsioen B, Aspalter IM, et al. Endothelial cells dynamically compete for the tip cell position during angiogenic sprouting. Nature Cell Biology. 2010;12(10):943-953. DOI: 10.1038/ncb2103
  42. 42. De Bock K, Georgiadou M, Carmeliet P. Role of endothelial cell metabolism in vessel sprouting. Cell Metabolism. 2013;18(5):634-647. DOI: 10.1016/j.cmet.2013.08.001
  43. 43. De Bock K, Georgiadou M, Schoors S, Kuchnio A, Wong BW, Cantelmo AR, et al. Role of PFKFB3-driven glycolysis in vessel sprouting. Cell. 2013;154(3):651-663. DOI: 10.1016/j.cell.2013.06.037
  44. 44. Rohlenova K, Veys K, Miranda-Santos I, De Bock K, Carmeliet P. Endothelial cell metabolism in health and disease. Trends in Cell Biology. 2018;28(3):224-236. DOI: 10.1016/j.tcb.2017.10.010
  45. 45. Schoors S, De Bock K, Cantelmo AR, Georgiadou M, Ghesquière B, Cauwenberghs S, et al. Partial and transient reduction of glycolysis by PFKFB3 blockade reduces pathological angiogenesis. Cell Metabolism. 2014;19(1):37-48. DOI: 10.1016/j.cmet.2013.11.008
  46. 46. Yu P, Wilhelm K, Dubrac A, Tung JK, Alves TC, Fang JS, et al. FGF-dependent metabolic control of vascular development. Nature. 2017;545(7653):224-241. DOI: 10.1038/nature22322
  47. 47. Doddaballapur A, Michalik KM, Manavski Y, Lucas T, Houtkooper RH, You X, et al. Laminar shear stress inhibits endothelial cell metabolism via KLF2-mediated repression of PFKFB3. Arteriosclerosis, Thrombosis, and Vascular Biology. 2015;35(1):137-145. DOI: 10.1161/ATVBAHA.114.304277
  48. 48. Wilhelm K, Happel K, Eelen G, Schoors S, Oellerich MF, Lim R, et al. FOXO1 couples metabolic activity and growth state in the vascular endothelium. Nature. 2016;529(7585):216-220. DOI: 10.1038/nature16498
  49. 49. Heiden MGV, Cantley LC, Thompson CB. Understanding the Warburg effect: The metabolic requirements of cell proliferation. Science. 2009;324(5930):1029-1033. DOI: 10.1126/science.1160809
  50. 50. Ghesquière B, Wong BW, Kuchnio A, Carmeliet P. Metabolism of stromal and immune cells in health and disease. Nature. 2014;511:167-176. DOI: 10.1038/nature13312
  51. 51. Vizán P, Sánchez-Tena S, Alcarraz-Vizán G, Soler M, Messeguer R, Pujol MD, et al. Characterization of the metabolic changes underlying growth factor angiogenic activation: Identification of new potential therapeutic targets. Carcinogenesis. 2009;30(6):946-952. DOI: 10.1093/carcin/bgp083
  52. 52. Riganti C, Gazzano E, Polimeni M, Aldieri E, Ghigo D. The pentose phosphate pathway: An antioxidant defense and a crossroad in tumor cell fate. Free Radical Biology & Medicine. 2012;53(3):421-436. DOI: 10.1016/j.freeradbiomed.2012.05.006
  53. 53. Hardie DG, Schaffer BE, Brunet A. AMPK: An energy-sensing pathway with multiple inputs and outputs. Trends in Cell Biology. 2016;26(3):190-201. DOI: 10.1016/j.tcb.2015.10.013
  54. 54. Dagher Z, Ruderman N, Tornheim K, Ido Y. The effect of AMP-activated protein kinase and its activator AICAR on the metabolism of human umbilical vein endothelial cells. Biochemical and Biophysical Research Communications. 1999;265(1):112-115. DOI: 10.1006/bbrc.1999.1635
  55. 55. Nagata D, Mogi M, Walsh K. AMP-activated protein kinase (AMPK) signaling in endothelial cells is essential for angiogenesis in response to hypoxic stress. Journal of Biological Chemistry. 2003;278(33):31000-31006. DOI: 10.1074/jbc.M300643200
  56. 56. Kadenbach B. Mitochondrial Oxidative Phosphorylation: Nuclear-Encoded Genes, Enzyme Regulation, and Pathophysiology. New York: Springer; 2012
  57. 57. Boyer PD. Conformational coupling in oxidative phosphorylation and photophosphorylation. Trends in Biochemical Sciences. 1977;2(2):38-41. DOI: 10.1016/0968-0004(77)90254-7
  58. 58. Rees DM, Leslie AG, Walker JE. The structure of the membrane extrinsic region of bovine ATP synthase. Proceedings of the National Academy of Sciences of the United States of America. 2009;106(51):21597-21601. DOI: 10.1073/pnas.0910365106
  59. 59. Jastroch M, Divakaruni AS, Mookerjee S, Treberg JR, Brand MD. Mitochondrial proton and electron leaks. Essays in Biochemistry. 2010;47:53-67. DOI: 10.1042/bse0470053
  60. 60. Arnold S, Kadenbach B. Cell respiration is controlled by ATP, an allosteric inhibitor of cytochrome-c oxidase. European Journal of Biochemistry. 1997;249(1):350-354. DOI: 10.1111/j.1432-1033.1997.t01-1-00350.x
  61. 61. Devlin TM. Textbook of Biochemistry with Clinical Correlation. New York: John Wiley and Sons; 1982
  62. 62. Lane AN, Fan TW. Regulation of mammalian nucleotide metabolism and biosynthesis. Nucleic Acids Research. 2015;43(4):2466-2485. DOI: 10.1093/nar/gkv047
  63. 63. Nordlund P, Reichard P. Ribonucleotide reductases. Annual Review of Biochemistry. 2006;75:681-706. DOI: 10.1146/annurev.biochem.75.103004.142443
  64. 64. Fairman JW, Wijerathna SR, Ahmad MF, Xu H, Nakano R, Jha S, et al. Structural basis for allosteric regulation of human ribonucleotide reductase by nucleotide-induced oligomerization. Nature Structural and Molecular Biology. 2011;18(3):316-322. DOI: 10.1038/nsmb.2007
  65. 65. Stodola JL, Burgers PM. Resolving individual steps of Okazaki-fragment maturation at a millisecond timescale. Nature Structural & Molecular Biology. 2016;23(5):402-408. DOI: 10.1038/nsmb.3207
  66. 66. Kunkel TA, Sabatino RD, Bambara RA. Exonucleolytic proofreading by calf thymus DNA polymerase delta. Proceedings of the National Academy of Sciences of the United States of America. 1987;84(14):4865-4869. DOI: 10.1073/pnas.84.14.4865
  67. 67. Chabes A, Stillman B. Constitutively high dNTP concentration inhibits cell cycle progression and the DNA damage checkpoint in yeast Saccharomyces cerevisiae. Proceedings of the National Academy of Sciences of the United States of America. 2007;104(4):1183-1188. DOI: 10.1073/pnas.0610585104
  68. 68. Franzolin E, Pontarin G, Rampazzo C, Miazzi C, Ferraro P, Palumbo E, et al. The deoxynucleotide triphosphohydrolase SAMHD1 is a major regulator of DNA precursor pools in mammalian cells. Proceedings of the National Academy of Sciences of the United States of America. 2013;110(35):14272-14277. DOI: 10.1073/pnas.1312033110
  69. 69. Koç A, Wheeler LJ, Mathews CK, Merrill GF. Hydroxyurea arrests DNA replication by a mechanism that preserves basal dNTP pools. The Journal of Biological Chemistry. 2004;279(1):223-230. DOI: 10.1074/jbc.M303952200
  70. 70. Aye Y, Li M, Long MJ, Weiss RS. Ribonucleotide reductase and cancer: Biological mechanisms and targeted therapies. Oncogene. 2015;34(16):2011-2021. DOI: 10.1038/onc.2014.155
  71. 71. Friedman JR, Nunnari J. Mitochondrial form and function. Nature. 2014;505(7483):335-343. DOI: 10.1038/nature12985
  72. 72. Dromparis P, Michelakis ED. Mitochondria in vascular health and disease. Annual Review of Physiology. 2013;75:95-126. DOI: 10.1146/annurev-physiol-030212-183804
  73. 73. Osellame LD, Blacker TS, Duchen MR. Cellular and molecular mechanisms of mitochondrial function. Best practice & research. The Journal of Clinical Endocrinology and Metabolism. 2012;26(6):711-723. DOI: 10.1016/j.beem.2012.05.003
  74. 74. Galluzzi L, Kepp O, Trojel-Hansen C, Kroemer G. Mitochondrial control of cellular life, stress, and death. Circulation Research. 2012;111(9):1198-1207. DOI: 10.1161/CIRCRESAHA.112.268946
  75. 75. Bao XR, Ong SE, Goldberger O, Peng J, Sharma R, Thompson DA, et al. Mitochondrial dysfunction remodels one-carbon metabolism in human cells. eLife. 2016;5:e10575. DOI: 10.7554/eLife.10575
  76. 76. Nikkanen J, Forsström S, Euro L, Paetau I, Kohnz RA, Wang L, et al. Mitochondrial DNA replication defects disturb cellular dNTP pools and remodel one-carbon metabolism. Cell Metabolism. 2016;23(4):635-648. DOI: 10.1016/j.cmet.2016.01.019
  77. 77. Pestana CR, Silva CH, Uyemura SA, Santos AC, Curti C. Impact of adenosine nucleotide translocase (ANT) proline isomerization on Ca2+−induced cysteine relative mobility/mitochondrial permeability transition pore. Journal of Bioenergetics and Biomembranes. 2010;42(4):329-335. DOI: 10.1007/s10863-010-9297-4
  78. 78. Sharma S, Dewald O, Adrogue J, Salazar RL, Razeghi P, Crapo JD, et al. Induction of antioxidant gene expression in a mouse model of ischemic cardiomyopathy is dependent on reactive oxygen species. Free Radical Biology & Medicine. 2006;40(12):2223-2231. DOI: 10.1016/j.freeradbiomed.2006.02.019
  79. 79. Singh M, Sharma H, Singh N. Hydrogen peroxide induces apoptosis in HeLa cells through mitochondrial pathway. Mitochondrion. 2007;7(6):367-373. DOI: 10.1016/j.mito.2007.07.003
  80. 80. Patella F, Schug ZT, Persi E, Neilson LJ, Erami Z, Avanzato D, et al. Proteomics-based metabolic modeling reveals that fatty acid oxidation (FAO) controls endothelial cell (EC) permeability. Molecular & Cellular Proteomics. 2015;14(3):621-634. DOI: 10.1074/mcp.M114.045575
  81. 81. Dagher Z, Ruderman N, Tornheim K, Ido Y. Acute regulation of fatty acid oxidation and amp-activated protein kinase in human umbilical vein endothelial cells. Circulation Research. 2001;88(12):1276-1282. DOI: 10.1161/hh1201.092998
  82. 82. Schoors S, Bruning U, Missiaen R, Queiroz KC, Borgers G, Elia I, et al. Fatty acid carbon is essential for dNTP synthesis in endothelial cells. Nature. 2015;520(7546):192-197. DOI: 10.1038/nature14362
  83. 83. Hülsmann WC, Dubelaar ML. Aspects of fatty acid metabolism in vascular endothelial cells. Biochimie. 1988;70(5):681-686. DOI: 10.1016/0300-9084(88)90253-2
  84. 84. Teuwen LA, Draoui N, Dubois C, Carmeliet P. Endothelial cell metabolism: An update anno 2017. Current Opinion in Hematology. 2017;24(3):240-247. DOI: 10.1097/MOH.0000000000000335
  85. 85. Kalucka J, Bierhansl L, Conchinha NV, Missiaen R, Elia I, Brüning U, et al. Quiescent endothelial cells upregulate fatty acid β-oxidation for vasculoprotection via redox homeostasis. Cell Metabolism. 2018;28(6):881-894.e13. DOI: 10.1016/j.cmet.2018.07.016
  86. 86. Kawashima S. Malfunction of vascular control in lifestyle-related diseases: Endothelial nitric oxide (NO) synthase/NO system in atherosclerosis. Journal of Pharmacological Sciences. 2004;96(4):411-419. DOI: 10.1254/jphs.fmj04006x6
  87. 87. Böger RH, Bode-Böger SM, Szuba A, Tsao PS, Chan JR, Tangphao O, et al. Asymmetric dimethylarginine (ADMA): A novel risk factor for endothelial dysfunction - Its role in hypercholesterolemia. Circulation. 1998;98(18):1842-1847. DOI: 10.1161/01.CIR.98.18.1842
  88. 88. Böger RH. Asymmetric dimethylarginine: Understanding the physiology, genetics, and clinical relevance of this novel biomarker. In: Proceedings of the 4th International Symposium on ADMA. Pharmacological Research; 2009. DOI: 10.1016/j.phrs.2009.10.001
  89. 89. Chan JR, Böger RH, Bode-Böger SM, Tangphao O, Tsao PS, Blaschke TF, et al. Asymmetric dimethylarginine increases mononuclear cell adhesiveness in hypercholesterolemic humans. Arteriosclerosis, Thrombosis, and Vascular Biology. 2000;20(4):1040-1046. DOI: 10.1161/01.ATV.20.4.1040

Written By

Sarah R. Aldosari, Maan A. Awad, Frank W. Sellke and Md. Ruhul Abid

Submitted: August 4th, 2020 Reviewed: August 10th, 2020 Published: September 21st, 2020