Lipopolysaccharide, inflammation and mitochondria-targeted antioxidants: History
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Subjects: Pharmacology & Pharmacy
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Rimma Parnova

Lipopolysaccharide (LPS), the major component of the outer membrane of Gram-negative bacteria, is the most abundant proinflammatory agent. Considerable evidence indicates that LPS challenge inescapably causes oxidative stress and mitochondrial dysfunction, leading to cell and tissue damage. Increased mitochondrial reactive oxygen species (mtROS) generation triggered by LPS is known to play a key role in the progression of the inflammatory response. mtROS at excessive levels impair electron transport chain functioning, reduce the mitochondrial membrane potential, and initiate lipid peroxidation and oxidative damage of mitochondrial proteins and mtDNA. Over the past 20 years, a large number of mitochondria-targeted antioxidants (mito-AOX) of different structures that can accumulate inside mitochondria and scavenge free radicals have been synthesized. Their protective role based on the prevention of oxidative stress and the restoration of mitochondrial function has been demonstrated in a variety of common diseases and pathological states.

  • mitochondria-targeted antioxidants
  • inflammation
  • LPS

1. Mechanisms of LPS-Triggered Inflammation

Inflammatory reactions play a critical role in LPS-induced tissue injury, which is caused by the adhesion and migration of leucocytes through the epithelium, the production of a variety of proinflammatory mediators by monocytes/macrophages, and oxidative stress driven by excess generation of reactive oxygen and nitrogen species. As the most abundant proinflammatory agent, LPS activates the systemic and cellular inflammatory response largely due to signaling through Toll-like receptor 4 (TLR4), which is expressed not only in immune cells but also in almost all cell types [1][2][3]. In mammalian cells, LPS-induced activation of TLR4 occurs through a series of interactions with several adapter proteins to facilitate its recognition by the TLR4/MD-2 receptor complex, which causes TLR4 oligomerization and recruitment of its numerous downstream adaptors. LPS/TLR4 signaling can be divided into MyD88-dependent and MyD88-independent (TRIF-dependent) pathways, leading to nuclear transcription factors NF-κB-, AP-1-, and IRF3-mediated induction of a wide range of pro- and anti-inflammatory mediators, such as cytokines, chemokines, eicosanoids, and others [4][5]. Stimulation of immune, epithelial, endothelial, and other cells by pro-inflammatory mediators results in excessive ROS/RNS generation, which is potentially damaging to all cellular compartments.

Recently, TLR4-independent LPS sensing pathways have been described [6]. First, transient receptor potential (TRP) channels have been identified as non-TLR membrane-bound sensors of LPS, and second, caspase-4/5 (and caspase-11 in mice) have been established as cytoplasmic sensors for LPS [6].

LPS has been shown to activate inflammasomes—cytosolic multiprotein complexes that mediate the propagation of the inflammatory response within the innate and the adaptive immune system as well as in epithelial cells [7][8][9][10]. Inflammasome activation is evoked by extra- and intracellular pathogens, such as Gram-negative bacteria, and/or by danger signals to activate caspase-1, resulting in a caspase-1-dependent, highly inflammatory form of cell death—pyroptosis [9][10]. It is thought that in Gram-negative infections, pyroptosis plays a key role in the destruction of intracellular bacterial replication niches. Inflammasomes are responsible for the maturation of proinflammatory cytokines, including IL-1β and IL-18, and for their secretion through the formation of pores in the plasma membrane [9][11][12]. The most studied inflammasome, NLRP3, is implicated in many different pathologies such as cancer and metabolic, neurodegenerative, and inflammatory diseases. Activation of the NLRP3 inflammasome occurs in two steps via different activating signals. Through TLR4 activation, LPS participates in the priming of inflammasome activation to initiate the transcription of NLRP3, inactive pro-IL-1β, and pro-IL-18 via NF-κB [11]. The second signals, which are notably diverse, include mtROS, derived mainly from ETC electron leakage [11], and initiate assembly of the inflammasome complex and, among other things, maturation of pro-IL-1β and pro-IL-18.

2. Generation of mtROS and Their Role in Normal and Pathological Conditions

ROS is a general term encompassing oxygen-free radicals, including superoxide ion (O2−) and hydroxyl radical (·OH), and nonradical oxygen substrates, e.g., hydrogen peroxide (H2O2) and singlet oxygen (1O2). ROS are produced during a wide range of biochemical reactions within the cell and within different cell compartments (mitochondria, peroxisomes, and endoplasmic reticulum). There are many cell proteins, bearing thiols, catecholamines, hydroquinones, and flavins, that may participate in intracellular ROS production. Although ROS can be produced in the cytosol by NADPH oxidases and xanthine oxidase, the major sources of ROS are mitochondria [13][14]. Simultaneously, mitochondria are a major target for damages by their own ROS.

Mitochondria can contain more than a dozen enzymatic sources of mtROS [15][16]. However, there is a consensus now that the mitochondrial respiratory chain is a primary source of mtROS, which are generated by the leakage of electrons from the ETC, resulting in a partial reduction in molecular oxygen. Mitochondrial complexes I (NADH-ubiquinone oxidoreductase) and III (cytochrome C oxidoreductase) are assumed to be a predominant source of superoxide generation under pathological conditions [13][17][18][19]. In particular, superoxide is generated by the reaction of O2 with electrons originating either from direct transport from NADH under conditions of a slow respiration rate (due to low ATP demand or mitochondrial damage) or from the reverse electron transport (RET) from complex II (succinate dehydrogenase) to complex I under conditions of a high protonmotive force or when there is a high NADH/NAD+ ratio in the matrix [13][17][18][19].

In healthy tissues under physiological conditions, mtROS generation and their removal by endogenous scavenging compounds and enzymes are tightly controlled by a complex antioxidant defense network including superoxide dismutase (SOD), catalase, ascorbic acid, tocopherol, reduced glutathione (GSH), etc. SOD catalyzes the conversion of superoxide to hydrogen peroxide, which, in turn, is converted to water by glutathione peroxidase (GPx). Under oxidative stress, the expression of antioxidant genes is activated by translocation of nuclear factor E2-related factor 2 (Nrf2) from the cytoplasm to the nucleus [20][21].

mtROS are necessary for the cell due to their essential role in various intracellular signaling pathways, including mitochondrial quality control by autophagy [22][23]. Disturbance of the fine-tuning of the equilibrium between mtROS generation and scavenging leads to excessive ROS production, resulting in severe damage of single cells and whole organs, loss of function, and then organism failure [24][25]. Numerous pathological conditions and diseases such as sepsis, cancer, metabolic diseases, neurodegenerative diseases, and others are triggered or accompanied by an increased level of mtROS [26][27][28]. At the cellular level, the most harmful effects of mtROS are associated with oxidation-triggered sustained damage of mitochondrial nucleic acids, proteins, and lipids, resulting in the impairment of ETC functioning and ATP production [28][29][30][31][32][33]. For example, mtROS-induced oxidation of cardiolipin polyunsaturated fatty acids damages cristae curvatures, reduces ETC complex activity, initiates cardiolipin translocation from the inner to the outer mitochondrial membrane, and triggers cytochrome C release into the cytosol, which is critical for the mitochondrial cell death pathway [34]. Furthermore, mtROS can oxidize proteins of the mitochondrial permeability transition pore, enhancing cytochrome C and mitochondrial DNA (mtDNA) release. mtDNA is especially sensitive to damage by ROS due to a lack of protective histones and its proximity to the ETC [35][36]. mtROS-initiated release of mtDNA is an important trigger of systemic inflammation, which is recognized by cells as a virulent pathogen motive [32]. Inflammatory conditions are commonly accompanied by excessive NO production, which, in turn, can interact with superoxide to generate the highly toxic peroxynitrite (ONOO-) [37][38].

3. LPS Triggers mtROS Generation

Enhancement of ROS generated in both the cytosol and the mitochondria is an important step in LPS signaling that links the activation of TLR4 with the NF-kB-driven expression of proinflammatory mediators. Numerous animal models of sepsis (LPS or CLP) have demonstrated common abnormalities: increased level of ROS, decreased antioxidant capacity, and mitochondrial oxidative damage [37][39][40]. LPS-triggered generation of mitochondrial superoxide measured usually with the fluorogenic dye MitoSOX was demonstrated in different cell types, such as microglia [41][42], muscle myoblasts [43], gingival fibroblasts [44], human pulmonary bronchial epithelial cells [45], macrophages [46], and others. In addition, the decline in antioxidative enzymes and glutathione content caused by LPS contributes to the impairment of endogenous antioxidant defense and the subsequent increase in mtROS generation [47][48][49][50][51].

In a wide range of cell types, LPS application disturbs cellular energetics, which manifests itself in a decline in respiratory complex activity, decline in the mitochondrial membrane potential, reduction in mitochondrial respiration, and suppression of ATP production in a tissue-, time-, and dose-dependent manner [52][53][54][55][56][57][58][59][60]. A critical step in these disturbances is associated with excessive ROS generation [61][54][62][13][14]. In innate immune cells, LPS has been shown to switch the metabolic reprogramming from oxidative phosphorylation to aerobic glycolysis as a survival response maintaining the cellular ATP level [41][46][63][64], which results in the slowing or reversing of electron transport through respiratory complex I and a subsequent increase in ROS production.

The intrinsic mechanisms linking TLR4 signaling and mtROS have been studied mainly in phagocytic cells such as macrophages in which both cytosolic and mitochondrial ROS generation is also related to their bactericidal activity [65][66]. Thus, it has been shown that LPS-triggered induction of mtROS is mediated by a complex I-associated protein evolutionarily conserved signaling intermediate in Toll pathways (ECSIT), which plays a key role in complex I assembly and stability [66][67]. Upon LPS stimulation, the TLR signaling adapter tumor necrosis factor receptor-associated factor 6 (TRAF6) is translocated to the mitochondria and ubiquitinates ECSIT, resulting in its enrichment at the mitochondrial periphery, thus leading to the augmentation of mitochondrial and cellular ROS generation [66]. TRAF6 also mediates the ubiquitination of the small GTPase Rac, maintaining it in an active GTP-loaded state, which is necessary for the full activation of the ROS-producing machinery [68]. In addition, it was demonstrated in macrophages that LPS activates mitochondrial mitofusin2 (Mfn2) expression, which has been shown to be a required step for mtROS generation. Mfn2−/− macrophages are not be able to produce mtROS and proinflammatory mediators in response to LPS [69]. Whether the above mechanisms exist in other cell types remains unknown.

4. Mitochondria-Targeted Antioxidants and Their Application against LPS-Triggered Inflammation

Generally, antioxidants are substances that can accept or donate electron(s) to eliminate the unpaired condition of free radicals, thus neutralizing them. Antioxidant drugs can either scavenge free radicals or turn into new free radicals, which in some cases are less active and dangerous than initial ones [70]. Antioxidants can also break chain reactions [71] as well as affect ROS-regulated enzymes through controlling the cellular level of free radicals [72][73][74][75]. The natural defense mechanisms are supplied by enzymatic and non-enzymatic antioxidants, which are distributed within the cell cytoplasm and organelles (such as SOD, GPx and reductase, catalase, vitamins, minerals, polyphenols, albumin, transferrin, ferritin, and a variety of others), whereas foods and supplements provide a wide variety of exogenous natural ones (e.g., vitamins B, C, and E; ions Zn, Cu, and Se; flavonoids; omega-3 fatty acids; L-carnitine; and Q-enzyme Q10).

4.1. Conjugates with Lipophilic Cations

An era of investigations applying mito-AOX conjugates began half a century ago when V. Skulachev and colleagues demonstrated the ability of lipophilic cations, such as triphenylphosphonium (TPP+), to accumulate within mitochondria due to the large MMP (negative inside) [76]. In numerous chimeric mito-AOX compounds synthesized later, lipophilic cations, which provide drug accumulation several hundred-fold in the mitochondrial matrix, were grafted to antioxidant moieties, which quenched electrons from the respiratory chain, thus diminishing ROS elevation [77]. Conjugates designed in such a way are widely used as a tool for research, as well as for diagnostic and therapeutic purposes, including drug delivery (for review, see [78][79].

Triphenylphosphonium derivatives have been mainly used as a mitochondrial targeting moiety. They are conjugated with quinone derivatives (ubiquinone in MitoQs [80] or plastoquinone in SkQs [14][81]); with superoxide dismutase and catalase mimetics in MitoTEMPO and MitoTEMPOL, respectively [82]; and with vitamin E in MitoVitE [83], etc. Inside mitochondria, these chimeric mito-AOX undergo red/ox cycling: they not only quench radicals but also can be reduced afterward by the ETC.

Most of the in vivo studies and clinical trials were performed with MitoQ and SkQ1. As both compounds are found to be localized at the matrix-facing side of the inner mitochondrial membrane with their antioxidant portion and alkyl chain, their main protective activity is to prevent lipid peroxidation [84][85]. TPP-based mito-AOX accumulate preferentially in healthy and hyperpolarized mitochondria but not within injured mitochondria, which carry a lower membrane potential and therefore are capable of taking up lower doses of therapeutic antioxidants than normal ones [86][87].

TPP-based antioxidants have been widely used in both in vitro experiments on various types of cells exposed to LPS (Table 1) and in vivo experiments on typical animal models of inflammation or sepsis (administration of LPS or cecal ligation and puncture; Table 2). Each cellular model mimics a specific pathological state or disorder associated with mtROS-induced inflammation. Among examples in Table 1 is LPS- or bacteria-induced mitochondrial dysfunction in oligodendrocytes (model of multiple sclerosis) [85], renal tubular cells (pyelonephritis and acute kidney injury) [88][89], microglia (neurodegeneration) [42], hepatocytes (liver failure) [90], endothelial cells (vascular abnormalities) [60][91], muscle myoblasts (diaphragm weakness) [43], gingival fibroblasts (periodontitis) [44], intestinal epithelial cells (impaired gut barrier function) [20], and others.

Table 1. In vitro effects of mitochondria-targeted antioxidants.

Inflammatory Model Cells Mito-AOX Major Findings Reference
LPS E. coli Macrophages MitoQ
50-100 nM		 ↓ Cellular ROS; ↓ IL-1β mRNA and protein expression [65]
LPS E. coli BV-2 murine microglial cells MitoTEMPO
200 µM		 ↓ Mitochondrial and cellular ROS; ↓ iNOS and COX-2 expression; ↓ TNF-α, IL-1β, IL-6 content; ↓ NF-ḵB activation [42]
LPS E. coli Primary cultured frog urinary bladder epithelial cells MitoQ
25 nM		 ↓ Cellular ROS; prevention of fatty acid oxidation decline and lipid droplet accumulation [56]
LPS E. coli Intestinal epithelial cell line-6 (IEC-6) MitoQ
1 µM		 Stimulation of nuclear translocation of Nrf2 [20]
LPS E. coli NRK-52E (rat renal proximal tubular cell line) MitoQ
1 µM		 ↓ Cellular ROS; ↓NLRP3 inflammasome activation; ↓ IL-1β, IL-18, and caspase-1 [89]
Mixture of cytokines + LPS E. coli C2C12 muscle myoblasts MitoTEMPOL
10 mg/L		 ↓ Mitochondrial superoxide generation; prevention of reduction in cell width [43]
LPS E. coli + PepG S. aureus HUVEC-C MitoQ
1 µM		 ↓ Cellular ROS; restoration of MMP; ↓ IL-1β, IL-6, and IL-8 [91]
LPS E. coli + PepG S. aureus HUVEC-C Melatonin, 0.1, 1.0, 10, 100, and 500 µM; MitoVitE, 5 µM ↓ IL-6 and IL-8; ↓ NF-ḵB activation; ↓ loss of MMP; ↑ GSH level; ↓ decline in metabolic activity [48][92]
LPS P. gingivalis Human gingival fibroblasts MitoTEMPO
50 µM		 ↓ Mitochondrial ROS; ↓ IL-6, IL-1β, and TNF-α production; ↓ activation of NF-B [44]
Inflammatory mediators generated by incubation of white blood cells with LPS E. coli Buffalo rat liver cell line-3A (BRL-3A) MitoTEMPO,
500 nM		 ↓ Mitochondrial and cellular ROS; ↓iNOS mRNA; ↓ IL-6 [90]
LPS E. coli + IFN-γ + TNFα HUVEC Mitoquinone (MQ)
1 µM		 ↓ Cellular ROS; ↓ tyrosine nitration and iNOS protein expression; recovery of O2 consumption and complex I activity [60]
LPS E. coli + succinate Bone marrow-derived macrophages Dimethyl malonate, 10 mM; MitoQ, 500 nM; MitoTEMPO, 0.5–1 mM ↓ Cellular ROS, IL-1β, and HIF-1α; ↑ IL-1RA and IL-10 [46]
E. coli lysate + activated leucocytes Primary culture rat kidney cells SKQR1 10 nM ↓ Cellular ROS; ↓ cell dearth [88]
LPS E. coli Primary oligodendrocytes SkQ1 5-10 nM Restoration of myelin synthesis [85]
LPS E. coli Cardiomyocytes Melatonin 100 μM ↓ Cellular ROS; ↓ loss of MMP; ↑ content of GSH, SOD; ↓ decline in BAP31 expression; ↑ cell viability [47]
LPS E. coli Primary neonatal rat cardiomyocytes Melatonin 100 μM ↓ IL-6, TNF-α, mRNA levels; ↓Bax and ↑ Bcl-2 expression; ↑ autophagy; [93]
LPS E. coli Human alveolar epithelial cells Melatonin, 800 μM ↓ cellular ROS; ↓ MDA; ↑ SOD and GPx levels; prevention of LPS-induced epithelial–mesenchymal transition through Nrf2 activation [49]
LPS E. coli Macrophages RAW 264.7 Mn-porphyrin-oligopeptide conjugate, 10 µM ↓ LPS-induced cell dearth [94]
LPS E. coli Cardiomyocytes (H9C2 cell line) SS-31, 10 μM ↓ ROS; ↓ MDA;
↓ mRNA level of IL-6, IL-1β, and TNF-α;
normalized activity of GPx and SOD;
↓ MMP decline; ↑ ATP		 [50]
LPS E. coli Murine microglial cells (BV-2) SS-31, 100 nM ↓ ROS; effect is mediated by Fis1;
↓ Fis1 expression;
↓ COX-2 and iNOS expression		 [95]
LPS E. coli Macrophages RAW 264.7 XJB-5-131 2 µmol/kg ↓ NO and inflammatory cytokines [96]
E. coli 0157:H7 Human colonic epithelial cell line (Caco-2) MitoTEMPO ↓ Cellular ROS; ↓NLRP3 inflammasome activation; ↓ IL-1β and IL-18 [97]

Table 2. In vivo effects of mitochondria-targeted antioxidants.

Model of Infection Species Organ Investigated Mito-AOX Mode of Antioxidant Application Major Findings Reference
LPS E. coli Rat, mouse Heart MitoQ
500 μM		 Given water orally for 2 days ↓ Oxidative stress; ↓ mitochondrial dysfunction; ↓ cardiac TNF-α level; ↓ reductions in cardiac pressure generation; ↓ caspase 3 and 9 activity [98]
LPS E. coli Mouse Gut, serum MitoQ
4 mg/kg		 i.v. injection 15 min before LPS ↓ Gut barrier dysfunction, restoration of the level of tight junction proteins (ZO-1 and occludin); ↓ intestinal inflammatory response; ↑ SOD and GSH level; ↓TNF-α, IL-1, IL-6, and NO in intestines and plasma [20]
LPS E. coli Rat Liver,
serum		 MitoTEMPO 50 nmol/kg;
SKQ1 5 nmol/kg		 i.p., 24 and 1 h before LPS ↓ iNOS expression; ↓ plasma NO; ↓liver damage [90]
LPS E. coli Rat (7-day-old pups) Kidney SkQR1
100 nmol/kg		 i.p., 3 h before LPS ↓Acute kidney injury; preservation of cell proliferative activity [99]
E. coli lysate Rat Kidney SkQR1
500 nmol/kg in total		 i.p., 1, 12, 24, 36, and 48 h after intraurethral bacteria injection ↓ Renal cell dearth and animal mortality, restoration of Bcl-2 level in kidney; ↓TNF-α in kidney [88]
LPS E. coli + PepG S. aureus Rat Liver, kidney, lungs, heart, gut 7.5 μmol/kg MitoQ,
then 5 μmol/kg/h MitoQ		 As a bolus
i.v. infusion for 6 h after LPS+PepG		 ↓ Acute liver and renal dysfunction; ↑ MMP in most organs [91]
LPS E. coli + PepG S. aureus Rat Liver, kidney 1.5 μmol/kg MitoQ or MitoVitE or melatonin,
then 1 μmol/kg/h MitoQ, MitoVitE, or melatonin		 As a bolus
i.v. infusion for 5 h after LPS+PepG		 ↓ Mitochondrial damage; ↓ organ dysfunction; ↓ inflammatory response [100]
CLP Mouse Kidney MitoTEMPO
10 mg/kg		 i.p., 6 h after operation ↓ Mitochondrial ROS, protection of complex I and II/III respiration; ↑ SOD; ↓ renal dysfunction (improved renal microcirculation and GFR); ↑ survival of animals [53]
CLP Mouse Diaphragm MitoTEMPOL
10 mg/kg/d		 i.p., immediately after operation and 24 h later or only 6 h after operation ↓ Diaphragm weakness; ↓ mitochondrial superoxide generation; prevention of mitochondrial dysfunction; ↓ proteolytic enzyme activities; ↓ depletion of myosin heavy-chain protein content [43]
LPS E. coli Mouse Heart Melatonin
20 mg/kg/d		 i.p., 48 h before LPS ↓ Cardiomyopathy; ↓ caspase 3 activation and cardiomyocyte apoptosis [47]
LPS E. coli Mouse Heart Melatonin
20 mg/kg/d		 i.p., for 7 days before LPS ↓ Myocardial dysfunction and inflammation; ↓ cardiomyocyte apoptosis; ↑ AMPK activity and autophagy [93]
LPS E. coli Mouse
(pregnant)		 Placenta Melatonin
5.0 mg/kg		 i.p., 30 min before and 150 min after LPS ↓ Placental oxidative stress, hypoxic stress, and ER stress [101]
CLP Mouse Diaphragm Melatonin
30 mg/kg		 i.p., four doses: 30 min before operation, just after operation, and 4 and 8 h after operation ↓ Respiratory chain failure; restoration of the redox status [102]
CLP Rat Liver, kidney, lung, heart, diaphragm Melatonin
10 mg/kg		 i.p. 30 min before and 6 h after operation ↑ Level of GSH; ↓ MDA; ↓ tissue oxidative damage [51]
LPS E. coli Mouse Heart SS-31
5 mg/kg		 i.p., 30 min after LPS ↓ ROS; restoration of myocardial damage; ↑ ATP; ↓ mRNA level of IL-6, IL-1β, and TNF-α; ↓ apoptosis;↑ SOD and GPx [50]
LPS E. coli Mouse Liver, serum XJB-5-131
2 μmol/kg		 i.v. 1 h before LPS ↓ Hepatic iNOS expression, ↓ blood nitrite level [96]
LPS E. coli Mouse Hippocampus SS-31
5 mg/kg		 LPS microinjection in the hippocampi
SS-31 i.p. 30 min before LPS and then once daily for 3 days thereafter		 ↓ ROS, MDA, IL-6, and TNF-α; ↑ SOD; ↓ hippocampal cell apoptosis; ↑ BDNF expression and synaptic protein levels, maintenance of hippocampal neuron morphology; ↓ memory impairment [103]
CLP Mouse Lung, kidney, liver SS-31
5 mg/kg		 i.p, immediately and 5 h after operation ↓ ROS, MDA, TNF-α, MPO activity, iNOS, and NF-κB p65; ↑ ATP; ↓ apoptosis, ↓ the histological damage; ↓ organ dysfunction, no result on mouse survival rate [104]
CLP Mouse Hippocampus SS-31
5 mg/kg		 i.p, immediately after operation and once daily for 6 days thereafter ↓ ROS; ↓ NLRP3 and IL-1β; ↑ ATP; ↓ mitochondrial dysfunction; ↓ apoptosis; ↓ behavior and cognitive deficits; ↓ mortality rate [105]
Live E. coli bacteria Rat Serum M40401 (SOD mimetic)
0.25, 2.5, 25 μmol/kg/h		 i.v. infusion 0.5 and 3 h after bacterial challenge Maintenance of a normal mean arterial pressure; ↓ TNF-α and IL-1β; ↓ mortality [106]

The application of different TPP-based mito-AOX (MitoQ, MitoVitE, MitoTEMPO, or MitoTEMPOL) to primary cultured cells or to cell lines exposed to LPS convincingly evidences their antioxidant and mitochondria-protective properties. As shown in Table 1, TPP-based compounds commonly demonstrate a decrease in mitochondrial/cellular ROS generation, the enhancement of the content of GSH and antioxidant enzymes such as SOD and GPx, and decreased accumulation of lipid peroxidation products such as MDA, as well as restoration of mitochondrial function. These antioxidants decrease the production of proinflammatory cytokines such as IL-1β and IL-18 and prevent NF-kB and caspase activation, leading to the inhibition of apoptosis and the increase in cell survival. MitoTEMPO or MitoQ application highlights the critical role of mtROS in LPS/E. coli-induced inflammasome activation, as shown in colonic epithelial cells [97] and renal proximal tubular cells [89].

Examples of the beneficial application of TPP-based antioxidants in different murine and rat acute inflammation models are summarized in Table 2. TPP-based antioxidants have been shown to accumulate in all major animal organs, such as the heart, kidney, liver, lung, and others, after oral, i.v., or i.p. administration [77][107].

The heart and the cardiovascular system suffer seriously during sepsis. MitoQ administration largely prevents LPS-induced cardiac mitochondrial dysfunction and reduction in cardiac pressure-generating capacity, inhibiting caspase 9 and 3 activity [98]. The septic response is well known to be related to widespread vascular endothelial injury, which plays a key role in the progression of multiple-organ failure [108]. Results obtained on human endothelial cells (HUVECs) exposed to LPS+PepG showed that MitoQ decreases cellular ROS generation, restores the MMP, and attenuates pro-inflammatory mediator production [91] (Table 1). A protective effect of mito-AOX has been demonstrated in an animal model of acute kidney injury caused by CLP following MitoTEMPO i.p. injection six hours after operation [53] or by LPS administration following i.p. injection of SKQR1 (plastoquinol conjugated with decylrhodamine) three hours before LPS administration [99]. In both protocols, despite their differences, mito-AOX were nephroprotective (Table 2). SKQR1 was also highly protective against acute pyelonephritis induced by intraurethral infection [88]. In the frog urinary bladder epithelium, which possesses the characteristics of the mammalian kidney collecting duct, MitoQ effectively inhibited LPS-induced ROS generation, the decline in fatty acid oxidation, and subsequent accumulation of lipid droplets, demonstrating a key role of mtROS in the shift of intracellular lipid metabolism under the influence of bacterial stimuli [56].

The impairment of gut permeability is a serious consequence of dysbiosis. MitoQ has been shown to improve intestinal permeability and inhibit LPS-induced bacterial translocation via a decrease in oxidative stress and restoration of the level of tight junction proteins (occludin and ZO-1) in the gut epithelium [20]. The authors showed that MitoQ alleviates LPS-induced oxidative stress in intestinal epithelial cells, triggering the nuclear translocation of the nuclear factor Nrf2, which, in turn, stimulates the expression of its downstream antioxidant genes [20].

Data presented in Table 2 indicate that the protective effect of mito-AOX can be observed independently on the differences in the administration protocol (application of mito-AOX before, immediately after LPS administration /CLP or some time later). Even a six-hour delay in therapy with a single dose of MitoTEMPO significantly increased mitochondrial respiration and improved renal function and survival of animals [53]. Both immediate and delayed administration of the dismutase mimetic MitoTEMPOL was found to prevent sepsis-induced diaphragm weakness in a similar mode [40]. These observations are very important due to their clinical relevance.

However, some studies have reported that TPP-conjugated compounds fail to inhibit mtROS-mediated injuries [109] or even have a detrimental effect on mitochondrial function. For example, in cultured mesangial cells, MitoQ, MitoTEMPOL, and MitoVitE at a dose of 1 µM inhibited oxidative phosphorylation [110]. Application of MitoQ (500 nM) to proximal tubule cells led to mitochondrial swelling and depolarization [111]. Both MitoQ (500 nM) and MitoTEMPOL (10 µM) had a marked negative effect on the respiration of myoblasts compared to controls [112]. The studies mentioned above revealed that the negative effect of TPP-conjugated compounds on mitochondrial function is related to the toxicity of the carbon alkyl chain of the cation moiety itself [110][111][112]. Another reason for TPP-conjugated mito-AOX toxicity is their ability to be pro-oxidants that generate superoxide via redox cycling [84][113]. A high concentration of antioxidants as well as other factors (the redox potential of matrix environments, the presence of Cu, Fe, and Zn ions) could reverse their behavior from anti- to pro-oxidant, subsequently causing toxic effects [87][114]. The pro-oxidant effect of MitoQ and other related compounds applied at high concentrations (more than 1 µM) has been shown to kill tumor cells, considering mito-AOX as potential chemotherapeutic drugs [87][115][116]. However, no pro-oxidant effect of MitoQ and other targeted quinones was demonstrated in mice who were fed antioxidants [117].

Since the probability of an adverse side effect of cation-conjugated mito-AOX provided by either a cation moiety or pro-oxidative behavior depends critically on their concentration; when dealing with this type of mito-AOX, it is particularly important to choose the relevant concentration, which, in turn, depends on a given cell type. For example, our experiments revealed that frog urinary bladder epithelial cells, demonstrating high tolerance to LPS, are very sensitive to the toxic effect of MitoQ (IC50 = 400 nM) [56]. At doses higher than 25 nM, it reduced the oxygen consumption rate and cell viability, whereas the antioxidant potency of MitoQ and the ability to restore the LPS-induced decline in fatty acids oxidation were observed at a dose of 25 nM [56], which is much less than that in most other in vitro works [65][118][119][120]. Of note, the concentrations of mito-AOX used in in vitro experiments were much higher than those that can be achieved pharmacologically and were associated with protective effects in vivo [121].

4.2. Other Mitochondria-Targeted Conjugates

There was an attempt to design mito-AOX using the mitochondrial protein import machinery, which delivers nuclear-encoded mitochondrial proteins inside the mitochondria via translocase through the outer and inner membranes (TOM and TIM complexes, respectively). A mitochondria-targeted macrocyclic SOD mimetic was synthesized by attaching the mitochondria-targeting sequence peptide to the porphyrin ring of the manganese porphyrin complex MnMPy4P. The resulting construct MnMPy3P–MTS reportedly demonstrated a decrease in LPS-induced cell death in activated macrophages [94].

Another example of the successful application of mito-AOX against LPS is the hemigramicidin–TEMPO conjugate XJB-5-131, which consists of a stable nitroxide radical and a portion of the membrane-active cyclopeptide antibiotic gramicidin S. The gramicidin segment was used to target the nitroxide payload to mitochondria because antibiotics of this type have a high affinity for bacterial membranes [96]. XJB-5-131 limited the LPS-induced inflammatory response both in vitro in macrophages and in vivo in a mice septic model [96].

4.3. Melatonin

Melatonin is a natural antioxidant produced mainly by the pineal gland as well as by most of the organs and tissues. Frequent use of melatonin for treatment of insomnia is based on its traditionally accepted role as a hormonal regulator of the circadian rhythm. Besides this, melatonin possesses antiapoptotic, anti-inflammatory, and antitumor activity, as well as powerful antioxidant properties. These facts alongside its profoundly safe side-effect profile make it possible to propose melatonin as a promising adjunctive drug for different pathological states, including inflammation and sepsis (for review, see [122][123][124][125][126]).

Melatonin was first reported as a potent, broad-spectrum antioxidant and free-radical scavenger in the early 1990s [127]. The electron-rich melatonin molecule provides its antioxidant power via a cascade of scavenging reactions. Unlike classical antioxidants that have the potential to act as anti- and pro-oxidants via redox cycling [128], melatonin forms several stable end products excreted in the urine, which is believed to exclude its pro-oxidant effect [129]. Although the high lipid solubility of melatonin favors its entering all cells and subcellular compartments, melatonin is specifically targeted to mitochondria, where it enters via the oligopeptide transporters PEPT1 and PEPT2 [130]. In addition, melatonin is produced within mitochondria, and its generation can be inducible [131][132]. For these reasons, mitochondria have the highest level of melatonin.

Melatonin is one of the most important endogenous factors in limiting oxidative stress. It provides antioxidant defense via a plethora of mechanisms. Melatonin by itself and also its endogenous metabolites directly scavenge free radicals, bind heavy metals associated with radical production, reduce the membrane potential, and stimulate ETC complex activity and ATP synthesis [133][134][135]. Moreover, melatonin potentiates the activity of a wide variety of antioxidant enzymes. It inhibits the ubiquitination of Nrf2, allowing its binding with the antioxidant response element, which, in turn, activates the transcription of antioxidant genes [136][137]. Melatonin augments the SIRT3 signaling pathway, which protects mitochondria from oxidative damage, upregulates the synthesis of GSH, and acts synergistically with vitamin C, vitamin E, and GSH to scavenge free radicals [128][138].

Numerous experimental studies have revealed the antioxidant and anti-inflammatory properties of melatonin, both in vitro and in vivo. Typical examples are presented in Table 1 and Table 2. On different cells challenged with LPS (HUVECs, cardiomyocytes, alveolar epithelial cells), it was shown that melatonin decreases ROS generation [47][49] and production of proinflammatory cytokines [48][49][93] and increases cellular antioxidant content (SOD, GSH) [47][48][92] through upregulation of Nrf2 expression [49]. Interestingly, not only melatonin but also its structurally related indolamine compounds (6-hydroxymelatonin, tryptamine or indole-3-carboxylic acid) possess antioxidant properties [48].

The beneficial application of melatonin was demonstrated in two animal models of sepsis—administration of LPS and CLP. Melatonin, being commonly injected i.p. before or after sepsis initiation, significantly improved sepsis-induced organ dysfunction (heart, kidney, liver, lung, placenta) by decreasing oxidative tissue damage and the inflammatory response, preserving mitochondrial function [47][51][93][100][101][139]. In the latest works on the septic cardiomyopathy model, it was shown that LPS suppresses the expression of B cell receptor-associated protein 31 (BAP31), a key regulator of endoplasmic reticulum stress, and melatonin could restore BAP31 expression. The knockdown of BAP31 attenuated the beneficial effects of melatonin on mitochondrial function and endoplasmic reticulum homeostasis under LPS [47], suggesting that, at least in part, melatonin contributes to the preservation of cardiac function in septic cardiomyopathy via regulation of BAP31 expression and stability. Another work demonstrated that autophagy plays a critical role in melatonin-induced myocardial protection. Thus, melatonin protects against LPS-induced septic myocardial injury by activating the AMPK-mediated autophagy pathway and further inhibiting mitochondrial injury and myocardial apoptosis [93].

4.4. Cell-Permeable Peptide Antioxidants

In the middle of the 2000s, a family of cell-permeable small synthetic tetrapeptides (Szeto–Schiller peptides (SS peptides)) was introduced as mitochondria-targeted antioxidants. The electron-scavenging abilities of SS peptides were provided by aromatic–cationic motifs in their molecules [140][141][142]. SS peptides readily penetrate the cell via diffusion, selectively accumulate within mitochondria, and concentrate in the IMM without reaching the mitochondrial matrix. In contrast to the MMP-driven entry of triphenylphosphonium-based conjugates into the mitochondria, the accumulation of SS peptides is independent of the MMP and does not depolarize the mitochondrial membrane. For this reason, SS peptides can penetrate not only normal mitochondria but also damaged ones with a low MMP [140].

The most studied peptide of this family is SS-31 (elamipretide, Bendavia™, MTP-131, d-Arg-Dmt-Lys-Phe-NH2), which, in addition to its mtROS-scavenging ability, links selectively to cardiolipin by electrostatic and hydrophobic interactions [143][144]. Thus, SS-31 is now positioned more as a cardiolipin stabilizer/protector than as a mtROS scavenger.

Cardiolipin is readily oxidized by mtROS, which leads to multiple injuries. Oxidized cardiolipin disrupts the structure of respiratory supercomplexes to inhibit electron transfer and oxidative phosphorylation [34]. Translocation of oxidized cardiolipin from the IMM into the OMM provides a docking station for NLRP3 inflammasome assembly, and it can trigger mitochondrial fission and initiate mitophagy [145]. Binding of SS-31 to cardiolipin inhibits cardiolipin peroxidation, stabilizes cristae curvatures [143][144][146][147], and restores the stability and activity of respiratory complexes [148].

The linking of SS-31 to cardiolipin also inhibits the peroxidase activity of cytochrome C to result in decreasing mtROS production and improving the coupling between oxygen consumption and ATP synthesis [144]. SS-31 enhances ATP levels even under conditions of low substrate and oxygen supply, such as ischemia [143][146], or in increased energy demand states, such as sepsis and others pathologies [50][149][150]. The restoration of mitochondrial functioning by SS-31 can prevent a wide range of downstream cellular events, e.g., inflammasome activation and cytokine expression, autophagy, apoptosis, and necrosis. The beneficial effects of SS-31 were reported in different disease models (for review, see [151]), demonstrating the existence of a common mechanism mediating its action in different pathological conditions.

The protective effect of SS-31 against LPS was demonstrated in several in vitro and in vivo models (see Table 1 and Table 2). In LPS-treated cells and CPL/LPS-challenged mice, SS-31 decreased apoptosis, improved sepsis-induced organ dysfunction, restored morphological damage, and reversed mitochondrial dysfunction [50][103][104][105]. It also attenuated ROS and MDA levels [50][103][104][105], maintained ATP production [50][104][105], and suppressed pro-inflammatory cytokine expression [50][103][104][105].

Several successive clinical trials in phases 1-3 were conducted in patients with cardiac, renal, skeletal muscle, and ophthalmic problems, as well as in mitochondrial myopathy patients (for review, see [151]). No adverse side effects of SS-31 were found until now. The safety of using SS-31, a drug with multiple beneficial pharmacological properties, for organs most affected by sepsis is particularly important. Very promising preclinical and clinical trial findings encourage to develop SS-31-based therapeutic approaches for the treatment of sepsis and other pathologies.

4.5. Suppressors of Site IQ and IIIQ Electron Leakage

Recently, small molecules from different chemical families that specifically suppress mitochondrial superoxide/H2O2 production (S1QELs for site IQ [152] and S3QELs for site IIIQo [153]) were identified by chemical screening. They bind directly to complex I or III and selectively suppress electron leakage without inhibiting oxidative phosphorylation [152][153], as well as inhibit the reverse electron flow through complex I [154]. They do not cause cytotoxicity at their effective concentrations [153] and do not participate in redox recycling [155].

The cytoprotective effect of S1QELs against oxidative damage has been demonstrated in animal (rat, mouse), human, and different cellular models [153][156][157]. S1QELs protected against ischemia-reperfusion injury in a perfused mouse heart [158]. In a murine model of asystolic cardiac arrest, S1QELs diminished myocardial ROS, as well as improved myocardial function after cardiopulmonary resuscitation, neurologic outcomes, and survival [159]. In recent papers, S1QELs and S3QELs have been offered as promising investigation tools for elucidating the functioning of IQ and IIIQo sites in normal and pathological conditions, opening up new possibilities for better therapy [155][160]. Given the fact that LPS-driven mtROS are generated predominantly by mitochondrial complex I, S1QELs can potentially be specific suppressors of LPS-induced mtROS production, gently withstanding LPS-induced oxidative stress. However, the efficiency of S1QELs and S3QELs in a sepsis animal model or LPS-induced injury remains poorly investigated and warrants further research.

This entry is adapted from the peer-reviewed paper 10.3390/pharmaceutics13020144

References

  1. Miller, S.I.; Ernst, R.K.; Bader, M.W. LPS, TLR4 and infectious disease diversity. Nat. Rev. Microbiol. 2005, 3, 36–46.
  2. Nikolaeva, S.; Bachteeva, V.; Fock, E.; Herterich, S.; Lavrova, E.; Borodkina, A.; Gambaryan, S.; Parnova, R. Frog urinary bladder epithelial cells express TLR4 and respond to bacterial LPS by increase of iNOS expression and L-arginine uptake. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2012, 303, R1042–R1052.
  3. Lien, E.; Ingalls, R.R. Toll-like receptors. Crit. Care Med. 2002, 30, S1–S11.
  4. Lu, Y.C.; Yeh, W.C.; Ohashi, P.S. LPS/TLR4 signal transduction pathway. Cytokine 2008, 42, 145–151.
  5. Takeda, K.; Akira, S. TLR signaling pathways. Semin. Immunol. 2004, 16, 3–9.
  6. Mazgaeen, L.; Gurung, P. Recent Advances in Lipopolysaccharide Recognition Systems. Int. J. Mol. Sci 2020, 21, 379.
  7. Tschopp, J. Mitochondria: Sovereign of inflammation? Eur. J. Immunol. 2011, 41, 1196–1202.
  8. Evavold, C.L.; Kagan, J.C. How Inflammasomes Inform Adaptive Immunity. J. Mol. Biol. 2018, 430, 217–237.
  9. Dagenais, M.; Skeldon, A.; Saleh, M. The inflammasome: In memory of Dr. Jurg Tschopp. Cell Death Differ. 2012, 19, 5–12.
  10. Demirel, I.; Persson, A.; Brauner, A.; Sarndahl, E.; Kruse, R.; Persson, K. Activation of the NLRP3 Inflammasome Pathway by Uropathogenic Escherichia coli Is Virulence Factor-Dependent and Influences Colonization of Bladder Epithelial Cells. Front. Cell Infect. Microbiol. 2018, 8, 81.
  11. Swanson, K.V.; Deng, M.; Ting, J.P. The NLRP3 inflammasome: Molecular activation and regulation to therapeutics. Nat. Rev. Immunol. 2019, 19, 477–489.
  12. Liu, X.; Zhang, Z.; Ruan, J.; Pan, Y.; Magupalli, V.G.; Wu, H.; Lieberman, J. Inflammasome-activated gasdermin D causes pyroptosis by forming membrane pores. Nature 2016, 535, 153–158.
  13. Murphy, M.P. How mitochondria produce reactive oxygen species. Biochem. J. 2009, 417, 1–13.
  14. Skulachev, V.P. Cationic antioxidants as a powerful tool against mitochondrial oxidative stress. Biochem. Biophys. Res. Commun. 2013, 441, 275–279.
  15. Sies, H.; Berndt, C.; Jones, D.P. Oxidative Stress. Annu. Rev. Biochem. 2017, 86, 715–748.
  16. Andreyev, A.Y.; Kushnareva, Y.E.; Murphy, A.N.; Starkov, A.A. Mitochondrial ROS Metabolism: 10 Years Later. Biochemistry (Moscow) 2015, 80, 517–531.
  17. Robb, E.L.; Hall, A.R.; Prime, T.A.; Eaton, S.; Szibor, M.; Viscomi, C.; James, A.M.; Murphy, M.P. Control of mitochondrial superoxide production by reverse electron transport at complex I. J. Biol. Chem. 2018, 293, 9869–9879.
  18. Treberg, J.R.; Quinlan, C.L.; Brand, M.D. Evidence for two sites of superoxide production by mitochondrial NADH-ubiquinone oxidoreductase (complex I). J. Biol. Chem. 2011, 286, 27103–27110.
  19. Hirst, J.; King, M.S.; Pryde, K.R. The production of reactive oxygen species by complex I. Biochem. Soc. Trans. 2008, 36, 976–980.
  20. Zhang, S.; Zhou, Q.; Li, Y.; Zhang, Y.; Wu, Y. MitoQ Modulates Lipopolysaccharide-Induced Intestinal Barrier Dysfunction via Regulating Nrf2 Signaling. Mediat. Inflamm. 2020, 2020, 3276148.
  21. Bellezza, I.; Giambanco, I.; Minelli, A.; Donato, R. Nrf2-Keap1 signaling in oxidative and reductive stress. Biochim. Biophys. Acta Mol. Cell Res. 2018, 1865, 721–733.
  22. Zorov, D.B.; Bannikova, S.Y.; Belousov, V.V.; Vyssokikh, M.Y.; Zorova, L.D.; Isaev, N.K.; Krasnikov, B.F.; Plotnikov, E.Y. Reactive oxygen and nitrogen species: Friends or foes? Biochemistry (Moscow) 2005, 70, 215–221.
  23. Droge, W. Free radicals in the physiological control of cell function. Physiol. Rev. 2002, 82, 47–95.
  24. Kozlov, A.V.; Bahrami, S.; Calzia, E.; Dungel, P.; Gille, L.; Kuznetsov, A.V.; Troppmair, J. Mitochondrial dysfunction and biogenesis: Do ICU patients die from mitochondrial failure? Ann. Intensive Care 2011, 1, 41.
  25. Zorov, D.B.; Juhaszova, M.; Sollott, S.J. Mitochondrial reactive oxygen species (ROS) and ROS-induced ROS release. Physiol. Rev. 2014, 94, 909–950.
  26. Prauchner, C.A. Oxidative stress in sepsis: Pathophysiological implications justifying antioxidant co-therapy. Burns 2017, 43, 471–485.
  27. Di Meo, S.; Reed, T.; Venditti, P.; Victor, V. Role of ROS and RNS sources in physiological and pathological conditions. Oxidative Med. Cell Longev. 2016, 2016, 1245049.
  28. Yang, Y.; Karakhanova, S.; Hartwig, W.; D’Haese, J.G.; Philippov, P.P.; Werner, J.; Bazhin, A.V. Mitochondria and Mitochondrial ROS in Cancer: Novel Targets for Anticancer Therapy. J. Cell. Physiol. 2016, 231, 2570–2581.
  29. Robinson, A.R.; Yousefzadeh, M.J.; Rozgaja, T.A.; Wang, J.; Li, X.; Tilstra, J.S.; Feldman, C.H.; Gregg, S.Q.; Johnson, C.H.; Skoda, E.M.; et al. Spontaneous DNA damage to the nuclear genome promotes senescence, redox imbalance and aging. Redox Biol. 2018, 17, 259–273.
  30. Zhang, D.; Li, Y.; Heims-Waldron, D.; Bezzerides, V.; Guatimosim, S.; Guo, Y.; Gu, F.; Zhou, P.; Lin, Z.; Ma, Q.; et al. Mitochondrial Cardiomyopathy Caused by Elevated Reactive Oxygen Species and Impaired Cardiomyocyte Proliferation. Circ. Res. 2018, 122, 74–87.
  31. De, I.; Dogra, N.; Singh, S. The Mitochondrial Unfolded Protein Response: Role in Cellular Homeostasis and Disease. Curr. Mol. Med. 2017, 17, 587–597.
  32. Yao, X.; Carlson, D.; Sun, Y.; Ma, L.; Wolf, S.E.; Minei, J.P.; Zang, Q.S. Mitochondrial ROS Induces Cardiac Inflammation via a Pathway through mtDNA Damage in a Pneumonia-Related Sepsis Model. PLoS ONE 2015, 10, e0139416.
  33. Callahan, L.A.; Supinski, G.S. Sepsis induces diaphragm electron transport chain dysfunction and protein depletion. Am. J. Respir. Crit. Care Med. 2005, 172, 861–868.
  34. Paradies, G.; Petrosillo, G.; Paradies, V.; Ruggiero, F.M. Role of cardiolipin peroxidation and Ca2+ in mitochondrial dysfunction and disease. Cell Calcium. 2009, 45, 643–650.
  35. Orrenius, S.; Gogvadze, V.; Zhivotovsky, B. Calcium and mitochondria in the regulation of cell death. Biochem. Biophys. Res. Commun. 2015, 460, 72–81.
  36. Williamson, J.; Davison, G. Targeted Antioxidants in Exercise-Induced Mitochondrial Oxidative Stress: Emphasis on DNA Damage. Antioxidants (Basel) 2020, 9, 142.
  37. Galley, H.F. Oxidative stress and mitochondrial dysfunction in sepsis. Br. J. Anaesth. 2011, 107, 57–64.
  38. Poderoso, J.J. The formation of peroxynitrite in the applied physiology of mitochondrial nitric oxide. Arch. Biochem. Biophys. 2009, 484, 214–220.
  39. Rocha, M.; Herance, R.; Rovira, S.; Hernandez-Mijares, A.; Victor, V.M. Mitochondrial dysfunction and antioxidant therapy in sepsis. Infect. Disord. Drug Targets 2012, 12, 161–178.
  40. Supinski, G.S.; Schroder, E.A.; Callahan, L.A. Mitochondria and Critical Illness. Chest 2020, 157, 310–322.
  41. Voloboueva, L.A.; Emery, J.F.; Sun, X.; Giffard, R.G. Inflammatory response of microglial BV-2 cells includes a glycolytic shift and is modulated by mitochondrial glucose-regulated protein 75/mortalin. FEBS Lett. 2013, 587, 756–762.
  42. Park, J.; Min, J.S.; Kim, B.; Chae, U.B.; Yun, J.W.; Choi, M.S.; Kong, I.K.; Chang, K.T.; Lee, D.S. Mitochondrial ROS govern the LPS-induced pro-inflammatory response in microglia cells by regulating MAPK and NF-kappaB pathways. Neurosci. Lett. 2015, 584, 191–196.
  43. Supinski, G.S.; Wang, L.; Schroder, E.A.; Callahan, L.A.P. MitoTEMPOL, a mitochondrial targeted antioxidant, prevents sepsis-induced diaphragm dysfunction. Am. J. Physiol. Lung Cell Mol. Physiol. 2020, 319, L228–L238.
  44. Li, X.; Wang, X.; Zheng, M.; Luan, Q.X. Mitochondrial reactive oxygen species mediate the lipopolysaccharide-induced pro-inflammatory response in human gingival fibroblasts. Exp. Cell Res. 2016, 347, 212–221.
  45. Jiao, P.; Li, W.; Shen, L.; Li, Y.; Yu, L.; Liu, Z. The protective effect of doxofylline against lipopolysaccharides (LPS)-induced activation of NLRP3 inflammasome is mediated by SIRT1 in human pulmonary bronchial epithelial cells. Artif. Cells Nanomed. Biotechnol. 2020, 48, 687–694.
  46. Mills, E.L.; Kelly, B.; Logan, A.; Costa, A.S.; Varma, M.; Bryant, C.E.; Tourlomousis, P.; Däbritz, J.H.M.; Gottlieb, E.; Latorre, I. Succinate dehydrogenase supports metabolic repurposing of mitochondria to drive inflammatory macrophages. Cell 2016, 167, 457–470.e413.
  47. Zhang, J.; Wang, L.; Xie, W.; Hu, S.; Zhou, H.; Zhu, P.; Zhu, H. Melatonin attenuates ER stress and mitochondrial damage in septic cardiomyopathy: A new mechanism involving BAP31 upregulation and MAPK-ERK pathway. J. Cell. Physiol. 2020, 235, 2847–2856.
  48. Lowes, D.A.; Almawash, A.M.; Webster, N.R.; Reid, V.L.; Galley, H.F. Melatonin and structurally similar compounds have differing effects on inflammation and mitochondrial function in endothelial cells under conditions mimicking sepsis. Br. J. Anaesth. 2011, 107, 193–201.
  49. Ding, Z.; Wu, X.; Wang, Y.; Ji, S.; Zhang, W.; Kang, J.; Li, J.; Fei, G. Melatonin prevents LPS-induced epithelial-mesenchymal transition in human alveolar epithelial cells via the GSK-3beta/Nrf2 pathway. Biomed. Pharmacother. 2020, 132, 110827.
  50. Liu, Y.; Yang, W.; Sun, X.; Xie, L.; Yang, Y.; Sang, M.; Jiao, R. SS31 Ameliorates Sepsis-Induced Heart Injury by Inhibiting Oxidative Stress and Inflammation. Inflammation 2019, 42, 2170–2180.
  51. Sener, G.; Toklu, H.; Kapucu, C.; Ercan, F.; Erkanli, G.; Kacmaz, A.; Tilki, M.; Yegen, B.C. Melatonin protects against oxidative organ injury in a rat model of sepsis. Surg. Today 2005, 35, 52–59.
  52. Vanasco, V.; Magnani, N.D.; Cimolai, M.C.; Valdez, L.B.; Evelson, P.; Boveris, A.; Alvarez, S. Endotoxemia impairs heart mitochondrial function by decreasing electron transfer, ATP synthesis and ATP content without affecting membrane potential. J. Bioenerg. Biomembr. 2012, 44, 243–252.
  53. Patil, N.K.; Parajuli, N.; MacMillan-Crow, L.A.; Mayeux, P.R. Inactivation of renal mitochondrial respiratory complexes and manganese superoxide dismutase during sepsis: Mitochondria-targeted antioxidant mitigates injury. Am. J. Physiol. Renal. Physiol. 2014, 306, F734–F743.
  54. Chandel, N.S.; McClintock, D.S.; Feliciano, C.E.; Wood, T.M.; Melendez, J.A.; Rodriguez, A.M.; Schumacker, P.T. Reactive oxygen species generated at mitochondrial Complex III stabilize HIF-1-alpha during hypoxia: A mechanism of O2 sensing. J. Biol. Chem. 2000, 275, 25130–25138.
  55. Quoilin, C.; Mouithys-Mickalad, A.; Duranteau, J.; Gallez, B.; Hoebeke, M. Endotoxin-induced basal respiration alterations of renal HK-2 cells: A sign of pathologic metabolism down-regulation. Biochem. Biophys. Res. Commun. 2012, 423, 350–354.
  56. Fock, E.; Bachteeva, V.; Lavrova, E.; Parnova, R. Mitochondrial-Targeted Antioxidant MitoQ Prevents E. coli Lipopolysaccharide-Induced Accumulation of Triacylglycerol and Lipid Droplets Biogenesis in Epithelial Cells. J. Lipids 2018, 2018, 5745790.
  57. Jeger, V.; Brandt, S.; Porta, F.; Jakob, S.M.; Takala, J.; Djafarzadeh, S. Dose response of endotoxin on hepatocyte and muscle mitochondrial respiration in vitro. Biomed. Res. Int. 2015, 2015, 353074.
  58. Karlsson, M.; Hara, N.; Morata, S.; Sjovall, F.; Kilbaugh, T.; Hansson, M.J.; Uchino, H.; Elmer, E. Diverse and Tissue-Specific Mitochondrial Respiratory Response in a Mouse Model of Sepsis-Induced Multiple Organ Failure. Shock 2016, 45, 404–410.
  59. Frisard, M.I.; Wu, Y.; McMillan, R.P.; Voelker, K.A.; Wahlberg, K.A.; Anderson, A.S.; Boutagy, N.; Resendes, K.; Ravussin, E.; Hulver, M.W. Low levels of lipopolysaccharide modulate mitochondrial oxygen consumption in skeletal muscle. Metabolism 2015, 64, 416–427.
  60. Apostolova, N.; Garcia-Bou, R.; Hernandez-Mijares, A.; Herance, R.; Rocha, M.; Victor, V.M. Mitochondrial antioxidants alleviate oxidative and nitrosative stress in a cellular model of sepsis. Pharm. Res. 2011, 28, 2910–2919.
  61. Qiu, Z.; He, Y.; Ming, H.; Lei, S.; Leng, Y.; Xia, Z.Y. Lipopolysaccharide (LPS) Aggravates High Glucose- and Hypoxia/Reoxygenation-Induced Injury through Activating ROS-Dependent NLRP3 Inflammasome-Mediated Pyroptosis in H9C2 Cardiomyocytes. J. Diabetes Res. 2019, 2019, 8151836.
  62. Zhang, H.; Zhang, W.; Jiao, F.; Li, X.; Zhang, H.; Wang, L.; Gong, Z. The nephroprotective effect of MS-275 on lipopolysaccharide (LPS)-induced acute kidney injury by inhibiting reactive oxygen species (ROS)-oxidative stress and endoplasmic reticulum stress. Med. Sci. Mon. Int. Med. J. Exp. Clin. Res. 2018, 24, 2620.
  63. Raulien, N.; Friedrich, K.; Strobel, S.; Rubner, S.; Baumann, S.; von Bergen, M.; Korner, A.; Krueger, M.; Rossol, M.; Wagner, U. Fatty Acid Oxidation Compensates for Lipopolysaccharide-Induced Warburg Effect in Glucose-Deprived Monocytes. Front. Immunol. 2017, 8, 609.
  64. Palsson-McDermott, E.M.; Curtis, A.M.; Goel, G.; Lauterbach, M.A.; Sheedy, F.J.; Gleeson, L.E.; van den Bosch, M.W.; Quinn, S.R.; Domingo-Fernandez, R.; Johnston, D.G. Pyruvate kinase M2 regulates Hif-1α activity and IL-1β induction and is a critical determinant of the warburg effect in LPS-activated macrophages. Cell Metab. 2015, 21, 65–80.
  65. Kelly, B.; Tannahill, G.M.; Murphy, M.P.; O’Neill, L.A. Metformin Inhibits the Production of Reactive Oxygen Species from NADH:Ubiquinone Oxidoreductase to Limit Induction of Interleukin-1β (IL-1β) and Boosts Interleukin-10 (IL-10) in Lipopolysaccharide (LPS)-activated Macrophages. J. Biol. Chem. 2015, 290, 20348–20359.
  66. West, A.P.; Brodsky, I.E.; Rahner, C.; Woo, D.K.; Erdjument-Bromage, H.; Tempst, P.; Walsh, M.C.; Choi, Y.; Shadel, G.S.; Ghosh, S. TLR signalling augments macrophage bactericidal activity through mitochondrial ROS. Nature 2011, 472, 476–480.
  67. Kopp, E.; Medzhitov, R.; Carothers, J.; Xiao, C.; Douglas, I.; Janeway, C.A.; Ghosh, S. ECSIT is an evolutionarily conserved intermediate in the Toll/IL-1 signal transduction pathway. Genes Dev. 1999, 13, 2059–2071.
  68. Geng, J.; Sun, X.; Wang, P.; Zhang, S.; Wang, X.; Wu, H.; Hong, L.; Xie, C.; Li, X.; Zhao, H.; et al. Kinases Mst1 and Mst2 positively regulate phagocytic induction of reactive oxygen species and bactericidal activity. Nat. Immunol. 2015, 16, 1142–1152.
  69. Tur, J.; Pereira-Lopes, S.; Vico, T.; Marin, E.A.; Munoz, J.P.; Hernandez-Alvarez, M.; Cardona, P.J.; Zorzano, A.; Lloberas, J.; Celada, A. Mitofusin 2 in Macrophages Links Mitochondrial ROS Production, Cytokine Release, Phagocytosis, Autophagy, and Bactericidal Activity. Cell Rep. 2020, 32, 108079.
  70. Williams, R.J.; Spencer, J.P.; Rice-Evans, C. Flavonoids: Antioxidants or signalling molecules? Free Radic. Biol. Med. 2004, 36, 838–849.
  71. Murphy, M.P. Antioxidants as therapies: Can we improve on nature? Free Radical. Biol. Med. 2014, 66, 20–23.
  72. Haddad, J.J.; Land, S.C. Redox/ROS regulation of lipopolysaccharide-induced mitogen-activated protein kinase (MAPK) activation and MAPK-mediated TNF-alpha biosynthesis. Br. J. Pharmacol. 2002, 135, 520–536.
  73. Runkel, E.D.; Liu, S.; Baumeister, R.; Schulze, E. Surveillance-activated defenses block the ROS-induced mitochondrial unfolded protein response. PLoS Genet. 2013, 9, e1003346.
  74. Martindale, J.L.; Holbrook, N.J. Cellular response to oxidative stress: Signaling for suicide and survival. J. Cell. Physiol. 2002, 192, 1–15.
  75. Chen, L.; Liu, P.; Feng, X.; Ma, C. Salidroside suppressing LPS-induced myocardial injury by inhibiting ROS-mediated PI3K/Akt/mTOR pathway in vitro and in vivo. J. Cell. Mol. Med. 2017, 21, 3178–3189.
  76. Liberman, E.A.; Topaly, V.P. Permeability of bimolecular phospholipid membranes for fat-soluble ions. Biofizika 1969, 14, 452–461.
  77. Kelso, G.F.; Porteous, C.M.; Hughes, G.; Ledgerwood, E.C.; Gane, A.M.; Smith, R.A.; Murphy, M.P. Prevention of mitochondrial oxidative damage using targeted antioxidants. Ann. N. Y. Acad. Sci. 2002, 959, 263–274.
  78. Battogtokh, G.; Choi, Y.S.; Kang, D.S.; Park, S.J.; Shim, M.S.; Huh, K.M.; Cho, Y.Y.; Lee, J.Y.; Lee, H.S.; Kang, H.C. Mitochondria-targeting drug conjugates for cytotoxic, anti-oxidizing and sensing purposes: Current strategies and future perspectives. Acta Pharm. Sin. B 2018, 8, 862–880.
  79. Zielonka, J.; Joseph, J.; Sikora, A.; Hardy, M.; Ouari, O.; Vasquez-Vivar, J.; Cheng, G.; Lopez, M.; Kalyanaraman, B. Mitochondria-Targeted Triphenylphosphonium-Based Compounds: Syntheses, Mechanisms of Action, and Therapeutic and Diagnostic Applications. Chem. Rev. 2017, 117, 10043–10120.
  80. Murphy, M.P.; Smith, R.A. Targeting antioxidants to mitochondria by conjugation to lipophilic cations. Annu. Rev. Pharmacol. Toxicol. 2007, 47, 629–656.
  81. Skulachev, M.V.; Antonenko, Y.N.; Anisimov, V.N.; Chernyak, B.V.; Cherepanov, D.A.; Chistyakov, V.A.; Egorov, M.V.; Kolosova, N.G.; Korshunova, G.A.; Lyamzaev, K.G.; et al. Mitochondrial-targeted plastoquinone derivatives. Effect on senescence and acute age-related pathologies. Curr. Drug Targets 2011, 12, 800–826.
  82. Trnka, J.; Blaikie, F.H.; Smith, R.A.; Murphy, M.P. A mitochondria-targeted nitroxide is reduced to its hydroxylamine by ubiquinol in mitochondria. Free Radic. Biol. Med. 2008, 44, 1406–1419.
  83. Smith, R.A.; Porteous, C.M.; Coulter, C.V.; Murphy, M.P. Selective targeting of an antioxidant to mitochondria. Eur. J. Biochem. 1999, 263, 709–716.
  84. James, A.M.; Cocheme, H.M.; Smith, R.A.; Murphy, M.P. Interactions of mitochondria-targeted and untargeted ubiquinones with the mitochondrial respiratory chain and reactive oxygen species. Implications for the use of exogenous ubiquinones as therapies and experimental tools. J. Biol. Chem. 2005, 280, 21295–21312.
  85. Fetisova, E.K.; Muntyan, M.S.; Lyamzaev, K.G.; Chernyak, B.V. Therapeutic Effect of the Mitochondria-Targeted Antioxidant SkQ1 on the Culture Model of Multiple Sclerosis. Oxidative Med. Cell. Longev. 2019, 2019, 2082561.
  86. Ross, M.F.; Da Ros, T.; Blaikie, F.H.; Prime, T.A.; Porteous, C.M.; Severina, I.I.; Skulachev, V.P.; Kjaergaard, H.G.; Smith, R.A.; Murphy, M.P. Accumulation of lipophilic dications by mitochondria and cells. Biochem. J. 2006, 400, 199–208.
  87. Plotnikov, E.Y.; Zorov, D.B. Pros and Cons of Use of Mitochondria-Targeted Antioxidants. Antioxidants 2019, 8, 316.
  88. Plotnikov, E.Y.; Morosanova, M.A.; Pevzner, I.B.; Zorova, L.D.; Manskikh, V.N.; Pulkova, N.V.; Galkina, S.I.; Skulachev, V.P.; Zorov, D.B. Protective effect of mitochondria-targeted antioxidants in an acute bacterial infection. Proc. Natl. Acad. Sci. USA 2013, 110, E3100–E3108.
  89. Lei, X.; Li, S.; Luo, C.; Wang, Y.; Liu, Y.; Xu, Z.; Huang, Q.; Zou, F.; Chen, Y.; Peng, F.; et al. Micheliolide Attenuates Lipopolysaccharide-Induced Inflammation by Modulating the mROS/NF-kappaB/NLRP3 Axis in Renal Tubular Epithelial Cells. Mediat. Inflamm. 2020, 2020, 3934769.
  90. Weidinger, A.; Mullebner, A.; Paier-Pourani, J.; Banerjee, A.; Miller, I.; Lauterbock, L.; Duvigneau, J.C.; Skulachev, V.P.; Redl, H.; Kozlov, A.V. Vicious inducible nitric oxide synthase-mitochondrial reactive oxygen species cycle accelerates inflammatory response and causes liver injury in rats. Antioxid. Redox Signal. 2015, 22, 572–586.
  91. Lowes, D.A.; Thottakam, B.M.; Webster, N.R.; Murphy, M.P.; Galley, H.F. The mitochondria-targeted antioxidant MitoQ protects against organ damage in a lipopolysaccharide-peptidoglycan model of sepsis. Free Radic. Biol. Med. 2008, 45, 1559–1565.
  92. Minter, B.E.; Lowes, D.A.; Webster, N.R.; Galley, H.F. Differential Effects of MitoVitE, alpha-Tocopherol and Trolox on Oxidative Stress, Mitochondrial Function and Inflammatory Signalling Pathways in Endothelial Cells Cultured under Conditions Mimicking Sepsis. Antioxidants 2020, 9, 195.
  93. Di, S.; Wang, Z.; Hu, W.; Yan, X.; Ma, Z.; Li, X.; Li, W.; Gao, J. The Protective Effects of Melatonin Against LPS-Induced Septic Myocardial Injury: A Potential Role of AMPK-Mediated Autophagy. Front. Endocrinol. (Lausanne) 2020, 11, 162.
  94. Asayama, S.; Kawamura, E.; Nagaoka, S.; Kawakami, H. Design of manganese porphyrin modified with mitochondrial signal peptide for a new antioxidant. Mol. Pharm. 2006, 3, 468–470.
  95. Mo, Y.N.; Deng, S.Y.; Zhang, L.N.; Huang, Y.; Li, W.C.; Peng, Q.Y.; Liu, Z.Y.; Ai, Y.H. SS-31 reduces inflammation and oxidative stress through the inhibition of Fis1 expression in lipopolysaccharide-stimulated microglia. Biochem. Biophys. Res. Commun. 2019, 520, 171–178.
  96. Fink, M.P.; Macias, C.A.; Xiao, J.; Tyurina, Y.Y.; Delude, R.L.; Greenberger, J.S.; Kagan, V.E.; Wipf, P. Hemigramicidin-TEMPO conjugates: Novel mitochondria-targeted antioxidants. Crit. Care Med. 2007, 35, S461–S467.
  97. Xue, Y.; Du, M.; Zhu, M.J. Quercetin suppresses NLRP3 inflammasome activation in epithelial cells triggered by Escherichia coli O157:H7. Free Radic. Biol. Med. 2017, 108, 760–769.
  98. Supinski, G.S.; Murphy, M.P.; Callahan, L.A. MitoQ administration prevents endotoxin-induced cardiac dysfunction. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2009, 297, R1095–R1102.
  99. Plotnikov, E.Y.; Pevzner, I.B.; Zorova, L.D.; Chernikov, V.P.; Prusov, A.N.; Kireev, I.I.; Silachev, D.N.; Skulachev, V.P.; Zorov, D.B. Mitochondrial Damage and Mitochondria-Targeted Antioxidant Protection in LPS-Induced Acute Kidney Injury. Antioxidants 2019, 8, 176.
  100. Lowes, D.; Webster, N.; Murphy, M.; Galley, H. Antioxidants that protect mitochondria reduce interleukin-6 and oxidative stress, improve mitochondrial function, and reduce biochemical markers of organ dysfunction in a rat model of acute sepsis. Br. J. Anaesth. 2013, 110, 472–480.
  101. Wang, H.; Li, L.; Zhao, M.; Chen, Y.H.; Zhang, Z.H.; Zhang, C.; Ji, Y.L.; Meng, X.H.; Xu, D.X. Melatonin alleviates lipopolysaccharide-induced placental cellular stress response in mice. J. Pineal. Res. 2011, 50, 418–426.
  102. Lopez, L.C.; Escames, G.; Tapias, V.; Utrilla, P.; Leon, J.; Acuna-Castroviejo, D. Identification of an inducible nitric oxide synthase in diaphragm mitochondria from septic mice—Its relation with mitochondrial dysfunction and prevention by melatonin. Int. J. Biochem. Cell B 2006, 38, 267–278.
  103. Zhao, W.; Xu, Z.; Cao, J.; Fu, Q.; Wu, Y.; Zhang, X.; Long, Y.; Zhang, X.; Yang, Y.; Li, Y.; et al. Elamipretide (SS-31) improves mitochondrial dysfunction, synaptic and memory impairment induced by lipopolysaccharide in mice. J. Neuroinflamm. 2019, 16, 230.
  104. Li, G.; Wu, J.; Li, R.; Yuan, D.; Fan, Y.; Yang, J.; Ji, M.; Zhu, S. Protective Effects of Antioxidant Peptide SS-31 Against Multiple Organ Dysfunctions During Endotoxemia. Inflammation 2016, 39, 54–64.
  105. Wu, J.; Zhang, M.; Hao, S.; Jia, M.; Ji, M.; Qiu, L.; Sun, X.; Yang, J.; Li, K. Mitochondria-Targeted Peptide Reverses Mitochondrial Dysfunction and Cognitive Deficits in Sepsis-Associated Encephalopathy. Mol. Neurobiol. 2015, 52, 783–791.
  106. Macarthur, H.; Couri, D.M.; Wilken, G.H.; Westfall, T.C.; Lechner, A.J.; Matuschak, G.M.; Chen, Z.; Salvemini, D. Modulation of serum cytokine levels by a novel superoxide dismutase mimetic, M40401, in an Escherichia coli model of septic shock: Correlation with preserved circulating catecholamines. Crit. Care Med. 2003, 31, 237–245.
  107. Smith, R.A.; Porteous, C.M.; Gane, A.M.; Murphy, M.P. Delivery of bioactive molecules to mitochondria in vivo. Proc. Natl. Acad. Sci. USA 2003, 100, 5407–5412.
  108. Huet, O.; Dupic, L.; Harrois, A.; Duranteau, J. Oxidative stress and endothelial dysfunction during sepsis. Front. Biosci. (Landmark Ed.) 2011, 16, 1986–1995.
  109. Rademann, P.; Weidinger, A.; Drechsler, S.; Meszaros, A.; Zipperle, J.; Jafarmadar, M.; Dumitrescu, S.; Hacobian, A.; Ungelenk, L.; Rostel, F.; et al. Mitochondria-Targeted Antioxidants SkQ1 and MitoTEMPO Failed to Exert a Long-Term Beneficial Effect in Murine Polymicrobial Sepsis. Oxid. Med. Cell Longev. 2017, 2017, 6412682.
  110. Reily, C.; Mitchell, T.; Chacko, B.K.; Benavides, G.; Murphy, M.P.; Darley-Usmar, V. Mitochondrially targeted compounds and their impact on cellular bioenergetics. Redox Biol. 2013, 1, 86–93.
  111. Gottwald, E.M.; Duss, M.; Bugarski, M.; Haenni, D.; Schuh, C.D.; Landau, E.M.; Hall, A.M. The targeted anti-oxidant MitoQ causes mitochondrial swelling and depolarization in kidney tissue. Physiol. Rep. 2018, 6, e13667.
  112. Patkova, J.; Andel, M.; Trnka, J. Palmitate-induced cell death and mitochondrial respiratory dysfunction in myoblasts are not prevented by mitochondria-targeted antioxidants. Cell Physiol. Biochem. 2014, 33, 1439–1451.
  113. Doughan, A.K.; Dikalov, S.I. Mitochondrial redox cycling of mitoquinone leads to superoxide production and cellular apoptosis. Antioxid. Redox Signal. 2007, 9, 1825–1836.
  114. James, A.M.; Smith, R.A.; Murphy, M.P. Antioxidant and prooxidant properties of mitochondrial Coenzyme Q. Arch. Biochem. Biophys. 2004, 423, 47–56.
  115. Pokrzywinski, K.L.; Biel, T.G.; Kryndushkin, D.; Rao, V.A. Therapeutic Targeting of the Mitochondria Initiates Excessive Superoxide Production and Mitochondrial Depolarization Causing Decreased mtDNA Integrity. PLoS ONE 2016, 11, e0168283.
  116. Cheng, G.; Zielonka, J.; McAllister, D.M.; Mackinnon, A.C., Jr.; Joseph, J.; Dwinell, M.B.; Kalyanaraman, B. Mitochondria-targeted vitamin E analogs inhibit breast cancer cell energy metabolism and promote cell death. BMC Cancer 2013, 13, 285.
  117. Smith, R.A.; Murphy, M.P. Animal and human studies with the mitochondria-targeted antioxidant MitoQ. Ann. N. Y. Acad. Sci. 2010, 1201, 96–103.
  118. Fink, B.D.; Herlein, J.A.; Yorek, M.A.; Fenner, A.M.; Kerns, R.J.; Sivitz, W.I. Bioenergetic effects of mitochondrial-targeted coenzyme Q analogs in endothelial cells. J. Pharmacol. Exp. Ther. 2012, 342, 709–719.
  119. Lowes, D.A.; Wallace, C.; Murphy, M.P.; Webster, N.R.; Galley, H.F. The mitochondria targeted antioxidant MitoQ protects against fluoroquinolone-induced oxidative stress and mitochondrial membrane damage in human Achilles tendon cells. Free Radic. Res. 2009, 43, 323–328.
  120. McManus, M.J.; Murphy, M.P.; Franklin, J.L. Mitochondria-derived reactive oxygen species mediate caspase-dependent and -independent neuronal deaths. Mol. Cell Neurosci. 2014, 63, 13–23.
  121. Murphy, M.P.; Hartley, R.C. Mitochondria as a therapeutic target for common pathologies. Nat. Rev. Drug Discov. 2018, 17, 865–886.
  122. Socaciu, A.I.; Ionut, R.; Socaciu, M.A.; Ungur, A.P.; Barsan, M.; Chiorean, A.; Socaciu, C.; Rajnoveanu, A.G. Melatonin, an ubiquitous metabolic regulator: Functions, mechanisms and effects on circadian disruption and degenerative diseases. Rev. Endocr. Metab. Disord. 2020, 21, 465–478.
  123. Reiter, R.J.; Mayo, J.C.; Tan, D.X.; Sainz, R.M.; Alatorre-Jimenez, M.; Qin, L. Melatonin as an antioxidant: Under promises but over delivers. J. Pineal Res. 2016, 61, 253–278.
  124. Colunga Biancatelli, R.M.L.; Berrill, M.; Mohammed, Y.H.; Marik, P.E. Melatonin for the treatment of sepsis: The scientific rationale. J. Thorac. Dis. 2020, 12, S54–S65.
  125. Andersen, L.P.; Werner, M.U.; Rosenkilde, M.M.; Fenger, A.Q.; Petersen, M.C.; Rosenberg, J.; Gogenur, I. Pharmacokinetics of high-dose intravenous melatonin in humans. J. Clin. Pharmacol. 2016, 56, 324–329.
  126. Posadzki, P.P.; Bajpai, R.; Kyaw, B.M.; Roberts, N.J.; Brzezinski, A.; Christopoulos, G.I.; Divakar, U.; Bajpai, S.; Soljak, M.; Dunleavy, G.; et al. Melatonin and health: An umbrella review of health outcomes and biological mechanisms of action. BMC Med. 2018, 16, 18.
  127. Poeggeler, B.; Saarela, S.; Reiter, R.J.; Tan, D.X.; Chen, L.D.; Manchester, L.C.; Barlow-Walden, L.R. Melatonin—A highly potent endogenous radical scavenger and electron donor: New aspects of the oxidation chemistry of this indole accessed in vitro. Ann. N. Y. Acad. Sci. 1994, 738, 419–420.
  128. Tan, D.X.; Manchester, L.C.; Reiter, R.J.; Qi, W.B.; Karbownik, M.; Calvo, J.R. Significance of melatonin in antioxidative defense system: Reactions and products. Biol. Sign. Recept 2000, 9, 137–159.
  129. Tan, D.X.; Manchester, L.C.; Terron, M.P.; Flores, L.J.; Reiter, R.J. One molecule, many derivatives: A never-ending interaction of melatonin with reactive oxygen and nitrogen species? J. Pineal Res. 2007, 42, 28–42.
  130. Venegas, C.; Garcia, J.A.; Escames, G.; Ortiz, F.; Lopez, A.; Doerrier, C.; Garcia-Corzo, L.; Lopez, L.C.; Reiter, R.J.; Acuna-Castroviejo, D. Extrapineal melatonin: Analysis of its subcellular distribution and daily fluctuations. J. Pineal Res. 2012, 52, 217–227.
  131. Tan, D.-X.; Manchester, L.C.; Esteban-Zubero, E.; Zhou, Z.; Reiter, R.J. Melatonin as a potent and inducible endogenous antioxidant: Synthesis and metabolism. Molecules 2015, 20, 18886–18906.
  132. He, C.; Wang, J.; Zhang, Z.; Yang, M.; Li, Y.; Tian, X.; Ma, T.; Tao, J.; Zhu, K.; Song, Y.; et al. Mitochondria Synthesize Melatonin to Ameliorate Its Function and Improve Mice Oocyte’s Quality under in Vitro Conditions. Int. J. Mol. Sci. 2016, 17, 939.
  133. Lopez, A.; Garcia, J.A.; Escames, G.; Venegas, C.; Ortiz, F.; Lopez, L.C.; Acuna-Castroviejo, D. Melatonin protects the mitochondria from oxidative damage reducing oxygen consumption, membrane potential, and superoxide anion production. J. Pineal Res. 2009, 46, 188–198.
  134. Paradies, G.; Paradies, V.; Ruggiero, F.M.; Petrosillo, G. Protective role of melatonin in mitochondrial dysfunction and related disorders. Arch. Toxicol. 2015, 89, 923–939.
  135. Acuna Castroviejo, D.; Lopez, L.C.; Escames, G.; López, A.; Garcia, J.A.; Reiter, R.J. Melatonin-mitochondria interplay in health and disease. Curr. Top. Med. Chem. 2011, 11, 221–240.
  136. Vriend, J.; Reiter, R.J. Melatonin, bone regulation and the ubiquitin-proteasome connection: A review. Life Sci. 2016, 145, 152–160.
  137. Janjetovic, Z.; Jarrett, S.G.; Lee, E.F.; Duprey, C.; Reiter, R.J.; Slominski, A.T. Melatonin and its metabolites protect human melanocytes against UVB-induced damage: Involvement of NRF2-mediated pathways. Sci. Rep. 2017, 7, 1–13.
  138. Zhai, M.; Li, B.; Duan, W.; Jing, L.; Zhang, B.; Zhang, M.; Yu, L.; Liu, Z.; Yu, B.; Ren, K.; et al. Melatonin ameliorates myocardial ischemia reperfusion injury through SIRT3-dependent regulation of oxidative stress and apoptosis. J. Pineal Res. 2017, 63, e12419.
  139. Wu, J.Y.; Tsou, M.Y.; Chen, T.H.; Chen, S.J.; Tsao, C.M.; Wu, C.C. Therapeutic effects of melatonin on peritonitis-induced septic shock with multiple organ dysfunction syndrome in rats. J. Pineal Res. 2008, 45, 106–116.
  140. Zhao, K.; Zhao, G.M.; Wu, D.; Soong, Y.; Birk, A.V.; Schiller, P.W.; Szeto, H.H. Cell-permeable peptide antioxidants targeted to inner mitochondrial membrane inhibit mitochondrial swelling, oxidative cell death, and reperfusion injury. J. Biol. Chem. 2004, 279, 34682–34690.
  141. Szeto, H.H. Mitochondria-targeted cytoprotective peptides for ischemia-reperfusion injury. Antioxid. Redox Signal. 2008, 10, 601–619.
  142. Szeto, H.H. Development of mitochondria-targeted aromatic-cationic peptides for neurodegenerative diseases. Ann. N. Y. Acad. Sci. 2008, 1147, 112–121.
  143. Birk, A.V.; Liu, S.; Soong, Y.; Mills, W.; Singh, P.; Warren, J.D.; Seshan, S.V.; Pardee, J.D.; Szeto, H.H. The mitochondrial-targeted compound SS-31 re-energizes ischemic mitochondria by interacting with cardiolipin. J. Am. Soc. Nephrol. 2013, 24, 1250–1261.
  144. Birk, A.V.; Chao, W.M.; Bracken, C.; Warren, J.D.; Szeto, H.H. Targeting mitochondrial cardiolipin and the cytochrome c/cardiolipin complex to promote electron transport and optimize mitochondrial ATP synthesis. Br. J. Pharmacol. 2014, 171, 2017–2028.
  145. Iyer, S.S.; He, Q.; Janczy, J.R.; Elliott, E.I.; Zhong, Z.; Olivier, A.K.; Sadler, J.J.; Knepper-Adrian, V.; Han, R.Z.; Qiao, L.; et al. Mitochondrial Cardiolipin Is Required for Nlrp3 Inflammasome Activation. Immunity 2013, 39, 311–323.
  146. Birk, A.V.; Chao, W.M.; Liu, S.; Soong, Y.; Szeto, H.H. Disruption of cytochrome c heme coordination is responsible for mitochondrial injury during ischemia. Biochim. Biophys. Acta 2015, 1847, 1075–1084.
  147. Szeto, H.H. First-in-class cardiolipin-protective compound as a therapeutic agent to restore mitochondrial bioenergetics. Br. J. Pharmacol. 2014, 171, 2029–2050.
  148. Szeto, H.H. Pharmacologic Approaches to Improve Mitochondrial Function in AKI and CKD. J. Am. Soc. Nephrol. 2017, 28, 2856–2865.
  149. Sabbah, H.N.; Gupta, R.C.; Kohli, S.; Wang, M.; Hachem, S.; Zhang, K. Chronic Therapy With Elamipretide (MTP-131), a Novel Mitochondria-Targeting Peptide, Improves Left Ventricular and Mitochondrial Function in Dogs With Advanced Heart Failure. Circ. Heart Fail. 2016, 9, e002206.
  150. Righi, V.; Constantinou, C.; Mintzopoulos, D.; Khan, N.; Mupparaju, S.P.; Rahme, L.G.; Swartz, H.M.; Szeto, H.H.; Tompkins, R.G.; Tzika, A.A. Mitochondria-targeted antioxidant promotes recovery of skeletal muscle mitochondrial function after burn trauma assessed by in vivo 31P nuclear magnetic resonance and electron paramagnetic resonance spectroscopy. FASEB J. 2013, 27, 2521–2530.
  151. Szeto, H.H. Stealth Peptides Target Cellular Powerhouses to Fight Rare and Common Age-Related Diseases. Protein. Pept. Lett. 2018, 25, 1108–1123.
  152. Orr, A.L.; Ashok, D.; Sarantos, M.R.; Shi, T.; Hughes, R.E.; Brand, M.D. Inhibitors of ROS production by the ubiquinone-binding site of mitochondrial complex I identified by chemical screening. Free Radic. Biol. Med. 2013, 65, 1047–1059.
  153. Orr, A.L.; Vargas, L.; Turk, C.N.; Baaten, J.E.; Matzen, J.T.; Dardov, V.J.; Attle, S.J.; Li, J.; Quackenbush, D.C.; Goncalves, R.L. Suppressors of superoxide production from mitochondrial complex III. Nat. Chem. Biol. 2015, 11, 834.
  154. Wong, H.S.; Monternier, P.A.; Brand, M.D. S1QELs suppress mitochondrial superoxide/hydrogen peroxide production from site IQ without inhibiting reverse electron flow through Complex I. Free Radic. Biol. Med. 2019, 143, 545–559.
  155. Watson, M.A.; Wong, H.S.; Brand, M.D. Use of S1QELs and S3QELs to link mitochondrial sites of superoxide and hydrogen peroxide generation to physiological and pathological outcomes. Biochem. Soc. Trans. 2019, 47, 1461–1469.
  156. Brand, M.D. Mitochondrial generation of superoxide and hydrogen peroxide as the source of mitochondrial redox signaling. Free Radic. Biol. Med. 2016, 100, 14–31.
  157. Fang, J.; Wong, H.S.; Brand, M.D. Production of superoxide and hydrogen peroxide in the mitochondrial matrix is dominated by site IQ of complex I in diverse cell lines. Redox Biol. 2020, 37, 101722.
  158. Brand, M.D.; Goncalves, R.L.; Orr, A.L.; Vargas, L.; Gerencser, A.A.; Borch Jensen, M.; Wang, Y.T.; Melov, S.; Turk, C.N.; Matzen, J.T.; et al. Suppressors of Superoxide-H2O2 Production at Site IQ of Mitochondrial Complex I Protect against Stem Cell Hyperplasia and Ischemia-Reperfusion Injury. Cell Metab. 2016, 24, 582–592.
  159. Piao, L.; Fang, Y.H.; Hamanaka, R.B.; Mutlu, G.M.; Dezfulian, C.; Archer, S.L.; Sharp, W.W. Suppression of Superoxide-Hydrogen Peroxide Production at Site IQ of Mitochondrial Complex I Attenuates Myocardial Stunning and Improves Postcardiac Arrest Outcomes. Crit. Care Med. 2020, 48, e133–e140.
  160. Goncalves, R.L.; Watson, M.A.; Wong, H.-S.; Orr, A.L.; Brand, M.D. The use of site-specific suppressors to measure the relative contributions of different mitochondrial sites to skeletal muscle superoxide and hydrogen peroxide production. Redox Biol. 2020, 28, 101341.
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