Melatonin Protects Mitochondria and ATP Production: History
View Latest Version
Please note this is an old version of this entry, which may differ significantly from the current revision.
Contributor:
Doris Loh

Melatonin (N-acetyl-5-methoxytryptamine) is a ubiquitous, mitochondria-targeted molecule present in all tested eukarya and bacteria. In March 2022, the first discovery of the serotonin N-acetyltransferase (SNAT) gene—responsible for the penultimate formation of N-acetylserotonin (NAS) before its final conversion into melatonin — in archaea further consolidates the status of melatonin as a regulator of biomolecular condensates in all three domains of life in the cellular empire. RNA viruses including SARS-CoV-2 contain proteins with intrinsically disordered regions that undergo liquid-liquid phase separation (LLPS). Liquid-liquid phase separation (LLPS) of these proteins form membraneless condensates that act as “viral factories” to facilitate and enhance replication. Phase separation of the SARS-CoV-2 nucleocapsid (N) protein is associated with mitochondrial dysfunction and rewiring of energy production away from oxidative phosphorylation (OXPHOS) to favor aerobic glycolysis in cytoplasm. Increased ATP in cytoplasm supports viral replication. Melatonin protects mitochondria from damage, maintains adequate  adenosine triphosphate (ATP) levels to disassemble “viral factories”, and prevents suppression of host antiviral immune responses by inhibiting nucleocapsid phase separation via antioxidant-dependent and -independent means.

  • melatonin
  • Mitochondria
  • ATP

1. Introduction

Mitochondria are the “energy powerhouse of the cell” that control respiration and ATP synthesis [1][2], and mitochondria are directly targeted by viruses during infection to facilitate the modulation of cellular metabolism and innate immunity [3]. The fundamental features of optimal mitochondrial dynamics are characterized by the ability to connect and elongate (fusion), divide (fission), and turnover (mitophagy). Disruption of mitochondrial bioenergetics during viral infections may explain how RNA viruses hijack mitochondrial dynamics to support viral replication and persistence [4]. Both the hepatitis B and hepatitis C viruses promote chronic liver damage by altering the balance of mitochondrial dynamics towards fission and mitophagy in order to reduce virus-induced apoptosis, thereby enhancing viral persistence [5][6]. The SARS-CoV-2 virus relies on a sophisticated, multipronged approach to commandeer and manipulate mitochondrial dynamics and metabolism, evading mitochondria-dependent immune response to promote viral replication and pathogenesis [7]. The SARS-CoV-2 dsRNA, which is an intermediate of positive-strand RNA virus replication, has been found to localize in mitochondria [8], while computational modeling of SARS-CoV-2 viral RNA subcellular localization revealed much stronger transcript residency signals toward the mitochondrial matrix and nuclear compartments compared to other coronaviruses [9]. An analysis of changes in molecular composition of mitochondria captured by Raman microspectrometry and biomolecular component analysis (BCA) algorithm found a marked reduction in mtDNA content in microglia treated with spike protein or heat-inactivated SARS-CoV-2 virus [10].
Integrative imaging techniques provided evidence of extensive alterations to cellular organelles, including significant fragmentation of the Golgi apparatus and perturbation of mitochondrial morphology and function. Mitochondria in cells infected by SARS-CoV-2 displayed swollen cristae and matrix condensation, together with significant decreases in mitochondrial ATP synthase subunit 5B (ATP5B) that implies metabolic rewiring away from oxidative phosphorylation in favor of glycolysis [11][12]. The SARS-CoV-2 virus enhances replication by causing mitochondrial dysfunction via membrane depolarization and mitochondrial permeability transition pore (mPTP) opening in a time-dependent manner, with more damage observed at 12 h post-infection compared to 3 h. In order to prevent clearance and degradation of damaged mitochondria, the SARS-CoV-2 virus stalls initiated mitophagy to suppress mitochondrial quality control and clearance of virus by inhibiting binding of mitophagy mediator LC3 and its binding adaptor protein p62 [8][13]. In diabetic cardiomyopathy (DCM), the clearance of dysfunctional mitochondria by mitophagy is often impaired. In a DCM mouse model, melatonin supplementation at 20 mg/kg/day for 4 weeks increased the expression of both LC3-II and p62, resulting in upregulated Parkin-mitophagy that increased clearance of dysfunctional mitochondria to restore mitochondrial quality control [14].

2. Melatonin Rescues Mitochondrial Membrane Potential from SARS-CoV-2 Envelope Protein-Induced Depolarization

RNA viruses and bacterial infections promote ion channel activities, resulting in membrane depolarization that can activate pro-inflammatory, apoptotic NLR pyrin domain containing 3 (NLRP3) inflammasomes that are a major source of inflammatory IL-1β and IL-18 cytokines [15][16][17][18]. The SARS-CoV envelope (E) protein is a viroporin that regulates host cell microenvironment including pH and ion concentrations, causing death in humans and animal models by inducing the pro-inflammatory NLRP3 inflammasome response [19][20][21]. Using similar mechanisms, the SARS-CoV-2 E protein also increases pathogenicity by forming a homopentameric cation channel to modify host ion channel homeostasis in support of viral replication [22][23][24][25]. Mutations of the E protein can enhance the open channel conformation in ion-channel functionality, causing increased virulence and pathogenicity that are correlated with high COVID-19 mortalities [26]. Ion channels formed by viroporins not only allow water and ions to penetrate cell membranes [27], but also generate progressive membrane permeation and damage, disrupting membrane potential and collapsing ionic gradients that facilitate viral budding and release, spreading the virus to surrounding cells [28][29]. Molecular dynamic simulations demonstrated that the E protein can promote viral replication by reducing intracellular calcium in transfected cells and enhance viral budding by bending surrounding lipid bilayers [30].

2.1. Membrane Depolarization Impairs Oxidative Phosphorylation and Cation Homeostasis

Mitochondria infected by SARS-CoV-2 display swollen cristae [11][31][32]. Modulations to cristae topology directly affects mitochondrial function and bioenergetics [33]. ATP synthesis during oxidative phosphorylation (OXPHOS) in mitochondria is dependent upon the F1F0 ATP synthase (complex V) of the electron transport chain (ETC) to drive proton re-entry powered by chemical energy maintained by the negative membrane potential (ΔΨm) of inner mitochondrial membrane (IMM) consisting of inner boundary membranes (IBMs) and cristae—the principal site of oxidative phosphorylation in mitochondria [34][35][36][37]. Changes in the ΔΨm—depolarization or hyperpolarization—by a decrease (less negative) or an increase (more negative) of the ΔΨm, respectively, can alter mitochondrial homeostasis and bioenergetics [34]. Proper ΔΨm of IBM maintains a strong electrical force to keep protons close to cristae membrane within the intercristal space (ICS; cristae lumen) [38][39][40]. Depolarization of the mitochondria membrane can cause a partial or complete collapse of the ΔΨm [41], resulting in dysfunctional, swollen, unfolded cristae that no longer can maintain optimal ATP production via OXPHOS [42]. Decline of the ΔΨm causes matrix condensation, leading to the unfolding of cristae which expands matrix volume to cause mitochondrial swelling [43][44]. Decreased ΔΨm reduces ATP production by lowering ETC activities, but targets damaged areas for clearance by mitophagy [45][46][47]. Yet, inhibition of mitophagy by SARS-CoV-2 prevents the timely clearance of dysfunctional mitochondria that prevents higher ATP production via OXPHOS in favor of glycolysis [12][48][49][50].
Membrane depolarization from viroporin ion channel activities can elevate production of reactive oxygen species (ROS) via increased matrix pH due to cation influx and/or anion efflux [51]. Depolarization opens different types of voltage-gated calcium channels (VGCCs) in a wide range of cell types including both excitable and nonexcitable cells [52][53]. Opening of VGCCs allows the rapid influx of extracellular calcium (Ca2+) that serves as electrical signaling messengers to initiate different important cellular processes [54]. Viruses—including the poliovirus [55], alphavirus [56], human immunodeficiency virus type I (HIV-1) [57], influenza virus [58], SARS-CoV [59], and SARS-CoV-2 [22]—encode viroporins to form ion channels in host cell membranes that facilitate membrane permeability to promote viral entry, replication, release, and dissemination to surrounding cells [59]. Dysregulated calcium signaling may underlie autonomic dysfunctions [60][61] often associated with post-acute sequelae of COVID-19 (PASC) [62][63][64], including postural orthostatic tachycardia syndrome (POTS) [65][66]. Unlike viroporins of other viruses that increase intracellular Ca2+ by modulating plasma membrane permeability [67][68], the SARS-CoV-2 E protein can decrease Ca2+ content in transfected cells by ~61.5% (0.1286 ± 0.0745 AU, N = 22) compared to nontransfected cells (0.2002 ± 0.096, N = 19; p = 0.01), indicating potential leakage, suppression, or sequestration of Ca2+ by the virus. Secondary osteoporosis often occurs with PASC, where a decrease in bone mineral density (BMD) by a mean of 8.6% (± 10.5%) could be detected in COVID-19 at a mean of 81 (± 48) days after hospital discharge. This significant loss in BMD far exceeded normal age-related annual BMD loss, resulting in a two-fold increase in the osteoporosis ratio [69].
Furthermore, the SARS-CoV-2 E protein is localized intracellularly and may be responsible for proton efflux in transfected cells [30]. An acidic pH can adjust the conductivity and ion selectivity of the ion-conducting transmembrane domain of E protein by protonating the Glu8 side chain carboxyl, altering the carboxy-terminal conformation [24]. The influenza B virus viroporin proton channel is pH-gated and mediates virus uncoating when activated by acidic pH [70]. Ionic imbalances in cells affecting the homeostasis of cations, including calcium (Ca2+), magnesium (Mg2+), zinc (Zn2+), potassium (K+), and sodium (Na+), can interfere with innate and adaptive immunity that affect the pathogenicity of viruses [19][71][72][73].

2.2. Viroporin Ion Channel Activities May Regulate Virus Phase Separation

Potassium (K+) efflux triggers the activation of the NLRP3 inflammasome upon infection by RNA viruses [15], including SARS-CoV-2 [74][75], where elevated urinary loss of potassium is often associated with COVID-19 disease severity [76]. Experimental work showed the SARS-CoV-2 ORF3a viroporin priming and activation of the NLRP3 inflammasome were dependent upon K+ efflux [77]. K+ is a rate-liming modulator of the glutamate transport cycle, where intracellular K+ relocates the glutamate binding site to the extracellular side of the membrane, and extracellular K+ induces glutamate release upon transporter relocation [78]. Glutamate promotes LLPS of the Escherichia coli single-stranded-DNA binding protein [79]. Thus, K+ efflux that can elevate glutamate availability [80] may enhance SARS-CoV-2 phase separation. Indeed, altered glutamine metabolism and dependence on glutamine receptor subtype 2 for internalization are associated with SARS-CoV-2 infections [81][82]. Mitochondrial dynamics dysfunction and Ca2+ dysregulation as a result of membrane depolarization induced by viroporin ion channel activities can also affect leucocyte functionality to suppress and evade immune responses during SARS-CoV-2 infection to enhance viral phase separation for viral replication.

3. Melatonin Attenuates Membrane Depolarization and Balances Ion Homeostasis by Antioxidant-Dependent and -Independent Mechanisms to Protect Mitochondria and Lymphocytes during Viral Infection and PASC

Leukocytes of patients recovered from COVID-19 presented loss of mitochondria membrane potential (ΔΨm) even at 11 months post-infection [83]. Leukocytes are responsible for the production of first line IFN-⍺ immune response [84][85], and the loss of ΔΨm caused by viroporin-mediated membrane depolarization may be one of the most important underlying causes for the development of PASC [83]. Lymphopenia and the depletion of T lymphocyte subsets were found in 98% (153/157) of patients infected by SARS-CoV in 2003 without any preexisting hematological disorders [86]. Correspondingly, patients infected by SARS-CoV-2 are associated with persistent lymphopenia [87][88] and functional exhaustion of lymphocytes [89]. COVID-19 disease progression is correlated with a nearly three-fold increased risk of severe COVID-19 (random effects model, OR = 2.99, 95% CI: 1.31–6.82) [90], while low lymphocyte counts in patients are deemed to be effective predictors of disease severity and hospitalization [91][92].
T lymphocytes are dependent upon functional mitochondria to supply local ATP and to maintain Ca2+ homeostasis and signaling during all stages of immune response [93][94]. In T lymphocytes, expression of 75% of the genes associated with survival and proliferation are dependent upon Ca2+ influx [95], while mitochondrial dynamics often affect T lymphocyte chemotaxis, where mitochondrial fusion protein OPA1 inhibits lymphocyte migration and chemotaxis, but fission enhances both migration and chemotaxis [96]. It is perhaps not a coincidence that depolarization of mitochondrial membranes can activate dynamin-related GTPase OPA1-dependent fusion to inhibit lymphocyte chemotaxis [97], and that the E protein viroporin can deplete intracellular Ca2+ content [30]. Stimulation of T lymphocytes triggers immediate accumulation of active mitochondria with elevated Ca2+ influx and heightened OXPHOS, which can also cause transient collapse of ΔΨm due to intense ETC activities, ion flux, and ATP release across the mitochondrial membrane [93]. Thus, inability to repolarize ΔΨm results in a reduction of ATP generation from the loss of electrochemical potential that maintains the gradient that drives the F1F0 ATP synthase (complex V) [45][98]. Moreover, membrane depolarization also prevents store-operated Ca2+ influx after store depletion [99]. Cell sorting experiments revealed that mtDNA damage occurs only in human fibroblast cells with low ΔΨm sustained for 24 h. These cells exhibited continuous, elevated production of hydrogen peroxide (H2O2) that potentially accentuated a feed-forward cascade of increasing ROS that impaired repair responses and increased mtDNA lesions, resulting in apoptosis [100]. Taken together, membrane depolarization by E protein suppresses not only ATP-dependent purinergic signaling that supports T lymphocyte immune response functions, but also T lymphocyte-mediated expression of genes that are dependent upon Ca2+ influx [93][95]. In its multipronged strategies against the SARS-CoV-2 virus, melatonin not only promotes the production of leukocytes [101], but also attenuates membrane depolarization to protect lymphocyte functionality (Figure 1).
Ijms 23 08122 g001 550
Figure 1. Schematic illustrating melatonin attenuation of acute infection, viral persistence, and post-acute sequelae COVID-19 (PASC) from potential alterations to the epitranscriptome and transcriptome via RNA m6A modifications and LINE1 derepression by the SARS-CoV-2 virus. The envelope (E) protein causes extensive mitochondrial distress and elevates oxidative stress via membrane depolarization and ionic imbalances that activate LINE1 derepression, NLRP3 inflammasome apoptotic signaling, stress granule formation, and nucleocapsid (N) protein liquid-liquid phase separation (LLPS). N protein LLPS forms membraneless condensates that not only facilitate viral transcription, genome packaging, and dissemination, but also enhance the suppression of host gene expression to evade innate immune responses via the disassembly of stress granules and the hijacking of DEAD-box RNA helicase DDX3X. Melatonin employs antioxidant-dependent and -independent strategies to modulate m6A modifications, suppress LINE1 derepression, rescue mitochondrial dysfunctions, and reduce oxidative stress. Melatonin regulates N protein LLPS to block the sequestration of DDX3X and the formation of NLRP3 inflammasome, as well as the disassembly of stress granules to support innate antiviral immune response, inhibiting viral transcription and replication, maintaining host gene stability and integrity to prevent severe disease and PASC (see Abbreviations for additional acronyms).
Melatonin is a pleiotropic molecule that can maintain optimal membrane potential by either increasing or reducing ΔΨm for maximum efficiency. In hyperpolarized, prorenin-treated microglia, treatment with 100 μM melatonin reduced ΔΨm and attenuated hyperpolarization and ROS overproduction [102]. Conversely, in mitochondria of human oocytes, 10 µM melatonin treatment decreased excessive intracellular Ca2+ levels to restore mitochondrial function and significantly increased membrane potential compared to control levels [103], while 1 μM melatonin added to post-thawed equine sperm increased mitochondrial membrane potential and improved mitochondrial function [104]. Membrane depolarization prevents store-operated Ca2+ influx after store depletion [99], disrupting T lymphocyte-mediated gene expressions [95]. However, treatment with 500 µM melatonin markedly elevated cytosolic calcium in human platelets by evoking store-operated calcium release from platelet mitochondria [105]. An analysis of human neutrophil respiratory burst and membrane potential changes found melatonin to increase depolarization at concentrations up to 0.5 mM, whereas 2 mM melatonin concentration decreased ΔΨm in neutrophils activated by phorbol 12-myristate 13-acetate (PMA) [106]. Mitochondrial inner membrane depolarization in human HaCaT keratinocytes irradiated with UVB radiation (50 mJ/cm2) was normalized by preincubation with 0.01 mM to 1 mM melatonin via the reduction of mitochondrial ROS (mROS) and inhibition of mitochondrial permeability transition pore (mPTP) opening [107].
Viroporin-induced membrane depolarization elevates production of ROS via ionic imbalances from dysregulated cation influx and/or anion efflux [51]. The SARS-CoV-2 virus can also escalate ROS release in Vero E6 cells via opening the mPTP, causing subsequent depolarization and further oxidative stress damage in a time-dependent manner [8][108]. In a self-perpetuating positive feedback loop, oxidative stress from unneutralized excess ROS leads to even more rapid depolarization of the inner mitochondrial membrane potential and subsequent disruption of OXPHOS and ATP production. Damaged mitochondria continue to produce more ROS, resulting in the dreaded ROS-induced ROS release (RIRR) loop [109]. ROS can also cause physiological lipid peroxidation [110], where oxidants attack the carbon-carbon double bond in lipids, initiating a cascading chain reaction that terminates in the formation of reactive aldehyde end products including 4-hydroxynonenal (HNE) [111]. In a pilot study of 21 critically ill COVID-19 patients admitted to the ICU, the only difference in clinical or laboratory parameters monitored between the 14 patients who recovered and the 7 who passed away was the significantly higher level of HNE-protein adducts (p < 0.05) obtained from the plasma of the deceased patients compared to levels in survivors during the initial 1–3 days in hospital [112].
Melatonin and its metabolites are potent inhibitors of lipid peroxidation cascades and are extremely effective at scavenging different types of ROS [113][114][115][116][117][118][119]. In leukocytes irradiated with 750 mJ/cm2 UVB light (280–360 nm, max: 310 nm), treatment with melatonin suppressed ROS directly in a dose-dependent manner where 10 mM melatonin reduced ROS formation in leukocytes by 260-fold, while 7.5 mM and 5 mM reduced ROS by 120- and 60-fold, respectively [120]. In addition to decreasing ROS via antioxidant-dependent mechanisms [121][122], the regulation of depolarization by melatonin may be via an ionic-based, antioxidant-independent mechanism. The repolarization of gonadotrophin-releasing hormone (GnRH)-induced membrane depolarization in neonatal rat pituitary cells by melatonin could be mediated through the inhibition of Ca2+ influx or a hyperpolarization mechanism that is sodium-dependent, involving modulation of the Na+/K+-dependent ATPase [123]. Jurkat cells undergo apoptosis from anti-Fas-induced mitochondrial membrane depolarization where inhibition of the Na+/K+ ATPase prevented membrane repolarization via the suppression of monovalent ion movements, particularly the intracellular accumulation of Na+ during sustained depolarization without repolarization [124].
Melatonin is an osmoregulator with pleiotropic effects on plasma sodium concentration in animal models [125][126]. This ancient molecule is indispensable in maintaining ion homeostasis in plants [127][128][129], and its comprehensive role as a “broad-based metabolic buffer” includes rhythmic circadian modulation of the Na+/K+-ATPase as well as the Na+/H+ exchanger ion-transport activities in human erythrocytes via antioxidant-dependent and -independent mechanisms [130][131]. Both Na+/K+-ATPase and Na+/H+ can influence transmembrane chemical gradients [132][133], as well as cytosolic pH and ionic balance [134][135][136]. Therefore, it is not inconceivable that melatonin can adjust salt homeostasis via Na+/K+-ATPase to regulate LLPS during viral infections as high salt or extremely low salt concentration can inhibit LLPS [137]. Hyponatremia where plasma sodium concentration is below 135 mmol/L is often associated with viral infections including COVID-19 [138]. Furthermore, in vitro experiments found 1.5% NaCl solution can achieve 100% inhibition of SARS-CoV-2 replication in nonhuman primate kidney Vero cells, while 1.1% of NaCl can inhibit viral replication by 88% in human epithelia lung Calu-3 cells [139].
The Na+/K+-ATPase is a P-type ATPase that utilizes energy from ATP hydrolysis to pump ions across membranes generating an electrochemical gradient [140]. Nonmitochondrial ATPases including P-type Na+/K+-ATPases are often localized in lipid raft microdomains in lipid bilayers of plasma membranes [141][142][143]. Increased ROS from oxidative stress can reduce membrane fluidity and performance of Na+/K+-ATPases [144][145][146][147]. Melatonin maintains membrane fluidity by inhibiting lipid peroxidation cascades in an antioxidant-dependent manner [114][115][148][149][150], while its ability to stabilize liquid-ordered (Lo)-liquid-disordered (Ld) phase separation in lipid bilayers (tested over a range of temperatures up to 45 °C) preserves necessary lipid raft composition and nanoscopic structure to support various ATPase activities, including those of Na+/K+-ATPases [130][151].
An analysis of information obtained from various neutron scattering techniques accessing membrane structure and dynamics from SARS-CoV-2 protein–host interactions revealed that molecular interactions during spike protein fusion peptide binding events could induce changes in membrane fluidity and rigidity where fusion peptide 1 increased rigidity while fusion peptide 2 reduced fluidity [152]. Other morphological changes induced by SARS-CoV-2 as a result of fusion events include modification of both lipid composition and membrane structure to produce non-lamellar cubic membranes that facilitate membrane fusion during viral infection [153]. The oxidation of high curvature lipids such as cardiolipin (CL) can result in the rearrangement of lipids in plasma membranes from a fluid lamellar phase to a non-lamellar cubic phase that can impact membrane integrity and stability. The fact that cubic membranes are usually found in membranes with high intrinsic curvature, such as mitochondrial inner membranes with deep cristae invaginations formed by high-curvature lipids that host ATP synthase dimers [154][155][156], further explains how SARS-CoV-2 and other viruses modulate mitochondrial function to favor glycolysis over OXPHOS.

4. Melatonin Protects Mitochondria Cristae Morphology and ATP Production via Antioxidant-Dependent and -Independent Mechanisms

Phase separation of SARS-CoV-2 N protein may be biphasically modulated by ATP where ATP can completely dissolve viral condensates, which promote pathogenicity and replication, formed by N protein LLPS at molar ratios of 1:500 (N-protein:ATP), but enhance assembly of condensates from low molar ratios of 1:25 up to 1:200 [157]. Hence, mechanisms associated with viral fusion and enhanced viral replication involve targeting of mitochondrial bioenergetics and the production of ATP. An analysis of bulk RNA-seq datasets from COVID-19 patients and healthy controls revealed a marked reduction of mtDNA gene expression in various types of cells including the immune system, with concomitant elevation of genes expressing glycolytic enzymes, and ROS production [48], while an interactome analysis identified multiple mitochondrial proteins that interact with the SARS-CoV-2 N protein [158]. Elevated glucose and sustained aerobic glycolysis in monocytes of COVID-19 patients are directly responsible for boosting viral replication, causing increased NLRP3 inflammasome and cytokine production, inhibition of T proliferation, and apoptosis of lung epithelial cells [50][159]. Metabolic alterations in live peripheral blood mononuclear cells (PBMC) obtained from patients with COVID-19 showed extensive mitochondrial dysfunction with compromised respiration but increased utilization of glucose serving as primary substrate for energy production in place of OXPHOS [160]. Substituting OXPHOS ATP production with aerobic glycolysis may lead to a more than 16-fold reduction of ATP. The theoretical maximum of ATP calculated from simultaneous measurements of oxygen consumption and extracellular acidification showed OXPHOS to yield 31.45 ATP/glucose (maximum total yield 33.45), whereas glucose yields only 2 ATP/glucose [161]. Considering ATP can completely dissolve N protein phase separation condensates at concentrations 2.5- to 20-fold above assembly concentrations, with disassembly starting beyond 8-fold increases, it is not surprising that the timely application of melatonin can effectively suppress viral replications.

4.1. Melatonin Suppresses Aerobic Glycolysis to Enhance Oxidative Phosphorylation

Melatonin is a powerful glycolytic that can inhibit aerobic glycolysis (the “Warburg effect”) by steering pyruvate metabolism towards the citric acid (tricarboxylic acid, Krebs) cycle and OXPHOS, and avoiding aerobic fermentation of glucose by glycolysis [162][163][164]. Melatonin can enhance mitochondrial OXPHOS ATP production [165] by different mechanisms including the stimulation of the SIRT3/PDH axis to reverse the Warburg phenotype in lung cancer cells in vitro [166]; and the suppression of hexokinase-2 overexpression to ameliorate glycolytic overload, improving mitochondrial ATP production and normalizing glycolysis to protect mitochondrial function in chronic kidney disease mesenchymal stem/stromal cells [167]. The SARS-CoV E protein ion channel induces membrane permeabilization that decreases ΔΨm in mitochondrial inner membranes [27][83]. Loss of membrane potential not only reduces ATP production due to impaired OXPHOS, but can induce the production of even more ROS due to accumulation of reducing equivalents from lower ETC activities that result in the creation of reductive stress that continues generate additional ROS to perpetuate the RIRR positive feedback loop [34][109][168][169]. The generation of excess ROS during SARS-CoV-2 infection [8] can initiate powerful lipid peroxidation cascades that damage lipid composition of the cristae, resulting in loss of ATP synthase function.

4.2. Melatonin and Metabolites Preserve Cardiolipin Function in Cristae by Preventing Lipid Peroxidation Cascades

The apex of deep IMM cristae invaginations provides the ideal location for hosting dimerized ATP synthases of eukaryotic mitochondria [154][170]. Dimerized ATP synthases are seven-fold more active than ATP monomers [171], and dimerization of ATP synthases is a prerequisite for shaping the high curvature cristae structure [172][173]. The deep negative membrane curvatures at the apexes of cristae are maintained by the unique cone-shaped structure of cardiolipin (CL) that not only increases bending elasticity of the IMM but also the regulation of formation and stability of respiratory chain complexes [174][175][176][177]. Accordingly, mitochondrial membranes can comprise up to 25% CL [178][179]. CL is a negatively charged, dianonic lipid that can dramatically lower pH at membrane interfaces to increase proton (H+) concentration (~700 to ~800) [180][181] to elevate ATP production [182]. The oxidation of just one fatty acid chain in CL can lead to vast conformational changes in the entire molecule, resulting in reduced membrane thickness, and potential impairment of proton and electron transport that are dependent on CL-mitochondrial protein interactions [183][184]. Elevation of ROS as a result of depolarized mitochondrial membranes during viral infection may increase peroxidation of cardiolipin. The destabilization of mitochondrial supercomplexes as a result of CL peroxidation affects mitochondrial bioenergetics, leading to impaired OXPHOS, reduced ATP production, and other mitochondrial dysfunctions in different tissues manifested in a range of pathophysiological conditions including heart ischemia/reperfusion, heart failure, diabetes, and Barth syndrome [185][186][187][188][189][190][191]. In Saccharomyces cerevisiae, disruption of the CRD1 gene responsible for encoding CL synthase resulting in the absence of CL in mitochondria membranes led to a loss of mitochondrial ΔΨm and mitochondrial genome when cultured at prolonged elevated temperature of 37 °C [192]. Interestingly, circulating anticardiolipin antibodies (aCL), which may cause endothelial dysfunction and elevated IgA-aCL, is often associated with increased ischemic burden in patients with coronary artery disease (CAD) [193].
Critically ill COVID-19 patients with coagulopathy and thrombocytopenia often manifest the presence of anticardiolipin antibodies in serum [194]. A meta-analysis and systematic review of 21 studies with 1159 hospitalized COVID-19 patients discovered the presence of antiphospholipid antibodies in ~50% of the patients. Severe disease was correlated with a higher prevalence of aCL (IgM or IgG) compared to noncritical disease (28.8% vs. 7.10%, p < 0.0001) [195]. Oxidized LDL bound by anti-lipoprotein antibodies are correlated with IgG-aCL and IgM-aCL [196]; thus, the presence of elevated aCL and other antiphospholipid antibodies is indicative of systemic lipid peroxidation, which may then explain the development of thromboses in the absence of correlated D dimer levels in about one-third of severely ill COVID-19 patients [193][197]. In fact, elevated lipid peroxidation is the only oxidative stress biomarker that is significantly different between intubated COVID-19 patients and/or those who died compared to patients with mild disease. In addition, patients whose lipid peroxidation rose above 1948.17 μM were either intubated or died 8.4 days earlier on average (mean survival time 15.4 vs. 23.8 days) [198]. Melatonin is a potent antioxidant that can protect mitochondrial function by neutralizing ROS to inhibit CL peroxidation [199]. The addition of 10 μM melatonin to rat heart mitochondria almost entirely prevented membrane depolarization induced by Ca2+/tert-Butylhydroperoxide (t-BuOOH), a peroxidation promoting peroxide, in addition to reversing cytochrome c release, and mitochondrial matrix swelling [116]. The reason why melatonin is uniquely suited to prevent lipid peroxidation cascades is in large part due to its preferential localization at hydrophilic/hydrophobic membrane interfaces.
Melatonin is uncharged in the entire pH range [200]. Even though melatonin is nonpolar, it can form strong H-bonds with hydrophilic lipid headgroups at hydrophilic/hydrophobic membrane interfaces [201]. Thus, melatonin becomes an efficient scavenger of both aqueous and lipophilic free radicals as a result of the presence of both hydrophilic and lipophilic moieties in the melatonin molecule [202]. As such, melatonin and its metabolites easily neutralize both the hydroxyl radical (•OH) and the hydroperoxyl radical (•OOH) [203][204]—two dominant ROS molecules that can initiate and sustain chain oxidation reactions of unsaturated phospholipids including CL in plasma membranes [205][206] and mitochondria [207][208]. During viral infections, ionic imbalances from viroporin ion channel activities activate the pro-inflammatory NLRP3 inflammasome which mediates the production of cytokines that can contribute to severe pathophysiology and disease [17][209]. Heightened expression of the NLRP3 inflammasome was detected in leukocytes in the lungs of all patients who did not survive COVID-19 [210]. Melatonin targets NLRP3 inflammasome-mediated cytokine release employing antioxidant-dependent and -independent mechanisms [211].

5. Melatonin Targets NLRP3 Inflammasomes via Cardiolipin and DDX3X

Cellular stress and dysfunction triggers prionoid-like phase transition of the NLR pyrin domain containing 3 (NLRP3) inflammasome to assemble supramolecular complexes responsible for mediating immune responses, including the release of inflammatory cytokines—IL-1β and IL-18 [212][213][214][215][216]. The NLRP3 inflammasome is a multiprotein complex comprising the NLRP3 sensor, the apoptosis-associated speck-like protein containing a C-terminal caspase recruitment domain (ASC) adaptor, and the caspase-1 (CASP1) protease [217][218]. The activation of NLRP3 inflammasomes is inextricably linked to various types of cell death, including pyroptosis, apoptosis, necroptosis, and ferroptosis [217]. Elevated ROS and mitochondrial distress translocate CL from the inner mitochondrial membrane (IMM) to the outer mitochondrial membrane (OMM) [219], and NLRP3 must be primed and directly bound by externalized CL before it can be activated [220]. As discussed in Section 5.1, viroporin ion channel activities activate NLRP3 inflammasome, and COVID-19 severe pathology resulting from an overactive immune-inflammatory response can be exacerbated by the activation of NLRP3 in infected macrophages in humanized mouse model of COVID-19 [221]. The SARS-CoV-2 E protein viroporin increases NLRP3 inflammasome activation in both murine and human macrophages in a biphasic manner [222] by first suppressing NLRP3 inflammasome activation to aid viral replication leading to advanced disease states that promote the activation of NLRP3 inflammasomes [222]. The activation of NLRP3 inflammasome is often associated with the development of severe COVID-19 [223][224][225] and increased oxidative stress [226], while the production of inflammatory cytokines, including IL-β, may fuel the development of cytokine storms and excess oxidative stress to complete a positive feedback cycle [227][228][229][230][231] that enhances N protein LLPS [232]. This unique biphasic effect may be a reflection of how the SARS-CoV-2 virus interacts with DDX3X and stress granules (SGs) during viral replication.
The regulation of the prionoid transition of NLRP3 inflammasome into supramolecular complexes is mediated by DEAD-box helicase 3 (DDX3X or DDX3)—a host X-chromosome encoded DEAD-box RNA helicase that is often hijacked by SARS-CoV-2 and other viruses [233][234]. In total, 18 species of virus from 12 genera—including the dengue virus [235], HIV-1 virus [236][237], hepatitis C virus [238], Japanese Encephalitis virus, and the Zika virus [239]—have been determined to be dependent upon DDX3X for virulence [240]. The ATP-bound form of DDX3X is necessary for the scaffolding of the ASC domain to transition into irreversible, stable, and insoluble supramolecular prionoid-like assemblies [213]. DDX3X is also a critical regulator of SGs requisite for proper SG maturation [241]. Therefore, the formation of SGs and the assembly of NLRP3 inflammasomes become mutually exclusive, since both SGs and NLRP3 inflammasomes compete for the same DDX3X. Consequently, loss of DDX3X will inhibit both SGs maturation and the scaffolding of ASCs to disrupt NLRP3 inflammasome supramolecular assembly [242][243], while the disassembly of SGs may encourage the aggregation of NLRP3 inflammasomes. Lipid peroxidation that can translocate CL from the IMM to OMM is regarded as a hallmark of severe COVID-19 [198]. Monocytes from severe COVID-19 patients exhibit elevated, persistent presence of ROS and lipid peroxidation compared to mild disease and health controls. The level of lipid peroxidation is strongly correlated with CASP1 activity and ASC aggregate formation, responsible for the NLRP3 inflammasome-dependent IL-β secretion by monocytes [244].
Melatonin targets DDX3X to regulate and enhance innate antiviral responses that suppress viral replication. Viral infection induces cellular stress and mitochondrial distress that activates the host integrated stress response (ISR) resulting in the formation of SGs. The timely, adequate presence of melatonin can reduce ROS and lipid peroxidation to prevent the translocation of CL to OMM, thus inhibiting the activation of NLRP3 and its prionoid phase transition to form inflammasome supramolecular complexes [211][245][246][247][248]. This effectively allows DDX3X to accelerate the formation and maturation of SGs that can enhance antiviral innate immune signaling [249] and also inhibit viral protein accumulation and replication [250][251]. As such, viruses including SARS-CoV-2 have evolved sophisticated mechanisms to hijack DDX3X by disrupting SG formation. The SARS-CoV-2 N protein not only phase separates to form “viral factory” condensates [252] that protect the viral genomic RNA by packing them into distinct RNP complexes [253][254], but also acts as the central hub for DDX3X interactions [255]. In Vero E6 cells infected by SARS-CoV-2, mass spectrometry analysis revealed DDX3X localizes with viral RNA foci in cytoplasm, and enhances viral infection via interactions with N protein [256]. The fact that the immunopurified complexes were harvested 24 h post-infection may also imply that the N protein has already undergone phase separation to form viable “viral factories” that can interact with DDX3X, facilitating immune evasion and suppression.

6. DDX3X Is a “Double-Edged Sword” That Mediates Host Antiviral Immunity and Viral Replication

DDX3X is not only an essential mediator of host innate immunity, but also acts as host factors that assist viral replication [257][258]. Therefore, DDX3X is often targeted and hijacked by viruses during infection to evade immune response and promote replication [240][259][260]. SARS-CoV-2 N protein sequesters and potentially binds to DDX3X in order to inhibit host antiviral pathways [255]. The induction of first line IFN immune defense requires the synergistic activation of the type I IFN-β promoter by DDX3X, and TANK-binding kinase 1 (TBK1) and its interaction partner, DDX3X [154][261]. This synergistic effect on IFN induction is mediated by the recruitment of DDX3X into mitochondrial antiviral-signaling protein (MAVS, IPS-1) to promote the scaffolding and aggregation of MAVS into prion-like complexes that can then activate TBK1 and interferon regulatory factor 3 (IRF3) for type I IFN responses [262][263]. LLPS of N protein inhibits both the polyubiquitination and formation of prion-like aggregates in MAVS, effectively suppressing the host innate antiviral response [264]. The prion-like conformational switch of MAVS on the mitochondrial membrane is the lynchpin that propagates antiviral signaling cascades that can inhibit viral infections [265] and is mediated by DDX3X. Nevertheless, in order to hijack DDX3X, viruses including SARS-CoV-2 must first dismantle the assembly of host SGs that are associated with DDX3X.

7. N Protein Must Phase Separation to Target G3BP1 and Disassemble Stress Granules

Stress granules (SGs) are membraneless organelles assembled as a result of LLPS activated by cellular stress, including viral infections [266][267]. Ras-GTPase-activating protein SH3 domain-binding protein 1 (G3BP1) [268] is the molecular switch that regulates RNA-dependent LLPS of SGs, and its effects on SG LLPS can be tuned by phosphorylation of IDRs in G3BP1 as well as extrinsic binding factors that can strengthen or weaken the SG assembly [269]. G3BP1 promotes SG IFN signaling, enhancing innate antiviral response via positive regulation of RIG-1—an upstream target of MAVS [270][271][272]. Recent biochemical and structural analyses of the interactions between SARS-CoV-2 N protein and G3BP1 revealed that N protein residues 1–25 (N1–25) occupies a conserved surface groove of the NTF2-like domain of G3BP1 (G3BP1NTF2). The interactions between the N1–25 and G3BP1NTF2 are enhanced by strong surface complementarity and hydrophobic groove-insertion mechanisms, resulting in the inhibition of SG assembly. However, the underlying mechanism for SG disassembly by SARS-CoV-2 N protein could not be determined [273]. N protein binding to G3BP1 also rewires the G3BP1 mRNA binding profile to suppress host cell stress response [274]. In order to target G3BP1, the SARS-CoV-2 N protein must first undergo LLPS, partitioning into SGs before it can bind and interact with G3BP1 to dismantle assembly of SGs [275].

8. The Formation of “Viral Factories” by N Protein LLPS Is Tuned by Phosphorylation

Oxidative stress induces the formation of SGs [276], and N protein LLPS induced by oxidative stress in vitro facilitates its partitioning into SGs to sequester G3BP1 [232][277]. Similar to other condensates formed via LLPS, N protein condensates can be tuned by the concentration of RNA where increasing RNA gradient with a fixed protein concentration at 10 μΜ caused N protein to increase viscosity from droplets to gel-like, and, eventually, solid assemblies [275][278], whereas phosphorylation of the serine/arginine (S/R)-rich region in the central IDR of the N protein can tune the viscosity and modulate N protein condensate assembly [279]. Phosphorylation is an ATP-dependent, post-translational modification that can fine-tune molecular interactions of condensate components by inducing nonequilibrium thermodynamic chemical reactions to control the size and number of condensates, acting somewhat like a rheostat that can adjust the dynamics of LLPS during condensate formation [280].
Unphosphorylated N protein facilitates tight association with host mRNAs, and thus, increases the propensity to form gel-like condensates; conversely, phosphorylation of N protein results in the formation of more dynamic, low-viscosity, liquid-like droplets [281]. The EBOV N protein-induced dynamic phosphorylation and dephosphorylation of VP30—the fourth N protein essential for viral transcription—take place in viral inclusion bodies [282][283]. Molecular dynamics simulations revealed that phosphorylation of the phosphate groups at different serine residues in the serine-arginine (SR)-rich domain in SARS-CoV-2 N protein induced the formation of dense salt bridge networks, increasing intra- and intermolecular contacts that impaired contact with RNA derived from SARS-CoV-2 genome, effectively preventing association with nonspecific RNA [253][284]. Thus, the tuning of the physical properties of N protein condensates via phosphorylation can determine whether viral transcription or packaging is favored via hyperphosphorylation (low-viscosity) or hypophosphorylation (high-viscosity), respectively [279][284]. Consequently, high-viscosity, unphosphorylated condensates are more effective at promoting viral packaging—the cytoplasmic compartmentalization of the viral genome—whereas low-viscosity, phosphorylated condensates operate as dynamic “viral factories” to promote viral transcription/replication and host immune evasion [279][281].

9. Melatonin Disrupts Formation of “Viral Factories” by Regulating GSK‐3 Phosphorylation of N Protein Condensates 

The GSK-3 kinase is implicated in enhancing virus replication, assembly, and release [285][286][287]. As part of the innate antiviral response, GSK-3 acts as a signaling molecule that may be involved in the sensing of nucleic acids of RNA and DNA viruses. It is not only responsible for the rapid activation of type I IFN signaling cascades [288], but also serves as the crux of multiple cell signaling pathways during various stages of viral replication [289]. The activation of GSK-3 in infected cells may be responsible for increased replication and pathophysiology by promoting systemic inflammation, renal dysfunction, and hepatotoxicity via the regulation of cytokine production and cell migration [290][291], as well as the transcriptional regulation of nuclear factor kappa B (NF-κB) [292]. GSK-3 also elevates oxidative stress in infected cells by downregulating the Nrf2 and the Nrf2/antioxidant response element (ARE) pathway [293][294]. GSK-3 directly inhibits nuclear factor erythroid 2-related factor (Nrf2) activation and indirectly inhibits Nrf2 post-induction [295]. The increased oxidative stress from GSK-3 activities may induce the assembly of SGs, but more importantly, the activation of GSK-3 may actually be the elusive, underlying mechanism that is responsible for the disassembly of SGs by SARS-CoV-2 N proteins [273]. The phosphorylation of N protein by GSK-3 not only determines the viscosity and function of condensates formed by N protein LLPS, but GSK-3 can also regulate DDX3X functions to control stress granule assembly and disassembly (Figure 2). GSK-3 is responsible for the phosphorylation of Gle1A which is is recruited to SGs in the cytoplasm during stress to regulate SG dynamics, assembly, and disassembly by controlling how DDX3X binds to RNA [296][297]. Phosphorylation of Gle1A by GSK-3 alters the biochemical properties and electrophoretic mobility that allows Gle1A to bind and inhibit DDX3X ATPase activities that ultimately results in the disassembly of SGs. 

Melatonin suppresses hyperphosphorylation of N protein by inhibiting the gene expression of GSK-3 and deactivating GSK-3 by promoting its phosphorylation. In Neuro2A cells subjected to okadaic acid (OA) treatment to induce phosphorylation of tau by GSK-3β exhibited elevated ROS and cytotoxicity, resulting in the loss of cell viability of up to 60%. Incubation with 200 μM melatonin for 24 h completely reversed tau-induced cytotoxicity, while at 100 μM concentration, melatonin completely restored cell viability, where melatonin effectively reduced the total mRNA expression level of GSK-3β [298].

Additionally, in human mesenchymal stem cells, melatonin attenuated adipogenic differentiation by suppressing GSK-3β activities [299]. Male Wistar rats subjected to bilateral renal ischemia to induce ischemia/reperfusion (I/R) injury showed increased lipid peroxidation and elevated lactate dehydrogenase (LDH) in plasma compared to controls. Treatment with melatonin (10 mg/kg, i.p.) 30 min before renal clamping markedly reduced lipid peroxidation and LDH levels in plasma, while the phosphorylation of GSK-3β was significantly enhanced via the restoration of AKT phosphorylation in the melatonin-treated group [300].

10. Conclusion

As COVID-19 transitions inevitably from pandemic to endemic, it is presently unclear how continued endemic infections from evolving SARS-CoV-2 variants will shape human health in the years to come. The detrimental effects of viral replication and persistence causing excess oxidative stress and mitochondrial distress can result in downstream effects that alter both host and viral RNA methylomes. Consequently, SARS-CoV-2 introduces a complex, fertile landscape that fosters a wide-array of challenging and often unexplained manifestations [879] during acute infection and post-infection. The timely application of melatonin as an essential adjuvant during acute infection and recovery can inhibit viral infection, replication, and persistence to prevent the hijacking of mitochondria and other vital host resources associated with immune evasion and suppression.

This entry is adapted from the peer-reviewed paper 10.3390/ijms23158122

References

  1. McBride, H.M.; Neuspiel, M.; Wasiak, S. Mitochondria: More than Just a Powerhouse. Curr. Biol. 2006, 16, R551–R560.
  2. Brown, G.C. Control of Respiration and ATP Synthesis in Mammalian Mitochondria and Cells. Biochem. J. 1992, 284 Pt 1, 1–13.
  3. Elesela, S.; Lukacs, N.W. Role of Mitochondria in Viral Infections. Life 2021, 11, 232.
  4. Khan, M.; Syed, G.H.; Kim, S.-J.; Siddiqui, A. Mitochondrial Dynamics and Viral Infections: A Close Nexus. Biochim. Biophys. Acta 2015, 1853 Pt B, 2822–2833.
  5. Kim, S.-J.; Khan, M.; Quan, J.; Till, A.; Subramani, S.; Siddiqui, A. Hepatitis B Virus Disrupts Mitochondrial Dynamics: Induces Fission and Mitophagy to Attenuate Apoptosis. PLoS Pathog. 2013, 9, e1003722.
  6. Kim, S.-J.; Syed, G.H.; Khan, M.; Chiu, W.-W.; Sohail, M.A.; Gish, R.G.; Siddiqui, A. Hepatitis C Virus Triggers Mitochondrial Fission and Attenuates Apoptosis to Promote Viral Persistence. Proc. Natl. Acad. Sci. USA 2014, 111, 6413–6418.
  7. Gatti, P.; Ilamathi, H.S.; Todkar, K.; Germain, M. Mitochondria Targeted Viral Replication and Survival Strategies-Prospective on SARS-CoV-2. Front. Pharmacol. 2020, 11, 578599.
  8. Shang, C.; Liu, Z.; Zhu, Y.; Lu, J.; Ge, C.; Zhang, C.; Li, N.; Jin, N.; Li, Y.; Tian, M.; et al. SARS-CoV-2 Causes Mitochondrial Dysfunction and Mitophagy Impairment. Front. Microbiol. 2021, 12, 780768.
  9. Wu, K.E.; Fazal, F.M.; Parker, K.R.; Zou, J.; Chang, H.Y. RNA-GPS Predicts SARS-CoV-2 RNA Residency to Host Mitochondria and Nucleolus. Cell Syst. 2020, 11, 102–108.e3.
  10. Pliss, A.; Kuzmin, A.N.; Prasad, P.N.; Mahajan, S.D. Mitochondrial Dysfunction: A Prelude to Neuropathogenesis of SARS-CoV-2. ACS Chem. Neurosci. 2022, 13, 308–312.
  11. Cortese, M.; Lee, J.-Y.; Cerikan, B.; Neufeldt, C.J.; Oorschot, V.M.J.; Köhrer, S.; Hennies, J.; Schieber, N.L.; Ronchi, P.; Mizzon, G.; et al. Integrative Imaging Reveals SARS-CoV-2-Induced Reshaping of Subcellular Morphologies. Cell Host Microbe 2020, 28, 853–866.e5.
  12. Santos, A.F.; Póvoa, P.; Paixão, P.; Mendonça, A.; Taborda-Barata, L. Changes in Glycolytic Pathway in SARS-COV 2 Infection and Their Importance in Understanding the Severity of COVID-19. Front. Chem. 2021, 9, 685196.
  13. Ma, K.; Wu, H.; Li, P.; Li, B. LC3-II May Mediate ATR-Induced Mitophagy in Dopaminergic Neurons through SQSTM1/p62 Pathway. Acta Biochim. Biophys. Sin. 2018, 50, 1047–1061.
  14. Wang, S.; Zhao, Z.; Feng, X.; Cheng, Z.; Xiong, Z.; Wang, T.; Lin, J.; Zhang, M.; Hu, J.; Fan, Y.; et al. Melatonin Activates Parkin Translocation and Rescues the Impaired Mitophagy Activity of Diabetic Cardiomyopathy through Mst1 Inhibition. J. Cell. Mol. Med. 2018, 22, 5132–5144.
  15. da Costa, L.S.; Outlioua, A.; Anginot, A.; Akarid, K.; Arnoult, D. RNA Viruses Promote Activation of the NLRP3 Inflammasome through Cytopathogenic Effect-Induced Potassium Efflux. Cell Death Dis. 2019, 10, 346.
  16. Muñoz-Planillo, R.; Kuffa, P.; Martínez-Colón, G.; Smith, B.L.; Rajendiran, T.M.; Núñez, G. K+ Efflux Is the Common Trigger of NLRP3 Inflammasome Activation by Bacterial Toxins and Particulate Matter. Immunity 2013, 38, 1142–1153.
  17. Ichinohe, T.; Pang, I.K.; Iwasaki, A. Influenza Virus Activates Inflammasomes via Its Intracellular M2 Ion Channel. Nat. Immunol. 2010, 11, 404–410.
  18. de Zoete, M.R.; Palm, N.W.; Zhu, S.; Flavell, R.A. Inflammasomes. Cold Spring Harb. Perspect. Biol. 2014, 6, a016287.
  19. DeDiego, M.L.; Nieto-Torres, J.L.; Jimenez-Guardeño, J.M.; Regla-Nava, J.A.; Castaño-Rodriguez, C.; Fernandez-Delgado, R.; Usera, F.; Enjuanes, L. Coronavirus Virulence Genes with Main Focus on SARS-CoV Envelope Gene. Virus Res. 2014, 194, 124–137.
  20. Farag, N.S.; Breitinger, U.; Breitinger, H.G.; El Azizi, M.A. Viroporins and Inflammasomes: A Key to Understand Virus-Induced Inflammation. Int. J. Biochem. Cell Biol. 2020, 122, 105738.
  21. McClenaghan, C.; Hanson, A.; Lee, S.-J.; Nichols, C.G. Coronavirus Proteins as Ion Channels: Current and Potential Research. Front. Immunol. 2020, 11, 573339.
  22. Arya, R.; Kumari, S.; Pandey, B.; Mistry, H.; Bihani, S.C.; Das, A.; Prashar, V.; Gupta, G.D.; Panicker, L.; Kumar, M. Structural Insights into SARS-CoV-2 Proteins. J. Mol. Biol. 2021, 433, 166725.
  23. Cao, Y.; Yang, R.; Lee, I.; Zhang, W.; Sun, J.; Wang, W.; Meng, X. Characterization of the SARS-CoV-2 E Protein: Sequence, Structure, Viroporin, and Inhibitors. Protein Sci. 2021, 30, 1114–1130.
  24. Mandala, V.S.; McKay, M.J.; Shcherbakov, A.A.; Dregni, A.J.; Kolocouris, A.; Hong, M. Structure and Drug Binding of the SARS-CoV-2 Envelope Protein Transmembrane Domain in Lipid Bilayers. Nat. Struct. Mol. Biol. 2020, 27, 1202–1208.
  25. Breitinger, U.; Ali, N.K.M.; Sticht, H.; Breitinger, H.-G. Inhibition of SARS CoV Envelope Protein by Flavonoids and Classical Viroporin Inhibitors. Front. Microbiol. 2021, 12, 692423.
  26. Rizwan, T.; Kothidar, A.; Meghwani, H.; Sharma, V.; Shobhawat, R.; Saini, R.; Vaishnav, H.K.; Singh, V.; Pratap, M.; Sihag, H.; et al. Comparative Analysis of SARS-CoV-2 Envelope Viroporin Mutations from COVID-19 Deceased and Surviving Patients Revealed Implications on Its Ion-Channel Activities and Correlation with Patient Mortality. J. Biomol. Struct. Dyn. 2021, 1–16.
  27. Cao, Y.; Yang, R.; Wang, W.; Lee, I.; Zhang, R.; Zhang, W.; Sun, J.; Xu, B.; Meng, X. Computational Study of the Ion and Water Permeation and Transport Mechanisms of the SARS-CoV-2 Pentameric E Protein Channel. Front. Mol. Biosci 2020, 7, 565797.
  28. Liao, Y.; Yuan, Q.; Torres, J.; Tam, J.P.; Liu, D.X. Biochemical and Functional Characterization of the Membrane Association and Membrane Permeabilizing Activity of the Severe Acute Respiratory Syndrome Coronavirus Envelope Protein. Virology 2006, 349, 264–275.
  29. Pervushin, K.; Tan, E.; Parthasarathy, K.; Lin, X.; Jiang, F.L.; Yu, D.; Vararattanavech, A.; Soong, T.W.; Liu, D.X.; Torres, J. Structure and Inhibition of the SARS Coronavirus Envelope Protein Ion Channel. PLoS Pathog. 2009, 5, e1000511.
  30. Mehregan, A.; Pérez-Conesa, S.; Zhuang, Y.; Elbahnsi, A.; Pasini, D.; Lindahl, E.; Howard, R.J.; Ulens, C.; Delemotte, L. Probing Effects of the SARS-CoV-2 E Protein on Membrane Curvature and Intracellular Calcium. Biochim. Biophys. Acta Biomembr. 2022, 1864, 183994.
  31. Nardacci, R.; Colavita, F.; Castilletti, C.; Lapa, D.; Matusali, G.; Meschi, S.; Del Nonno, F.; Colombo, D.; Capobianchi, M.R.; Zumla, A.; et al. Evidences for Lipid Involvement in SARS-CoV-2 Cytopathogenesis. Cell Death Dis. 2021, 12, 263.
  32. Wang, P.; Luo, R.; Zhang, M.; Wang, Y.; Song, T.; Tao, T.; Li, Z.; Jin, L.; Zheng, H.; Chen, W.; et al. A Cross-Talk between Epithelium and Endothelium Mediates Human Alveolar–capillary Injury during SARS-CoV-2 Infection. Cell Death Dis. 2020, 11, 1–17.
  33. Mannella, C.A.; Pfeiffer, D.R.; Bradshaw, P.C.; Moraru, I.I.; Slepchenko, B.; Loew, L.M.; Hsieh, C.E.; Buttle, K.; Marko, M. Topology of the Mitochondrial Inner Membrane: Dynamics and Bioenergetic Implications. IUBMB Life 2001, 52, 93–100.
  34. Zorova, L.D.; Popkov, V.A.; Plotnikov, E.Y.; Silachev, D.N.; Pevzner, I.B.; Jankauskas, S.S.; Babenko, V.A.; Zorov, S.D.; Balakireva, A.V.; Juhaszova, M.; et al. Mitochondrial Membrane Potential. Anal. Biochem. 2018, 552, 50–59.
  35. Zick, M.; Rabl, R.; Reichert, A.S. Cristae Formation-Linking Ultrastructure and Function of Mitochondria. Biochim. Biophys. Acta 2009, 1793, 5–19.
  36. Gilkerson, R.W.; Selker, J.M.L.; Capaldi, R.A. The Cristal Membrane of Mitochondria Is the Principal Site of Oxidative Phosphorylation. FEBS Lett. 2003, 546, 355–358.
  37. Mitchell, P. Coupling of Phosphorylation to Electron and Hydrogen Transfer by a Chemi-Osmotic Type of Mechanism. Nature 1961, 191, 144–148.
  38. Wolf, D.M.; Segawa, M.; Kondadi, A.K.; Anand, R.; Bailey, S.T.; Reichert, A.S.; van der Bliek, A.M.; Shackelford, D.B.; Liesa, M.; Shirihai, O.S. Individual Cristae within the Same Mitochondrion Display Different Membrane Potentials and Are Functionally Independent. EMBO J. 2019, 38, e101056.
  39. Rieger, B.; Junge, W.; Busch, K.B. Lateral pH Gradient between OXPHOS Complex IV and F(0)F(1) ATP-Synthase in Folded Mitochondrial Membranes. Nat. Commun. 2014, 5, 3103.
  40. Garcia, G.C.; Bartol, T.M.; Phan, S.; Bushong, E.A.; Perkins, G.; Sejnowski, T.J.; Ellisman, M.H.; Skupin, A. Mitochondrial Morphology Provides a Mechanism for Energy Buffering at Synapses. Sci. Rep. 2019, 9, 18306.
  41. Scott, I.D.; Nicholls, D.G. Energy Transduction in Intact Synaptosomes. Influence of Plasma-Membrane Depolarization on the Respiration and Membrane Potential of Internal Mitochondria Determined in Situ. Biochem. J. 1980, 186, 21–33.
  42. Mannella, C.A. Consequences of Folding the Mitochondrial Inner Membrane. Front. Physiol. 2020, 11, 536.
  43. Gottlieb, E.; Armour, S.M.; Harris, M.H.; Thompson, C.B. Mitochondrial Membrane Potential Regulates Matrix Configuration and Cytochrome c Release during Apoptosis. Cell Death Differ. 2003, 10, 709–717.
  44. Rasola, A.; Bernardi, P. Mitochondrial Permeability Transition in Ca(2+)-Dependent Apoptosis and Necrosis. Cell Calcium 2011, 50, 222–233.
  45. Liesa, M. Why Does a Mitochondrion Need Its Individual Cristae to Be Functionally Autonomous? Mol. Cell Oncol. 2020, 7, 1705119.
  46. Twig, G.; Elorza, A.; Molina, A.J.A.; Mohamed, H.; Wikstrom, J.D.; Walzer, G.; Stiles, L.; Haigh, S.E.; Katz, S.; Las, G.; et al. Fission and Selective Fusion Govern Mitochondrial Segregation and Elimination by Autophagy. EMBO J. 2008, 27, 433–446.
  47. Twig, G.; Shirihai, O.S. The Interplay between Mitochondrial Dynamics and Mitophagy. Antioxid. Redox Signal. 2011, 14, 1939–1951.
  48. Medini, H.; Zirman, A.; Mishmar, D. Immune System Cells from COVID-19 Patients Display Compromised Mitochondrial-Nuclear Expression Co-Regulation and Rewiring toward Glycolysis. iScience 2021, 24, 103471.
  49. Wu, M.; Neilson, A.; Swift, A.L.; Moran, R.; Tamagnine, J.; Parslow, D.; Armistead, S.; Lemire, K.; Orrell, J.; Teich, J.; et al. Multiparameter Metabolic Analysis Reveals a Close Link between Attenuated Mitochondrial Bioenergetic Function and Enhanced Glycolysis Dependency in Human Tumor Cells. Am. J. Physiol. Cell Physiol. 2007, 292, C125–C136.
  50. Codo, A.C.; Davanzo, G.G.; de Brito Monteiro, L.; de Souza, G.F.; Muraro, S.P.; Virgilio-da-Silva, J.V.; Prodonoff, J.S.; Carregari, V.C.; de Biagi Junior, C.A.O.; Crunfli, F.; et al. Elevated Glucose Levels Favor SARS-CoV-2 Infection and Monocyte Response through a HIF-1α/Glycolysis-Dependent Axis. Cell Metab. 2020, 32, 437–446.e5.
  51. Aklima, J.; Onojima, T.; Kimura, S.; Umiuchi, K.; Shibata, T.; Kuraoka, Y.; Oie, Y.; Suganuma, Y.; Ohta, Y. Effects of Matrix pH on Spontaneous Transient Depolarization and Reactive Oxygen Species Production in Mitochondria. Front. Cell Dev. Biol. 2021, 9, 692776.
  52. Moreno Davila, H. Molecular and Functional Diversity of Voltage-Gated Calcium Channels. Ann. N. Y. Acad. Sci. 1999, 868, 102–117.
  53. Pitt, G.S.; Matsui, M.; Cao, C. Voltage-Gated Calcium Channels in Nonexcitable Tissues. Annu. Rev. Physiol. 2021, 83, 183–203.
  54. Catterall, W.A. Voltage-Gated Calcium Channels. Cold Spring Harb. Perspect. Biol. 2011, 3, a003947.
  55. Agirre, A.; Barco, A.; Carrasco, L.; Nieva, J.L. Viroporin-Mediated Membrane Permeabilization. Pore Formation by Nonstructural Poliovirus 2B Protein. J. Biol. Chem. 2002, 277, 40434–40441.
  56. Firth, A.E.; Chung, B.Y.; Fleeton, M.N.; Atkins, J.F. Discovery of Frameshifting in Alphavirus 6K Resolves a 20-Year Enigma. Virol. J. 2008, 5, 108.
  57. González, M.E. Vpu Protein: The Viroporin Encoded by HIV-1. Viruses 2015, 7, 4352–4368.
  58. To, J.; Torres, J. Viroporins in the Influenza Virus. Cells 2019, 8.
  59. Liao, Y.; Lescar, J.; Tam, J.P.; Liu, D.X. Expression of SARS-Coronavirus Envelope Protein in Escherichia Coli Cells Alters Membrane Permeability. Biochem. Biophys. Res. Commun. 2004, 325, 374–380.
  60. Landstrom, A.P.; Dobrev, D.; Wehrens, X.H.T. Calcium Signaling and Cardiac Arrhythmias. Circ. Res. 2017, 120, 1969–1993.
  61. Larsen, H.E.; Bardsley, E.N.; Lefkimmiatis, K.; Paterson, D.J. Dysregulation of Neuronal Ca2+ Channel Linked to Heightened Sympathetic Phenotype in Prohypertensive States. J. Neurosci. 2016, 36, 8562–8573.
  62. Jamal, S.M.; Landers, D.B.; Hollenberg, S.M.; Turi, Z.G.; Glotzer, T.V.; Tancredi, J.; Parrillo, J.E. Prospective Evaluation of Autonomic Dysfunction in Post-Acute Sequela of COVID-19. J. Am. Coll. Cardiol. 2022.
  63. Dani, M.; Dirksen, A.; Taraborrelli, P.; Torocastro, M.; Panagopoulos, D.; Sutton, R.; Lim, P.B. Autonomic Dysfunction in “Long COVID”: Rationale, Physiology and Management Strategies. Clin. Med. 2021, 21, e63–e67.
  64. Papadopoulou, M.; Bakola, E.; Papapostolou, A.; Stefanou, M.-I.; Gaga, M.; Zouvelou, V.; Michopoulos, I.; Tsivgoulis, G. Autonomic Dysfunction in Long-COVID Syndrome: A Neurophysiological and Neurosonology Study. J. Neurol. 2022, 1–2.
  65. Raj, S.R.; Arnold, A.C.; Barboi, A.; Claydon, V.E.; Limberg, J.K.; Lucci, V.-E.M.; Numan, M.; Peltier, A.; Snapper, H.; Vernino, S.; et al. Long-COVID Postural Tachycardia Syndrome: An American Autonomic Society Statement. Clin. Auton. Res. 2021, 31, 365–368.
  66. Bisaccia, G.; Ricci, F.; Recce, V.; Serio, A.; Iannetti, G.; Chahal, A.A.; Ståhlberg, M.; Khanji, M.Y.; Fedorowski, A.; Gallina, S. Post-Acute Sequelae of COVID-19 and Cardiovascular Autonomic Dysfunction: What Do We Know? J. Cardiovasc. Dev. Dis 2021, 8, 156.
  67. Chen, X.; Cao, R.; Zhong, W. Host Calcium Channels and Pumps in Viral Infections. Cells 2019, 9, 94.
  68. Hyser, J.M.; Estes, M.K. Pathophysiological Consequences of Calcium-Conducting Viroporins. Annu Rev. Virol. 2015, 2, 473–496.
  69. Berktaş, B.M.; Gökçek, A.; Hoca, N.T.; Koyuncu, A. COVID-19 Illness and Treatment Decrease Bone Mineral Density of Surviving Hospitalized Patients. Eur. Rev. Med. Pharmacol. Sci. 2022, 26, 3046–3056.
  70. Mandala, V.S.; Loftis, A.R.; Shcherbakov, A.A.; Pentelute, B.L.; Hong, M. Atomic Structures of Closed and Open Influenza B M2 Proton Channel Reveal the Conduction Mechanism. Nat. Struct. Mol. Biol. 2020, 27, 160–167.
  71. Gargan, S.; Stevenson, N.J. Unravelling the Immunomodulatory Effects of Viral Ion Channels, towards the Treatment of Disease. Viruses 2021, 13, 2165.
  72. Bohmwald, K.; Gálvez, N.M.S.; Andrade, C.A.; Mora, V.P.; Muñoz, J.T.; González, P.A.; Riedel, C.A.; Kalergis, A.M. Modulation of Adaptive Immunity and Viral Infections by Ion Channels. Front. Physiol. 2021, 12, 736681.
  73. Feske, S.; Wulff, H.; Skolnik, E.Y. Ion Channels in Innate and Adaptive Immunity. Annu. Rev. Immunol. 2015, 33, 291–353.
  74. Kaivola, J.; Nyman, T.A.; Matikainen, S. Inflammasomes and SARS-CoV-2 Infection. Viruses 2021, 13, 2513.
  75. Campbell, G.R.; To, R.K.; Hanna, J.; Spector, S.A. SARS-CoV-2, SARS-CoV-1, and HIV-1 Derived ssRNA Sequences Activate the NLRP3 Inflammasome in Human Macrophages through a Non-Classical Pathway. iScience 2021, 24, 102295.
  76. Causton, H.C. SARS-CoV2 Infection and the Importance of Potassium Balance. Front. Med. 2021, 8, 744697.
  77. Xu, H.; Akinyemi, I.A.; Chitre, S.A.; Loeb, J.C.; Lednicky, J.A.; McIntosh, M.T.; Bhaduri-McIntosh, S. SARS-CoV-2 Viroporin Encoded by ORF3a Triggers the NLRP3 Inflammatory Pathway. Virology 2022, 568, 13–22.
  78. Wang, J.; Zhang, K.; Goyal, P.; Grewer, C. Mechanism and Potential Sites of Potassium Interaction with Glutamate Transporters. J. Gen. Physiol. 2020, 152, e202012577.
  79. Kozlov, A.G.; Cheng, X.; Zhang, H.; Shinn, M.K.; Weiland, E.; Nguyen, B.; Shkel, I.A.; Zytkiewicz, E.; Finkelstein, I.J.; Record, M.T., Jr.; et al. How Glutamate Promotes Liquid-Liquid Phase Separation and DNA Binding Cooperativity of E. Coli SSB Protein. J. Mol. Biol. 2022, 434, 167562.
  80. Rimmele, T.S.; Rocher, A.-B.; Wellbourne-Wood, J.; Chatton, J.-Y. Control of Glutamate Transport by Extracellular Potassium: Basis for a Negative Feedback on Synaptic Transmission. Cereb. Cortex 2017, 27, 3272–3283.
  81. Bharadwaj, S.; Singh, M.; Kirtipal, N.; Kang, S.G. SARS-CoV-2 and Glutamine: SARS-CoV-2 Triggered Pathogenesis via Metabolic Reprograming of Glutamine in Host Cells. Front. Mol. Biosci 2020, 7, 627842.
  82. Wang, J.; Yang, G.; Wang, X.; Wen, Z.; Shuai, L.; Luo, J.; Wang, C.; Sun, Z.; Liu, R.; Ge, J.; et al. SARS-CoV-2 Uses Metabotropic Glutamate Receptor Subtype 2 as an Internalization Factor to Infect Cells. Cell Discov. 2021, 7, 119.
  83. Díaz-Resendiz, K.J.G.; Benitez-Trinidad, A.B.; Covantes-Rosales, C.E.; Toledo-Ibarra, G.A.; Ortiz-Lazareno, P.C.; Girón-Pérez, D.A.; Bueno-Durán, A.Y.; Pérez-Díaz, D.A.; Barcelos-García, R.G.; Girón-Pérez, M.I. Loss of Mitochondrial Membrane Potential (ΔΨm ) in Leucocytes as Post-COVID-19 Sequelae. J. Leukoc. Biol. 2022, 112, 23–29.
  84. Fitzgerald-Bocarsly, P. Human Natural Interferon-Alpha Producing Cells. Pharmacol. Ther. 1993, 60, 39–62.
  85. Decker, P. Neutrophils and Interferon-α-Producing Cells: Who Produces Interferon in Lupus? Arthritis Res. Ther. 2011, 13, 118.
  86. Wong, R.S.M.; Wu, A.; To, K.F.; Lee, N.; Lam, C.W.K.; Wong, C.K.; Chan, P.K.S.; Ng, M.H.L.; Yu, L.M.; Hui, D.S.; et al. Haematological Manifestations in Patients with Severe Acute Respiratory Syndrome: Retrospective Analysis. BMJ 2003, 326, 1358–1362.
  87. Zou, Z.-Y.; Ren, D.; Chen, R.-L.; Yu, B.-J.; Liu, Y.; Huang, J.-J.; Yang, Z.-J.; Zhou, Z.-P.; Feng, Y.-W.; Wu, M. Persistent Lymphopenia after Diagnosis of COVID-19 Predicts Acute Respiratory Distress Syndrome: A Retrospective Cohort Study. Eur. J. Inflam. 2021, 19, 20587392211036825.
  88. Ghizlane, E.A.; Manal, M.; Abderrahim, E.K.; Abdelilah, E.; Mohammed, M.; Rajae, A.; Amine, B.M.; Houssam, B.; Naima, A.; Brahim, H. Lymphopenia in Covid-19: A Single Center Retrospective Study of 589 Cases. Ann. Med. Surg. 2021, 69, 102816.
  89. Zheng, M.; Gao, Y.; Wang, G.; Song, G.; Liu, S.; Sun, D.; Xu, Y.; Tian, Z. Functional Exhaustion of Antiviral Lymphocytes in COVID-19 Patients. Cell. Mol. Immunol. 2020, 17, 533–535.
  90. Zhao, Q.; Meng, M.; Kumar, R.; Wu, Y.; Huang, J.; Deng, Y.; Weng, Z.; Yang, L. Lymphopenia Is Associated with Severe Coronavirus Disease 2019 (COVID-19) Infections: A Systemic Review and Meta-Analysis. Int. J. Infect. Dis. 2020, 96, 131–135.
  91. Tan, L.; Wang, Q.; Zhang, D.; Ding, J.; Huang, Q.; Tang, Y.-Q.; Wang, Q.; Miao, H. Lymphopenia Predicts Disease Severity of COVID-19: A Descriptive and Predictive Study. Signal Transduct. Target. Ther. 2020, 5, 33.
  92. Liu, J.; Li, S.; Liu, J.; Liang, B.; Wang, X.; Wang, H.; Li, W.; Tong, Q.; Yi, J.; Zhao, L.; et al. Longitudinal Characteristics of Lymphocyte Responses and Cytokine Profiles in the Peripheral Blood of SARS-CoV-2 Infected Patients. EBioMedicine 2020, 55, 102763.
  93. Ledderose, C.; Bao, Y.; Lidicky, M.; Zipperle, J.; Li, L.; Strasser, K.; Shapiro, N.I.; Junger, W.G. Mitochondria Are Gate-Keepers of T Cell Function by Producing the ATP That Drives Purinergic Signaling. J. Biol. Chem. 2014, 289, 25936–25945.
  94. Desdín-Micó, G.; Soto-Heredero, G.; Mittelbrunn, M. Mitochondrial Activity in T Cells. Mitochondrion 2018, 41, 51–57.
  95. Feske, S.; Giltnane, J.; Dolmetsch, R.; Staudt, L.M.; Rao, A. Gene Regulation Mediated by Calcium Signals in T Lymphocytes. Nat. Immunol. 2001, 2, 316–324.
  96. Campello, S.; Lacalle, R.A.; Bettella, M.; Mañes, S.; Scorrano, L.; Viola, A. Orchestration of Lymphocyte Chemotaxis by Mitochondrial Dynamics. J. Exp. Med. 2006, 203, 2879–2886.
  97. Zhang, K.; Li, H.; Song, Z. Membrane Depolarization Activates the Mitochondrial Protease OMA1 by Stimulating Self-Cleavage. EMBO Rep. 2014, 15, 576–585.
  98. Jahangir, A.; Ozcan, C.; Holmuhamedov, E.L.; Terzic, A. Increased Calcium Vulnerability of Senescent Cardiac Mitochondria: Protective Role for a Mitochondrial Potassium Channel Opener. Mech. Ageing Dev. 2001, 122, 1073–1086.
  99. Glitsch, M.D.; Bakowski, D.; Parekh, A.B. Store-Operated Ca2+ Entry Depends on Mitochondrial Ca2+ Uptake. EMBO J. 2002, 21, 6744–6754.
  100. Santos, J.H.; Hunakova, L.U.; Chen, Y.; Bortner, C.; Van Houten, B. Cell Sorting Experiments Link Persistent Mitochondrial DNA Damage with Loss of Mitochondrial Membrane Potential and Apoptotic Cell Death. J. Biol. Chem. 2003, 278, 1728–1734.
  101. Peña, C.; Rincon, J.; Pedreanez, A.; Viera, N.; Mosquera, J. Chemotactic Effect of Melatonin on Leukocytes. J. Pineal Res. 2007, 43, 263–269.
  102. Hu, L.; Zhang, S.; Wen, H.; Liu, T.; Cai, J.; Du, D.; Zhu, D.; Chen, F.; Xia, C. Melatonin Decreases M1 Polarization via Attenuating Mitochondrial Oxidative Damage Depending on UCP2 Pathway in Prorenin-Treated Microglia. PLoS ONE 2019, 14, e0212138.
  103. Liu, Y.-J.; Ji, D.-M.; Liu, Z.-B.; Wang, T.-J.; Xie, F.-F.; Zhang, Z.-G.; Wei, Z.-L.; Zhou, P.; Cao, Y.-X. Melatonin Maintains Mitochondrial Membrane Potential and Decreases Excessive Intracellular Ca2+ Levels in Immature Human Oocytes. Life Sci. 2019, 235, 116810.
  104. Lançoni, R.; Celeghini, E.C.C.; Alves, M.B.R.; Lemes, K.M.; Gonella-Diaza, A.M.; Oliveira, L.Z.; de Arruda, R.P. Melatonin Added to Cryopreservation Extenders Improves the Mitochondrial Membrane Potential of Postthawed Equine Sperm. J. Equine Vet. Sci. 2018, 69, 78–83.
  105. Kumari, S.; Dash, D. Melatonin Elevates Intracellular Free Calcium in Human Platelets by Inositol 1,4,5-Trisphosphate Independent Mechanism. FEBS Lett. 2011, 585, 2345–2351.
  106. Pieri, C.; Recchioni, R.; Moroni, F.; Marcheselli, F.; Marra, M.; Marinoni, S.; Di Primio, R. Melatonin Regulates the Respiratory Burst of Human Neutrophils and Their Depolarization. J. Pineal Res. 1998, 24, 43–49.
  107. Fischer, T.W.; Zmijewski, M.A.; Wortsman, J.; Slominski, A. Melatonin Maintains Mitochondrial Membrane Potential and Attenuates Activation of Initiator (casp-9) and Effector Caspases (casp-3/casp-7) and PARP in UVR-Exposed HaCaT Keratinocytes. J. Pineal Res. 2008, 44, 397–407.
  108. NavaneethaKrishnan, S.; Rosales, J.L.; Lee, K.-Y. mPTP Opening Caused by Cdk5 Loss Is due to Increased Mitochondrial Ca2+ Uptake. Oncogene 2020, 39, 2797–2806.
  109. Park, J.; Lee, J.; Choi, C. Mitochondrial Network Determines Intracellular ROS Dynamics and Sensitivity to Oxidative Stress through Switching Inter-Mitochondrial Messengers. PLoS ONE 2011, 6, e23211.
  110. Niki, E. Lipid Peroxidation: Physiological Levels and Dual Biological Effects. Free Radic. Biol. Med. 2009, 47, 469–484.
  111. Ayala, A.; Muñoz, M.F.; Argüelles, S. Lipid Peroxidation: Production, Metabolism, and Signaling Mechanisms of Malondialdehyde and 4-Hydroxy-2-Nonenal. Oxid. Med. Cell. Longev. 2014, 2014, 360438.
  112. Žarković, N.; Orehovec, B.; Milković, L.; Baršić, B.; Tatzber, F.; Wonisch, W.; Tarle, M.; Kmet, M.; Mataić, A.; Jakovčević, A.; et al. Preliminary Findings on the Association of the Lipid Peroxidation Product 4-Hydroxynonenal with the Lethal Outcome of Aggressive COVID-19. Antioxidants 2021, 10, 1341.
  113. Loh, D.; Reiter, R.J. Melatonin: Regulation of Biomolecular Condensates in Neurodegenerative Disorders. Antioxidants 2021, 10, 1483.
  114. Reiter, R.J.; Tan, D.-X.; Galano, A. Melatonin Reduces Lipid Peroxidation and Membrane Viscosity. Front. Physiol. 2014, 5, 377.
  115. García, J.J.; López-Pingarrón, L.; Almeida-Souza, P.; Tres, A.; Escudero, P.; García-Gil, F.A.; Tan, D.-X.; Reiter, R.J.; Ramírez, J.M.; Bernal-Pérez, M. Protective Effects of Melatonin in Reducing Oxidative Stress and in Preserving the Fluidity of Biological Membranes: A Review. J. Pineal Res. 2014, 56, 225–237.
  116. Petrosillo, G.; Moro, N.; Ruggiero, F.M.; Paradies, G. Melatonin Inhibits Cardiolipin Peroxidation in Mitochondria and Prevents the Mitochondrial Permeability Transition and Cytochrome c Release. Free Radic. Biol. Med. 2009, 47, 969–974.
  117. Livrea, M.A.; Tesoriere, L.; D’Arpa, D.; Morreale, M. Reaction of Melatonin with Lipoperoxyl Radicals in Phospholipid Bilayers. Free Radic. Biol. Med. 1997, 23, 706–711.
  118. Galano, A.; Reiter, R.J. Melatonin and Its Metabolites vs Oxidative Stress: From Individual Actions to Collective Protection. J. Pineal Res. 2018, 65, e12514.
  119. Tan, D.X.; Manchester, L.C.; Reiter, R.J.; Plummer, B.F.; Limson, J.; Weintraub, S.T.; Qi, W. Melatonin Directly Scavenges Hydrogen Peroxide: A Potentially New Metabolic Pathway of Melatonin Biotransformation. Free Radic. Biol. Med. 2000, 29, 1177–1185.
  120. Fischer, T.W.; Scholz, G.; Knöll, B.; Hipler, U.C.; Elsner, P. Melatonin Reduces UV-Induced Reactive Oxygen Species in a Dose-Dependent Manner in IL-3-Stimulated Leukocytes. J. Pineal Res. 2001, 31, 39–45.
  121. Reiter, R.J. Melatonin: Lowering the High Price of Free Radicals. News Physiol. Sci. 2000, 15, 246–250.
  122. 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.
  123. Vanecek, J.; Klein, D.C. Sodium-Dependent Effects of Melatonin on Membrane Potential of Neonatal Rat Pituitary Cells. Endocrinology 1992, 131, 939–946.
  124. Bortner, C.D.; Gomez-Angelats, M.; Cidlowski, J.A. Plasma Membrane Depolarization without Repolarization Is an Early Molecular Event in Anti-Fas-Induced Apoptosis. J. Biol. Chem. 2001, 276, 4304–4314.
  125. Ching, A.C.; Hughes, M.R.; Poon, A.M.; Pang, S.F. Melatonin Receptors and Melatonin Inhibition of Duck Salt Gland Secretion. Gen. Comp. Endocrinol. 1999, 116, 229–240.
  126. Hughes, M.R.; Kitamura, N.; Bennett, D.C.; Gray, D.A.; Sharp, P.J.; Poon, A.M.S. Effect of Melatonin on Salt Gland and Kidney Function of Gulls, Larus Glaucescens. Gen. Comp. Endocrinol. 2007, 151, 300–307.
  127. Farouk, S.; Al-Huqail, A.A. Sustainable Biochar And/or Melatonin Improve Salinity Tolerance in Borage Plants by Modulating Osmotic Adjustment, Antioxidants, and Ion Homeostasis. Plants 2022, 11, 765.
  128. Jiang, C.; Cui, Q.; Feng, K.; Xu, D.; Li, C.; Zheng, Q. Melatonin Improves Antioxidant Capacity and Ion Homeostasis and Enhances Salt Tolerance in Maize Seedlings. Acta Physiol. Plant 2016, 38, 82.
  129. Li, C.; Wang, P.; Wei, Z.; Liang, D.; Liu, C.; Yin, L.; Jia, D.; Fu, M.; Ma, F. The Mitigation Effects of Exogenous Melatonin on Salinity-Induced Stress in Malus Hupehensis. J. Pineal Res. 2012, 53, 298–306.
  130. Chakravarty, S.; Rizvi, S.I. Circadian Modulation of Sodium-Potassium ATPase and Sodium—Proton Exchanger in Human Erythrocytes: In Vitro Effect of Melatonin. Cell. Mol. Biol. 2011, 57, 80–86.
  131. Loh, D.; Reiter, R.J. Melatonin: Regulation of Prion Protein Phase Separation in Cancer Multidrug Resistance. Molecules 2022, 27, 705.
  132. Morth, J.P.; Pedersen, B.P.; Buch-Pedersen, M.J.; Andersen, J.P.; Vilsen, B.; Palmgren, M.G.; Nissen, P. A Structural Overview of the Plasma Membrane Na+,K+-ATPase and H+-ATPase Ion Pumps. Nat. Rev. Mol. Cell Biol. 2011, 12, 60–70.
  133. Noel, J.; Roux, D.; Pouysségur, J. Differential Localization of Na+/H+ Exchanger Isoforms (NHE1 and NHE3) in Polarized Epithelial Cell Lines. J. Cell Sci. 1996, 109 Pt 5, 929–939.
  134. Clausen, M.V.; Hilbers, F.; Poulsen, H. The Structure and Function of the Na,K-ATPase Isoforms in Health and Disease. Front. Physiol. 2017, 8, 371.
  135. Grinstein, S.; Rotin, D.; Mason, M.J. Na+/H+ Exchange and Growth Factor-Induced Cytosolic pH Changes. Role in Cellular Proliferation. Biochim. Biophys. Acta 1989, 988, 73–97.
  136. Cha, C.Y.; Oka, C.; Earm, Y.E.; Wakabayashi, S.; Noma, A. A Model of Na+/H+ Exchanger and Its Central Role in Regulation of pH and Na+ in Cardiac Myocytes. Biophys. J. 2009, 97, 2674–2683.
  137. Alberti, S.; Gladfelter, A.; Mittag, T. Considerations and Challenges in Studying Liquid-Liquid Phase Separation and Biomolecular Condensates. Cell 2019, 176, 419–434.
  138. Królicka, A.L.; Kruczkowska, A.; Krajewska, M.; Kusztal, M.A. Hyponatremia in Infectious Diseases-A Literature Review. Int. J. Environ. Res. Public Health 2020, 17.
  139. Machado, R.R.G.; Glaser, T.; Araujo, D.B.; Petiz, L.L.; Oliveira, D.B.L.; Durigon, G.S.; Leal, A.L.; Pinho, J.R.R.; Ferreira, L.C.S.; Ulrich, H.; et al. Inhibition of Severe Acute Respiratory Syndrome Coronavirus 2 Replication by Hypertonic Saline Solution in Lung and Kidney Epithelial Cells. ACS Pharmacol. Transl. Sci. 2021, 4, 1514–1527.
  140. Kühlbrandt, W. Biology, Structure and Mechanism of P-Type ATPases. Nat. Rev. Mol. Cell Biol. 2004, 5, 282–295.
  141. Dalskov, S.-M.; Immerdal, L.; Niels-Christiansen, L.-L.; Hansen, G.H.; Schousboe, A.; Danielsen, E.M. Lipid Raft Localization of GABA A Receptor and Na+, K+-ATPase in Discrete Microdomain Clusters in Rat Cerebellar Granule Cells. Neurochem. Int. 2005, 46, 489–499.
  142. Welker, P.; Geist, B.; Frühauf, J.-H.; Salanova, M.; Groneberg, D.A.; Krause, E.; Bachmann, S. Role of Lipid Rafts in Membrane Delivery of Renal Epithelial Na+-K+-ATPase, Thick Ascending Limb. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2007, 292, R1328–R1337.
  143. Fujii, T.; Takahashi, Y.; Itomi, Y.; Fujita, K.; Morii, M.; Tabuchi, Y.; Asano, S.; Tsukada, K.; Takeguchi, N.; Sakai, H. K+-Cl- Cotransporter-3a Up-Regulates Na+,K+-ATPase in Lipid Rafts of Gastric Luminal Parietal Cells. J. Biol. Chem. 2008, 283, 6869–6877.
  144. Pytel, E.; Olszewska-Banaszczyk, M.; Koter-Michalak, M.; Broncel, M. Increased Oxidative Stress and Decreased Membrane Fluidity in Erythrocytes of CAD Patients. Biochem. Cell Biol. 2013, 91, 315–318.
  145. Padmavathi, P.; Reddy, V.D.; Maturu, P.; Varadacharyulu, N. Smoking-Induced Alterations in Platelet Membrane Fluidity and Na(+)/K(+)-ATPase Activity in Chronic Cigarette Smokers. J. Atheroscler. Thromb. 2010, 17, 619–627.
  146. Yelinova, V.; Glazachev, Y.; Khramtsov, V.; Kudryashova, L.; Rykova, V.; Salganik, R. Studies of Human and Rat Blood under Oxidative Stress: Changes in Plasma Thiol Level, Antioxidant Enzyme Activity, Protein Carbonyl Content, and Fluidity of Erythrocyte Membrane. Biochem. Biophys. Res. Commun. 1996, 221, 300–303.
  147. Sutherland, E.; Dixon, B.S.; Leffert, H.L.; Skally, H.; Zaccaro, L.; Simon, F.R. Biochemical Localization of Hepatic Surface-Membrane Na+,K+-ATPase Activity Depends on Membrane Lipid Fluidity. Proc. Natl. Acad. Sci. USA 1988, 85, 8673–8677.
  148. García, J.J.; Reiter, R.J.; Guerrero, J.M.; Escames, G.; Yu, B.P.; Oh, C.S.; Muñoz-Hoyos, A. Melatonin Prevents Changes in Microsomal Membrane Fluidity during Induced Lipid Peroxidation. FEBS Lett. 1997, 408, 297–300.
  149. García, J.J.; Piñol-Ripoll, G.; Martínez-Ballarín, E.; Fuentes-Broto, L.; Miana-Mena, F.J.; Venegas, C.; Caballero, B.; Escames, G.; Coto-Montes, A.; Acuña-Castroviejo, D. Melatonin Reduces Membrane Rigidity and Oxidative Damage in the Brain of SAMP8 Mice. Neurobiol. Aging 2011, 32, 2045–2054.
  150. Ochoa, J.J.; Vílchez, M.J.; Palacios, M.A.; García, J.J.; Reiter, R.J.; Muñoz-Hoyos, A. Melatonin Protects against Lipid Peroxidation and Membrane Rigidity in Erythrocytes from Patients Undergoing Cardiopulmonary Bypass Surgery. J. Pineal Res. 2003, 35, 104–108.
  151. Bolmatov, D.; McClintic, W.T.; Taylor, G.; Stanley, C.B.; Do, C.; Collier, C.P.; Leonenko, Z.; Lavrentovich, M.O.; Katsaras, J. Deciphering Melatonin-Stabilized Phase Separation in Phospholipid Bilayers. Langmuir 2019, 35, 12236–12245.
  152. Santamaria, A.; Batchu, K.C.; Matsarskaia, O.; Prévost, S.F.; Russo, D.; Natali, F.; Seydel, T.; Hoffmann, I.; Laux, V.; Haertlein, M.; et al. Strikingly Different Roles of SARS-CoV-2 Fusion Peptides Uncovered by Neutron Scattering. J. Am. Chem. Soc. 2022, 144, 2968–2979.
  153. Deng, Y.; Angelova, A. Coronavirus-Induced Host Cubic Membranes and Lipid-Related Antiviral Therapies: A Focus on Bioactive Plasmalogens. Front. Cell Dev. Biol. 2021, 9, 630242.
  154. Strauss, M.; Hofhaus, G.; Schröder, R.R.; Kühlbrandt, W. Dimer Ribbons of ATP Synthase Shape the Inner Mitochondrial Membrane. EMBO J. 2008, 27, 1154–1160.
  155. Deng, Y.; Lee, E.L.-H.; Chong, K.; Almsherqi, Z.A. Evaluation of Radical Scavenging System in Amoeba Chaos Carolinense during Nutrient Deprivation. Interface Focus 2017, 7, 20160113.
  156. Mannella, C.A. Structure and Dynamics of the Mitochondrial Inner Membrane Cristae. Biochim. Biophys. Acta 2006, 1763, 542–548.
  157. Dang, M.; Li, Y.; Song, J. ATP Biphasically Modulates LLPS of SARS-CoV-2 Nucleocapsid Protein and Specifically Binds Its RNA-Binding Domain. Biochem. Biophys. Res. Commun. 2021, 541, 50–55.
  158. Zheng, X.; Sun, Z.; Yu, L.; Shi, D.; Zhu, M.; Yao, H.; Li, L. Interactome Analysis of the Nucleocapsid Protein of SARS-CoV-2 Virus. Pathogens 2021, 10, 1155.
  159. Yu, Q.; Guo, M.; Zeng, W.; Zeng, M.; Zhang, X.; Zhang, Y.; Zhang, W.; Jiang, X.; Yu, B. Interactions between NLRP3 Inflammasome and Glycolysis in Macrophages: New Insights into Chronic Inflammation Pathogenesis. Immun. Inflamm. Dis. 2022, 10, e581.
  160. Ajaz, S.; McPhail, M.J.; Singh, K.K.; Mujib, S.; Trovato, F.M.; Napoli, S.; Agarwal, K. Mitochondrial Metabolic Manipulation by SARS-CoV-2 in Peripheral Blood Mononuclear Cells of Patients with COVID-19. Am. J. Physiol. Cell Physiol. 2021, 320, C57–C65.
  161. Mookerjee, S.A.; Gerencser, A.A.; Nicholls, D.G.; Brand, M.D. Quantifying Intracellular Rates of Glycolytic and Oxidative ATP Production and Consumption Using Extracellular Flux Measurements. J. Biol. Chem. 2017, 292, 7189–7207.
  162. Reiter, R.J.; Sharma, R.; Rosales-Corral, S. Anti-Warburg Effect of Melatonin: A Proposed Mechanism to Explain Its Inhibition of Multiple Diseases. Int. J. Mol. Sci. 2021, 22, 764.
  163. Reiter, R.J.; Sharma, R.; Ma, Q.; Rosales-Corral, S.; Acuna-Castroviejo, D.; Escames, G. Inhibition of Mitochondrial Pyruvate Dehydrogenase Kinase: A Proposed Mechanism by Which Melatonin Causes Cancer Cells to Overcome Cytosolic Glycolysis, Reduce Tumor Biomass and Reverse Insensitivity to Chemotherapy. Melatonin Res. 2019, 2, 105–119.
  164. Gray, L.R.; Tompkins, S.C.; Taylor, E.B. Regulation of Pyruvate Metabolism and Human Disease. Cell. Mol. Life Sci. 2014, 71, 2577–2604.
  165. Martín, M.; Macías, M.; León, J.; Escames, G.; Khaldy, H.; Acuña-Castroviejo, D. Melatonin Increases the Activity of the Oxidative Phosphorylation Enzymes and the Production of ATP in Rat Brain and Liver Mitochondria. Int. J. Biochem. Cell Biol. 2002, 34, 348–357.
  166. Chen, X.; Hao, B.; Li, D.; Reiter, R.J.; Bai, Y.; Abay, B.; Chen, G.; Lin, S.; Zheng, T.; Ren, Y.; et al. Melatonin Inhibits Lung Cancer Development by Reversing the Warburg Effect via Stimulating the SIRT3/PDH Axis. J. Pineal Res. 2021, e12755.
  167. Go, G.; Yoon, Y.M.; Yoon, S.; Lee, G.; Lim, J.H.; Han, S.-Y.; Lee, S.H. Melatonin Protects Chronic Kidney Disease Mesenchymal Stem/stromal Cells against Accumulation of Methylglyoxal via Modulation of Hexokinase-2 Expression. Biomol. Ther. 2022, 30, 28.
  168. Pérez-Torres, I.; Guarner-Lans, V.; Rubio-Ruiz, M.E. Reductive Stress in Inflammation-Associated Diseases and the Pro-Oxidant Effect of Antioxidant Agents. Int. J. Mol. Sci. 2017, 18, 2098.
  169. Dawson, T.L.; Gores, G.J.; Nieminen, A.L.; Herman, B.; Lemasters, J.J. Mitochondria as a Source of Reactive Oxygen Species during Reductive Stress in Rat Hepatocytes. Am. J. Physiol. 1993, 264 Pt 1, C961–C967.
  170. Hahn, A.; Parey, K.; Bublitz, M.; Mills, D.J.; Zickermann, V.; Vonck, J.; Kühlbrandt, W.; Meier, T. Structure of a Complete ATP Synthase Dimer Reveals the Molecular Basis of Inner Mitochondrial Membrane Morphology. Mol. Cell 2016, 63, 445–456.
  171. Esparza-Perusquía, M.; Olvera-Sánchez, S.; Pardo, J.P.; Mendoza-Hernández, G.; Martínez, F.; Flores-Herrera, O. Structural and Kinetics Characterization of the F1F0-ATP Synthase Dimer. New Repercussion of Monomer-Monomer Contact. Biochim. Biophys. Acta Bioenerg. 2017, 1858, 975–981.
  172. Davies, K.M.; Anselmi, C.; Wittig, I.; Faraldo-Gómez, J.D.; Kühlbrandt, W. Structure of the Yeast F1Fo-ATP Synthase Dimer and Its Role in Shaping the Mitochondrial Cristae. Proc. Natl. Acad. Sci. USA 2012, 109, 13602–13607.
  173. Spikes, T.E.; Montgomery, M.G.; Walker, J.E. Interface Mobility between Monomers in Dimeric Bovine ATP Synthase Participates in the Ultrastructure of Inner Mitochondrial Membranes. Proc. Natl. Acad. Sci. USA 2021, 118.
  174. Elías-Wolff, F.; Lindén, M.; Lyubartsev, A.P.; Brandt, E.G. Curvature Sensing by Cardiolipin in Simulated Buckled Membranes. Soft Matter 2019, 15, 792–802.
  175. Ikon, N.; Ryan, R.O. Cardiolipin and Mitochondrial Cristae Organization. Biochim. Biophys. Acta Biomembr. 2017, 1859, 1156–1163.
  176. Mileykovskaya, E.; Dowhan, W. Cardiolipin-Dependent Formation of Mitochondrial Respiratory Supercomplexes. Chem. Phys. Lipids 2014, 179, 42–48.
  177. Pfeiffer, K.; Gohil, V.; Stuart, R.A.; Hunte, C.; Brandt, U.; Greenberg, M.L.; Schägger, H. Cardiolipin Stabilizes Respiratory Chain Supercomplexes. J. Biol. Chem. 2003, 278, 52873–52880.
  178. Horvath, S.E.; Daum, G. Lipids of Mitochondria. Prog. Lipid Res. 2013, 52, 590–614.
  179. Agrawal, A.; Ramachandran, R. Exploring the Links between Lipid Geometry and Mitochondrial Fission: Emerging Concepts. Mitochondrion 2019, 49, 305–313.
  180. Parui, P.P.; Sarakar, Y.; Majumder, R.; Das, S.; Yang, H.; Yasuhara, K.; Hirota, S. Determination of Proton Concentration at Cardiolipin-Containing Membrane Interfaces and Its Relation with the Peroxidase Activity of Cytochrome C. Chem. Sci. 2019, 10, 9140–9151.
  181. Haines, T.H.; Dencher, N.A. Cardiolipin: A Proton Trap for Oxidative Phosphorylation. FEBS Lett. 2002, 528, 35–39.
  182. Afzal, N.; Lederer, W.J.; Jafri, M.S.; Mannella, C.A. Effect of Crista Morphology on Mitochondrial ATP Output: A Computational Study. Curr. Res. Physiol. 2021, 4, 163–176.
  183. Vähäheikkilä, M.; Peltomaa, T.; Róg, T.; Vazdar, M.; Pöyry, S.; Vattulainen, I. How Cardiolipin Peroxidation Alters the Properties of the Inner Mitochondrial Membrane? Chem. Phys. Lipids 2018, 214, 15–23.
  184. Claypool, S.M. Cardiolipin, a Critical Determinant of Mitochondrial Carrier Protein Assembly and Function. Biochim. Biophys. Acta 2009, 1788, 2059–2068.
  185. Paradies, G.; Paradies, V.; De Benedictis, V.; Ruggiero, F.M.; Petrosillo, G. Functional Role of Cardiolipin in Mitochondrial Bioenergetics. Biochim. Biophys. Acta 2014, 1837, 408–417.
  186. Shi, Y. Emerging Roles of Cardiolipin Remodeling in Mitochondrial Dysfunction Associated with Diabetes, Obesity, and Cardiovascular Diseases. J. Biomed. Res. 2010, 24, 6–15.
  187. Chicco, A.J.; Sparagna, G.C. Role of Cardiolipin Alterations in Mitochondrial Dysfunction and Disease. Am. J. Physiol. Cell Physiol. 2007, 292, C33–C44.
  188. Paradies, G.; Paradies, V.; Ruggiero, F.M.; Petrosillo, G. Mitochondrial Bioenergetics and Cardiolipin Alterations in Myocardial Ischemia-Reperfusion Injury: Implications for Pharmacological Cardioprotection. Am. J. Physiol. Heart Circ. Physiol. 2018, 315, H1341–H1352.
  189. Dolinsky, V.W.; Cole, L.K.; Sparagna, G.C.; Hatch, G.M. Cardiac Mitochondrial Energy Metabolism in Heart Failure: Role of Cardiolipin and Sirtuins. Biochim. Biophys. Acta 2016, 1861, 1544–1554.
  190. Han, X.; Yang, J.; Yang, K.; Zhao, Z.; Abendschein, D.R.; Gross, R.W. Alterations in Myocardial Cardiolipin Content and Composition Occur at the Very Earliest Stages of Diabetes: A Shotgun Lipidomics Study. Biochemistry 2007, 46, 6417–6428.
  191. Schlame, M.; Ren, M. Barth Syndrome, a Human Disorder of Cardiolipin Metabolism. FEBS Lett. 2006, 580, 5450–5455.
  192. Jiang, F.; Ryan, M.T.; Schlame, M.; Zhao, M.; Gu, Z.; Klingenberg, M.; Pfanner, N.; Greenberg, M.L. Absence of Cardiolipin in the crd1 Null Mutant Results in Decreased Mitochondrial Membrane Potential and Reduced Mitochondrial Function. J. Biol. Chem. 2000, 275, 22387–22394.
  193. Ikonomidis, I.; Lekakis, J.; Vamvakou, G.; Loizou, S.; Revela, I.; Andreotti, F.; Kremastinos, D.T.; Nihoyannopoulos, P. IgA Anticardiolipin Antibody Is Associated with the Extent of Daily-Life Ischaemia in Patients with Chronic Coronary Artery Disease. Heart 2007, 93, 1412–1413.
  194. Saleh, J.; Peyssonnaux, C.; Singh, K.K.; Edeas, M. Mitochondria and Microbiota Dysfunction in COVID-19 Pathogenesis. Mitochondrion 2020, 54, 1–7.
  195. Taha, M.; Samavati, L. Antiphospholipid Antibodies in COVID-19: A Meta-Analysis and Systematic Review. RMD Open 2021, 7, e001580.
  196. Craig, W.Y.; Poulin, S.E.; Neveux, L.M.; Palomaki, G.E.; Dostal-Johnson, D.A.; Ledue, T.B.; Ritchie, R.F. Anti-Oxidized LDL Antibodies and Antiphospholipid Antibodies in Healthy Subjects: Relationship with Lipoprotein- and Oxidation-Related Analytes. J. Autoimmun. 1995, 8, 713–726.
  197. Hasan Ali, O.; Bomze, D.; Risch, L.; Brugger, S.D.; Paprotny, M.; Weber, M.; Thiel, S.; Kern, L.; Albrich, W.C.; Kohler, P.; et al. Severe Coronavirus Disease 2019 (COVID-19) Is Associated With Elevated Serum Immunoglobulin (Ig) A and Antiphospholipid IgA Antibodies. Clin. Infect. Dis. 2021, 73, e2869–e2874.
  198. Martín-Fernández, M.; Aller, R.; Heredia-Rodríguez, M.; Gómez-Sánchez, E.; Martínez-Paz, P.; Gonzalo-Benito, H.; Sánchez-de Prada, L.; Gorgojo, Ó.; Carnicero-Frutos, I.; Tamayo, E.; et al. Lipid Peroxidation as a Hallmark of Severity in COVID-19 Patients. Redox Biol. 2021, 48, 102181.
  199. Petrosillo, G.; Di Venosa, N.; Pistolese, M.; Casanova, G.; Tiravanti, E.; Colantuono, G.; Federici, A.; Paradies, G.; Ruggiero, F.M. Protective Effect of Melatonin against Mitochondrial Dysfunction Associated with Cardiac Ischemia- Reperfusion: Role of Cardiolipin. FASEB J. 2006, 20, 269–276.
  200. Römsing, S. Development and Validation of Bioanalytical Methods: Application to Melatonin and Selected Anti-Infective Drugs. Ph.D. Thesis, Acta Universitatis Upsaliensis, Uppsala, Sweden, 2010.
  201. Bongiorno, D.; Ceraulo, L.; Ferrugia, M.; Filizzola, F.; Giordano, C.; Ruggirello, A.; Liveri, V.T. H-NMR and FT-IR Study of the State of Melatonin Confined in Membrane Models: Location and Interactions of Melatonin in Water Free Lecithin and AOT Reversed Micelles. ARKIVOC 2004, 2004, 251–262.
  202. Ceraulo, L.; Ferrugia, M.; Tesoriere, L.; Segreto, S.; Livrea, M.A.; Turco Liveri, V. Interactions of Melatonin with Membrane Models: Portioning of Melatonin in AOT and Lecithin Reversed Micelles. J. Pineal Res. 1999, 26, 108–112.
  203. Galano, A.; Tan, D.X.; Reiter, R.J. Cyclic 3-Hydroxymelatonin, a Key Metabolite Enhancing the Peroxyl Radical Scavenging Activity of Melatonin. RSC Adv. 2014, 4, 5220.
  204. Tan, D.X.; Manchester, L.C.; Reiter, R.J.; Plummer, B.F. Cyclic 3-Hydroxymelatonin: A Melatonin Metabolite Generated as a Result of Hydroxyl Radical Scavenging. Biol. Signals Recept. 1999, 8, 70–74.
  205. Bielski, B.H.; Arudi, R.L.; Sutherland, M.W. A Study of the Reactivity of HO2/O2- with Unsaturated Fatty Acids. J. Biol. Chem. 1983, 258, 4759–4761.
  206. Aikens, J.; Dix, T.A. Perhydroxyl Radical (HOO.) Initiated Lipid Peroxidation. The Role of Fatty Acid Hydroperoxides. J. Biol. Chem. 1991, 266, 15091–15098.
  207. Ademowo, O.S.; Dias, H.K.I.; Burton, D.G.A.; Griffiths, H.R. Lipid (per) Oxidation in Mitochondria: An Emerging Target in the Ageing Process? Biogerontology 2017, 18, 859–879.
  208. Repetto, M.; Semprine, J.; Boveris, A. Lipid Peroxidation: Chemical Mechanism, Biological Implications and Analytical Determination. In Lipid Peroxidation; Catala, A., Ed.; IntechOpen: Rijeka, Croatia, 2012.
  209. Ito, M.; Yanagi, Y.; Ichinohe, T. Encephalomyocarditis Virus Viroporin 2B Activates NLRP3 Inflammasome. PLoS Pathog. 2012, 8, e1002857.
  210. Toldo, S.; Bussani, R.; Nuzzi, V.; Bonaventura, A.; Mauro, A.G.; Cannatà, A.; Pillappa, R.; Sinagra, G.; Nana-Sinkam, P.; Sime, P.; et al. Inflammasome Formation in the Lungs of Patients with Fatal COVID-19. Inflamm. Res. 2021, 70, 7–10.
  211. Arioz, B.I.; Tarakcioglu, E.; Olcum, M.; Genc, S. The Role of Melatonin on NLRP3 Inflammasome Activation in Diseases. Antioxidants 2021, 10, 1020.
  212. Seoane, P.I.; Lee, B.; Hoyle, C.; Yu, S.; Lopez-Castejon, G.; Lowe, M.; Brough, D. The NLRP3-Inflammasome as a Sensor of Organelle Dysfunction. J. Cell Biol. 2020, 219, 12.
  213. Samir, P.; Kanneganti, T.-D. DDX3X Sits at the Crossroads of Liquid-Liquid and Prionoid Phase Transitions Arbitrating Life and Death Cell Fate Decisions in Stressed Cells. DNA Cell Biol. 2020, 39, 1091–1095.
  214. Franklin, B.S.; Bossaller, L.; De Nardo, D.; Ratter, J.M.; Stutz, A.; Engels, G.; Brenker, C.; Nordhoff, M.; Mirandola, S.R.; Al-Amoudi, A.; et al. The Adaptor ASC Has Extracellular and “Prionoid” Activities That Propagate Inflammation. Nat. Immunol. 2014, 15, 727–737.
  215. Xia, S.; Chen, Z.; Shen, C.; Fu, T.-M. Higher-Order Assemblies in Immune Signaling: Supramolecular Complexes and Phase Separation. Protein Cell 2021, 12, 680–694.
  216. He, Y.; Hara, H.; Núñez, G. Mechanism and Regulation of NLRP3 Inflammasome Activation. Trends Biochem. Sci. 2016, 41, 1012–1021.
  217. Huang, Y.; Xu, W.; Zhou, R. NLRP3 Inflammasome Activation and Cell Death. Cell. Mol. Immunol. 2021, 18, 2114–2127.
  218. Franchi, L.; Eigenbrod, T.; Muñoz-Planillo, R.; Nuñez, G. The Inflammasome: A Caspase-1-Activation Platform That Regulates Immune Responses and Disease Pathogenesis. Nat. Immunol. 2009, 10, 241–247.
  219. Huang, Z.; Tyurina, Y.Y.; Jiang, J.; Tokarska-Schlattner, M.; Boissan, M.; Lacombe, M.-L.; Epand, R.; Schlattner, U.; Epand, R.M.; Kagan, V.E. Externalization of Cardiolipin as an “Eat-Me” Mitophageal Signal Is Facilitated by NDPK-D. Biophys. J. 2014, 106, 184a.
  220. Elliott, E.I.; Miller, A.N.; Banoth, B.; Iyer, S.S.; Stotland, A.; Weiss, J.P.; Gottlieb, R.A.; Sutterwala, F.S.; Cassel, S.L. Cutting Edge: Mitochondrial Assembly of the NLRP3 Inflammasome Complex Is Initiated at Priming. J. Immunol. 2018, 200, 3047–3052.
  221. Sefik, E.; Qu, R.; Junqueira, C.; Kaffe, E.; Mirza, H.; Zhao, J.; Brewer, J.R.; Han, A.; Steach, H.R.; Israelow, B.; et al. Inflammasome Activation in Infected Macrophages Drives COVID-19 Pathology. Nature 2022, 606, 585–593.
  222. Yalcinkaya, M.; Liu, W.; Islam, M.N.; Kotini, A.G.; Gusarova, G.A.; Fidler, T.P.; Papapetrou, E.P.; Bhattacharya, J.; Wang, N.; Tall, A.R. Modulation of the NLRP3 Inflammasome by Sars-CoV-2 Envelope Protein. Sci. Rep. 2021, 11, 24432.
  223. Freeman, T.L.; Swartz, T.H. Targeting the NLRP3 Inflammasome in Severe COVID-19. Front. Immunol. 2020, 11, 1518.
  224. Zeng, J.; Xie, X.; Feng, X.-L.; Xu, L.; Han, J.-B.; Yu, D.; Zou, Q.-C.; Liu, Q.; Li, X.; Ma, G.; et al. Specific Inhibition of the NLRP3 Inflammasome Suppresses Immune Overactivation and Alleviates COVID-19 like Pathology in Mice. EBioMedicine 2022, 75, 103803.
  225. Zhao, N.; Di, B.; Xu, L.-L. The NLRP3 Inflammasome and COVID-19: Activation, Pathogenesis and Therapeutic Strategies. Cytokine Growth Factor Rev. 2021, 61, 2–15.
  226. Abais, J.M.; Xia, M.; Zhang, Y.; Boini, K.M.; Li, P.-L. Redox Regulation of NLRP3 Inflammasomes: ROS as Trigger or Effector? Antioxid. Redox Signal. 2015, 22, 1111–1129.
  227. Mitroulis, I.; Skendros, P.; Ritis, K. Targeting IL-1beta in Disease; the Expanding Role of NLRP3 Inflammasome. Eur. J. Intern. Med. 2010, 21, 157–163.
  228. Tőzsér, J.; Benkő, S. Natural Compounds as Regulators of NLRP3 Inflammasome-Mediated IL-1β Production. Mediat. Inflamm. 2016, 2016, 5460302.
  229. Yang, D.; Elner, S.G.; Bian, Z.-M.; Till, G.O.; Petty, H.R.; Elner, V.M. Pro-Inflammatory Cytokines Increase Reactive Oxygen Species through Mitochondria and NADPH Oxidase in Cultured RPE Cells. Exp. Eye Res. 2007, 85, 462–472.
  230. Yoo, H.G.; Shin, B.A.; Park, J.S.; Lee, K.H.; Chay, K.O.; Yang, S.Y.; Ahn, B.W.; Jung, Y.D. IL-1beta Induces MMP-9 via Reactive Oxygen Species and NF-kappaB in Murine Macrophage RAW 264.7 Cells. Biochem. Biophys. Res. Commun. 2002, 298, 251–256.
  231. Ratajczak, M.Z.; Kucia, M. SARS-CoV-2 Infection and Overactivation of Nlrp3 Inflammasome as a Trigger of Cytokine “Storm” and Risk Factor for Damage of Hematopoietic Stem Cells. Leukemia 2020, 34, 1726–1729.
  232. Somasekharan, S.P.; Gleave, M. SARS-CoV-2 Nucleocapsid Protein Interacts with Immunoregulators and Stress Granules and Phase Separates to Form Liquid Droplets. FEBS Lett. 2021, 595, 2872–2896.
  233. Park, S.H.; Lee, S.G.; Kim, Y.; Song, K. Assignment of a Human Putative RNA Helicase Gene, DDX3, to Human X Chromosome Bands p11.3-->p11.23. Cytogenet. Cell Genet. 1998, 81, 178–179.
  234. Vesuna, F.; Akhrymuk, I.; Smith, A.; Winnard, P.T.; Lin, S.-C.; Scharpf, R.; Kehn-Hall, K.; Raman, V. RK-33, a Small Molecule Inhibitor of Host RNA Helicase DDX3, Suppresses Multiple Variants of SARS-CoV-2. bioRxiv 2022.
  235. Kumar, R.; Singh, N.; Abdin, M.Z.; Patel, A.H.; Medigeshi, G.R. Dengue Virus Capsid Interacts with DDX3X-A Potential Mechanism for Suppression of Antiviral Functions in Dengue Infection. Front. Cell. Infect. Microbiol. 2017, 7, 542.
  236. Brai, A.; Riva, V.; Saladini, F.; Zamperini, C.; Trivisani, C.I.; Garbelli, A.; Pennisi, C.; Giannini, A.; Boccuto, A.; Bugli, F.; et al. DDX3X Inhibitors, an Effective Way to Overcome HIV-1 Resistance Targeting Host Proteins. Eur. J. Med. Chem. 2020, 200, 112319.
  237. Yedavalli, V.S.R.K.; Neuveut, C.; Chi, Y.-H.; Kleiman, L.; Jeang, K.-T. Requirement of DDX3 DEAD Box RNA Helicase for HIV-1 Rev-RRE Export Function. Cell 2004, 119, 381–392.
  238. Pène, V.; Li, Q.; Sodroski, C.; Hsu, C.-S.; Liang, T.J. Dynamic Interaction of Stress Granules, DDX3X, and IKK-α Mediates Multiple Functions in Hepatitis C Virus Infection. J. Virol. 2015, 89, 5462–5477.
  239. Nelson, C.; Mrozowich, T.; Gemmill, D.L.; Park, S.M.; Patel, T.R. Human DDX3X Unwinds Japanese Encephalitis and Zika Viral 5′ Terminal Regions. Int. J. Mol. Sci. 2021, 22, 413.
  240. Winnard, P.T., Jr.; Vesuna, F.; Raman, V. Targeting Host DEAD-Box RNA Helicase DDX3X for Treating Viral Infections. Antivir. Res. 2021, 185, 104994.
  241. Saito, M.; Iestamantavicius, V.; Hess, D.; Matthias, P. Monitoring Acetylation of the RNA Helicase DDX3X, a Protein Critical for Formation of Stress Granules. In RNA Remodeling Proteins: Methods and Protocols; Boudvillain, M., Ed.; Springer: New York, NY, USA, 2021; pp. 217–234.
  242. Samir, P.; Kesavardhana, S.; Patmore, D.M.; Gingras, S.; Malireddi, R.K.S.; Karki, R.; Guy, C.S.; Briard, B.; Place, D.E.; Bhattacharya, A.; et al. DDX3X Acts as a Live-or-Die Checkpoint in Stressed Cells by Regulating NLRP3 Inflammasome. Nature 2019, 573, 590–594.
  243. Cui, B.C.; Sikirzhytski, V.; Aksenova, M.; Lucius, M.D.; Levon, G.H.; Mack, Z.T.; Pollack, C.; Odhiambo, D.; Broude, E.; Lizarraga, S.B.; et al. Pharmacological Inhibition of DEAD-Box RNA Helicase 3 Attenuates Stress Granule Assembly. Biochem. Pharmacol. 2020, 182, 114280.
  244. Lage, S.L.; Amaral, E.P.; Hilligan, K.L.; Laidlaw, E.; Rupert, A.; Namasivayan, S.; Rocco, J.; Galindo, F.; Kellogg, A.; Kumar, P.; et al. Persistent Oxidative Stress and Inflammasome Activation in CD14highCD16- Monocytes From COVID-19 Patients. Front. Immunol. 2021, 12, 799558.
  245. Mishra, S.R.; Mahapatra, K.K.; Behera, B.P.; Patra, S.; Bhol, C.S.; Panigrahi, D.P.; Praharaj, P.P.; Singh, A.; Patil, S.; Dhiman, R.; et al. Mitochondrial Dysfunction as a Driver of NLRP3 Inflammasome Activation and Its Modulation through Mitophagy for Potential Therapeutics. Int. J. Biochem. Cell Biol. 2021, 136, 106013.
  246. Favero, G.; Franceschetti, L.; Bonomini, F.; Rodella, L.F.; Rezzani, R. Melatonin as an Anti-Inflammatory Agent Modulating Inflammasome Activation. Int. J. Endocrinol. 2017, 2017, 1835195.
  247. Zhang, Y.; Li, X.; Grailer, J.J.; Wang, N.; Wang, M.; Yao, J.; Zhong, R.; Gao, G.F.; Ward, P.A.; Tan, D.-X.; et al. Melatonin Alleviates Acute Lung Injury through Inhibiting the NLRP3 Inflammasome. J. Pineal Res. 2016, 60, 405–414.
  248. Ma, S.; Chen, J.; Feng, J.; Zhang, R.; Fan, M.; Han, D.; Li, X.; Li, C.; Ren, J.; Wang, Y.; et al. Melatonin Ameliorates the Progression of Atherosclerosis via Mitophagy Activation and NLRP3 Inflammasome Inhibition. Oxid. Med. Cell. Longev. 2018, 2018, 9286458.
  249. Yoneyama, M.; Jogi, M.; Onomoto, K. Regulation of Antiviral Innate Immune Signaling by Stress-Induced RNA Granules. J. Biochem. 2016, 159, 279–286.
  250. McCormick, C.; Khaperskyy, D.A. Translation Inhibition and Stress Granules in the Antiviral Immune Response. Nat. Rev. Immunol. 2017, 17, 647–660.
  251. Miller, C.L. Stress Granules and Virus Replication. Future Virol. 2011, 6, 1329–1338.
  252. de Castro, I.F.; Volonté, L.; Risco, C. Virus Factories: Biogenesis and Structural Design. Cell. Microbiol. 2013, 15, 24–34.
  253. Savastano, A.; Ibáñez de Opakua, A.; Rankovic, M.; Zweckstetter, M. Nucleocapsid Protein of SARS-CoV-2 Phase Separates into RNA-Rich Polymerase-Containing Condensates. Nat. Commun. 2020, 11, 6041.
  254. Klein, S.; Cortese, M.; Winter, S.L.; Wachsmuth-Melm, M.; Neufeldt, C.J.; Cerikan, B.; Stanifer, M.L.; Boulant, S.; Bartenschlager, R.; Chlanda, P. SARS-CoV-2 Structure and Replication Characterized by in Situ Cryo-Electron Tomography. Nat. Commun. 2020, 11, 5885.
  255. Squeglia, F.; Romano, M.; Ruggiero, A.; Maga, G.; Berisio, R. Host DDX Helicases as Possible SARS-CoV-2 Proviral Factors: A Structural Overview of Their Hijacking Through Multiple Viral Proteins. Front. Chem. 2020, 8, 602162.
  256. Ciccosanti, F.; Di Rienzo, M.; Romagnoli, A.; Colavita, F.; Refolo, G.; Castilletti, C.; Agrati, C.; Brai, A.; Manetti, F.; Botta, L.; et al. Proteomic Analysis Identifies the RNA Helicase DDX3X as a Host Target against SARS-CoV-2 Infection. Antivir. Res. 2021, 190, 105064.
  257. Hernández-Díaz, T.; Valiente-Echeverría, F.; Soto-Rifo, R. RNA Helicase DDX3: A Double-Edged Sword for Viral Replication and Immune Signaling. Microorganisms 2021, 9, 1206.
  258. Valiente-Echeverría, F.; Hermoso, M.A.; Soto-Rifo, R. RNA Helicase DDX3: At the Crossroad of Viral Replication and Antiviral Immunity. Rev. Med. Virol. 2015, 25, 286–299.
  259. Riva, V.; Maga, G. From the Magic Bullet to the Magic Target: Exploiting the Diverse Roles of DDX3X in Viral Infections and Tumorigenesis. Future Med. Chem. 2019, 11, 1357–1381.
  260. Wang, W.; Jia, M.; Zhao, C.; Yu, Z.; Song, H.; Qin, Y.; Zhao, W. RNF39 Mediates K48-Linked Ubiquitination of DDX3X and Inhibits RLR-Dependent Antiviral Immunity. Sci. Adv. 2021, 7, eabe5877.
  261. Soulat, D.; Bürckstümmer, T.; Westermayer, S.; Goncalves, A.; Bauch, A.; Stefanovic, A.; Hantschel, O.; Bennett, K.L.; Decker, T.; Superti-Furga, G. The DEAD-Box Helicase DDX3X Is a Critical Component of the TANK-Binding Kinase 1-Dependent Innate Immune Response. EMBO J. 2008, 27, 2135–2146.
  262. Oshiumi, H.; Sakai, K.; Matsumoto, M.; Seya, T. DEAD/H BOX 3 (DDX3) Helicase Binds the RIG-I Adaptor IPS-1 to up-Regulate IFN-Beta-Inducing Potential. Eur. J. Immunol. 2010, 40, 940–948.
  263. Oshiumi, H.; Kouwaki, T.; Seya, T. Accessory Factors of Cytoplasmic Viral RNA Sensors Required for Antiviral Innate Immune Response. Front. Immunol. 2016, 7, 200.
  264. Wang, S.; Dai, T.; Qin, Z.; Pan, T.; Chu, F.; Lou, L.; Zhang, L.; Yang, B.; Huang, H.; Lu, H.; et al. Targeting Liquid-Liquid Phase Separation of SARS-CoV-2 Nucleocapsid Protein Promotes Innate Antiviral Immunity by Elevating MAVS Activity. Nat. Cell Biol. 2021, 23, 718–732.
  265. Hou, F.; Sun, L.; Zheng, H.; Skaug, B.; Jiang, Q.-X.; Chen, Z.J. MAVS Forms Functional Prion-like Aggregates to Activate and Propagate Antiviral Innate Immune Response. Cell 2011, 146, 448–461.
  266. John, L.; Samuel, C.E. Induction of Stress Granules by Interferon and down-Regulation by the Cellular RNA Adenosine Deaminase ADAR1. Virology 2014, 454–455, 299–310.
  267. Protter, D.S.W.; Parker, R. Principles and Properties of Stress Granules. Trends Cell Biol. 2016, 26, 668–679.
  268. Parker, F.; Maurier, F.; Delumeau, I.; Duchesne, M.; Faucher, D.; Debussche, L.; Dugue, A.; Schweighoffer, F.; Tocque, B. A Ras-GTPase-Activating Protein SH3-Domain-Binding Protein. Mol. Cell. Biol. 1996, 16, 2561–2569.
  269. Yang, P.; Mathieu, C.; Kolaitis, R.-M.; Zhang, P.; Messing, J.; Yurtsever, U.; Yang, Z.; Wu, J.; Li, Y.; Pan, Q.; et al. G3BP1 Is a Tunable Switch That Triggers Phase Separation to Assemble Stress Granules. Cell 2020, 181, 325–345.e28.
  270. Wu, J.; Liu, W.; Gong, P. A Structural Overview of RNA-Dependent RNA Polymerases from the Flaviviridae Family. Int. J. Mol. Sci. 2015, 16, 12943–12957.
  271. Deater, M.; Tamhankar, M.; Lloyd, R.E. TDRD3 Is an Antiviral Restriction Factor That Promotes IFN Signaling with G3BP1. PLoS Pathog. 2022, 18, e1010249.
  272. Yang, W.; Ru, Y.; Ren, J.; Bai, J.; Wei, J.; Fu, S.; Liu, X.; Li, D.; Zheng, H. G3BP1 Inhibits RNA Virus Replication by Positively Regulating RIG-I-Mediated Cellular Antiviral Response. Cell Death Dis. 2019, 10, 946.
  273. Biswal, M.; Lu, J.; Song, J. SARS-CoV-2 Nucleocapsid Protein Targets a Conserved Surface Groove of the NTF2-like Domain of G3BP1. J. Mol. Biol. 2022, 434, 167516.
  274. Nabeel-Shah, S.; Lee, H.; Ahmed, N.; Burke, G.L.; Farhangmehr, S.; Ashraf, K.; Pu, S.; Braunschweig, U.; Zhong, G.; Wei, H.; et al. SARS-CoV-2 Nucleocapsid Protein Binds Host mRNAs and Attenuates Stress Granules to Impair Host Stress Response. iScience 2022, 25, 103562.
  275. Wang, J.; Shi, C.; Xu, Q.; Yin, H. SARS-CoV-2 Nucleocapsid Protein Undergoes Liquid-Liquid Phase Separation into Stress Granules through Its N-Terminal Intrinsically Disordered Region. Cell Discov. 2021, 7, 5.
  276. Lian, X.J.; Gallouzi, I.-E. Oxidative Stress Increases the Number of Stress Granules in Senescent Cells and Triggers a Rapid Decrease in p21waf1/cip1 Translation. J. Biol. Chem. 2009, 284, 8877–8887.
  277. Luo, L.; Li, Z.; Zhao, T.; Ju, X.; Ma, P.; Jin, B.; Zhou, Y.; He, S.; Huang, J.; Xu, X.; et al. SARS-CoV-2 Nucleocapsid Protein Phase Separates with G3BPs to Disassemble Stress Granules and Facilitate Viral Production. Sci. Bull. 2021, 66, 1194–1204.
  278. Henninger, J.E.; Oksuz, O.; Shrinivas, K.; Sagi, I.; LeRoy, G.; Zheng, M.M.; Andrews, J.O.; Zamudio, A.V.; Lazaris, C.; Hannett, N.M.; et al. RNA-Mediated Feedback Control of Transcriptional Condensates. Cell 2021, 184, 207–225.e24.
  279. Lu, S.; Ye, Q.; Singh, D.; Cao, Y.; Diedrich, J.K.; Yates, J.R., 3rd; Villa, E.; Cleveland, D.W.; Corbett, K.D. The SARS-CoV-2 Nucleocapsid Phosphoprotein Forms Mutually Exclusive Condensates with RNA and the Membrane-Associated M Protein. Nat. Commun. 2021, 12, 502.
  280. Lu, S.; Deng, R.; Jiang, H.; Song, H.; Li, S.; Shen, Q.; Huang, W.; Nussinov, R.; Yu, J.; Zhang, J. The Mechanism of ATP-Dependent Allosteric Protection of Akt Kinase Phosphorylation. Structure 2015, 23, 1725–1734.
  281. Carlson, C.R.; Asfaha, J.B.; Ghent, C.M.; Howard, C.J.; Hartooni, N.; Safari, M.; Frankel, A.D.; Morgan, D.O. Phosphoregulation of Phase Separation by the SARS-CoV-2 N Protein Suggests a Biophysical Basis for Its Dual Functions. Mol. Cell 2020, 80, 1092–1103.e4.
  282. Lier, C.; Becker, S.; Biedenkopf, N. Dynamic Phosphorylation of Ebola Virus VP30 in NP-Induced Inclusion Bodies. Virology 2017, 512, 39–47.
  283. Mühlberger, E.; Weik, M.; Volchkov, V.E.; Klenk, H.D.; Becker, S. Comparison of the Transcription and Replication Strategies of Marburg Virus and Ebola Virus by Using Artificial Replication Systems. J. Virol. 1999, 73, 2333–2342.
  284. Nikolakaki, E.; Giannakouros, T. SR/RS Motifs as Critical Determinants of Coronavirus Life Cycle. Front. Mol. Biosci. 2020, 7, 219.
  285. Sarhan, M.A.; Abdel-Hakeem, M.S.; Mason, A.L.; Tyrrell, D.L.; Houghton, M. Glycogen Synthase Kinase 3β Inhibitors Prevent Hepatitis C Virus Release/assembly through Perturbation of Lipid Metabolism. Sci. Rep. 2017, 7, 1–12.
  286. Cuartas-López, A.M.; Gallego-Gómez, J.C. Glycogen Synthase Kinase 3ß Participates in Late Stages of Dengue Virus-2 Infection. Mem. Inst. Oswaldo Cruz 2020, 115, e190357.
  287. Guendel, I.; Iordanskiy, S.; Van Duyne, R.; Kehn-Hall, K.; Saifuddin, M.; Das, R.; Jaworski, E.; Sampey, G.C.; Senina, S.; Shultz, L.; et al. Novel Neuroprotective GSK-3β Inhibitor Restricts Tat-Mediated HIV-1 Replication. J. Virol. 2014, 88, 1189–1208.
  288. Marineau, A.; Khan, K.A.; Servant, M.J. Roles of GSK-3 and β-Catenin in Antiviral Innate Immune Sensing of Nucleic Acids. Cells 2020, 9, 897.
  289. Alfhili, M.A.; Alsughayyir, J.; McCubrey, J.A.; Akula, S.M. GSK-3-Associated Signaling Is Crucial to Virus Infection of Cells. Biochim. Biophys. Acta Mol. Cell Res. 2020, 1867, 118767.
  290. Jope, R.S.; Yuskaitis, C.J.; Beurel, E. Glycogen Synthase Kinase-3 (GSK3): Inflammation, Diseases, and Therapeutics. Neurochem. Res. 2007, 32, 577–595.
  291. Dugo, L.; Collin, M.; Allen, D.A.; Patel, N.S.A.; Bauer, I.; Mervaala, E.M.A.; Louhelainen, M.; Foster, S.J.; Yaqoob, M.M.; Thiemermann, C. GSK-3beta Inhibitors Attenuate the Organ Injury/dysfunction Caused by Endotoxemia in the Rat. Crit. Care Med. 2005, 33, 1903–1912.
  292. Hoeflich, K.P.; Luo, J.; Rubie, E.A.; Tsao, M.S.; Jin, O.; Woodgett, J.R. Requirement for Glycogen Synthase Kinase-3beta in Cell Survival and NF-kappaB Activation. Nature 2000, 406, 86–90.
  293. Chen, X.; Liu, Y.; Zhu, J.; Lei, S.; Dong, Y.; Li, L.; Jiang, B.; Tan, L.; Wu, J.; Yu, S.; et al. GSK-3β Downregulates Nrf2 in Cultured Cortical Neurons and in a Rat Model of Cerebral Ischemia-Reperfusion. Sci. Rep. 2016, 6, 20196.
  294. Lu, M.; Wang, P.; Qiao, Y.; Jiang, C.; Ge, Y.; Flickinger, B.; Malhotra, D.K.; Dworkin, L.D.; Liu, Z.; Gong, R. GSK3β-Mediated Keap1-Independent Regulation of Nrf2 Antioxidant Response: A Molecular Rheostat of Acute Kidney Injury to Chronic Kidney Disease Transition. Redox Biol. 2019, 26, 101275.
  295. Culbreth, M.; Aschner, M. GSK-3β, a Double-Edged Sword in Nrf2 Regulation: Implications for Neurological Dysfunction and Disease. F1000Research 2018, 7, 1043.
  296. Aryanpur, P.P.; Regan, C.A.; Collins, J.M.; Mittelmeier, T.M.; Renner, D.M.; Vergara, A.M.; Brown, N.P.; Bolger, T.A. Gle1 Regulates RNA Binding of the DEAD-Box Helicase Ded1 in Its Complex Role in Translation Initiation. Mol. Cell. Biol. 2017, 37.
  297. Aditi; Folkmann, A.W.; Wente, S.R. Cytoplasmic hGle1A Regulates Stress Granules by Modulation of Translation. Mol. Biol. Cell 2015, 26, 1476–1490.
  298. Das, R.; Balmik, A.A.; Chinnathambi, S. Melatonin Reduces GSK3β-Mediated Tau Phosphorylation, Enhances Nrf2 Nuclear Translocation and Anti-Inflammation. ASN Neuro 2020, 12, 1759091420981204.
  299. Rhee, Y.-H.; Ahn, J.-C. Melatonin Attenuated Adipogenesis through Reduction of the CCAAT/enhancer Binding Protein Beta by Regulating the Glycogen Synthase 3 Beta in Human Mesenchymal Stem Cells. J. Physiol. Biochem. 2016, 72, 145–155.
  300. Hadj Ayed Tka, K.; Mahfoudh Boussaid, A.; Zaouali, M.A.; Kammoun, R.; Bejaoui, M.; Ghoul Mazgar, S.; Rosello Catafau, J.; Ben Abdennebi, H. Melatonin Modulates Endoplasmic Reticulum Stress and Akt/GSK3-Beta Signaling Pathway in a Rat Model of Renal Warm Ischemia Reperfusion. Anal. Cell. Pathol. 2015, 2015, 635172.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
This entry is offline, you can click here to edit this entry!
ScholarVision Creations
Feedback