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Loh, D. Melatonin: ATP Regulation in MLOs. Encyclopedia. Available online: (accessed on 24 February 2024).
Loh D. Melatonin: ATP Regulation in MLOs. Encyclopedia. Available at: Accessed February 24, 2024.
Loh, Doris. "Melatonin: ATP Regulation in MLOs" Encyclopedia, (accessed February 24, 2024).
Loh, D. (2021, September 27). Melatonin: ATP Regulation in MLOs. In Encyclopedia.
Loh, Doris. "Melatonin: ATP Regulation in MLOs." Encyclopedia. Web. 27 September, 2021.
Melatonin: ATP Regulation in MLOs

Biomolecular condensates are membraneless organelles (MLOs) that form dynamic, chemically distinct subcellular compartments organizing macromolecules such as proteins, RNA, and DNA in unicellular prokaryotic bacteria and complex eukaryotic cells. Separated from surrounding environments, MLOs in the nucleoplasm, cytoplasm, and mitochondria assemble by liquid–liquid phase separation (LLPS) into transient, non-static, liquid-like droplets that regulate essential molecular functions. LLPS is primarily controlled by ATP-dependent post-translational modifications (PTMs) that fine-tune the balance between attractive and repulsive charge states and/or binding motifs of proteins. Aberrant phase separation due to the absence of adequate hydrotropic small molecules such as ATP can cause pathological protein aggregation in diseases such as neurodegenerative disorders. Melatonin is a potent antioxidant capable of protecting cardiolipin and membrane lipids raft domains from peroxidation to support ATPase functionality and ion channel activities that may exert a dominant influence over phase separation in biomolecular condensates during condensate coacervation or dissolution processes that are ATP-dependent. 

melatonin biomolecular condensate neurodegenerative disorder liquid–liquid phase separation ATP lipid raft post-translational modification m6A RNA

1. Introduction

Present in all cells, biomolecular condensates are membraneless organelles (MLOs) containing proteins, ribonucleic acids (RNAs), and other nucleic acids [1]. These micron-scale macromolecules that can assemble into liquid-like droplets have been proposed to be the origin of life [2]. Current cell and molecular biology reveal that liquid–liquid phase separation (LLPS) is the driving force behind the assembly or dissolution of biomolecules in energy-efficient, rapid, essential reactions to changing endogenous or exogenous conditions including stress response [3] and signal transduction [4][5], as well as genome expression, organization, and repair [6]. LLPS creates distinct compartments that enhance or restrict biochemical reactions by enriching or excluding biomolecules from their environment [7]. Increasing evidence associates diseases such as neurodegeneration and cancer with the formation of protein aggregates from dysregulated, aberrant transitions in phase separation [8][9][10][11][12].
Phase separation at its core is a thermodynamic process driven by the reduction or a negative change in global free energy [1][13]. LLPS is entropically unfavorable; therefore, multivalent protein–protein interactions that are energetically favorable may be necessary to offset energetic costs [14]. Adenosine triphosphate (ATP) is the molecule favored by most organisms for capturing and transferring free energy. During hydrolysis, ATP is transformed into adenosine diphosphate (ADP) and inorganic phosphate (Pi). The change in free energy of −7.3 kcal/mol associated with this chemical reaction is used by cells to perform energetically favorable reactions [15], including relevant post-translational modification (PTM) such as phosphorylation [16], ubiquitination [17][18], and SUMOylation that may regulate condensate nucleation, composition, and growth [19][20]. It is understood that most proteins in the human proteome can undergo LLPS, assembling into dense liquid-like, reversible droplets under most physiological conditions [21]. Thermodynamic non-equilibrium processes facilitate the constant exchange of substrates and information that allow these condensates to perform important biological functions [22]. The phase transition of these functionally relevant proteins from their native to droplet states are often mediated and stabilized by ATP-dependent factors such as PTM and RNA. RNAs are critical architectural components that can fine-tune biophysical properties such as viscosity and dynamics in the regulation of spatiotemporal distribution of condensates [23][24].
Aberrant phase separation leading to the pathological amyloid fibrillation of fused in sarcoma (FUS), TAR DNA-binding protein 43 (TDP-43), tau, and α-synuclein (α-Syn} are now associated with neurodegenerative disorders such as amyotrophic lateral sclerosis (ALS), frontotemporal dementia (FTD), Alzheimer’s disease (AD), and Parkinson’s disorder (PD) [25][26][27][28]. The timely dissolution of pathological amyloid fibrils may be dependent on cellular levels of ATP, which has recently been identified as a biological hydrotrope [29]—an amphiphilic molecule that may behave as a surfactant [30] which can reduce tension between solute and solvent, and increase solubility in an energy-independent manner.

2. ATP Regulates Biomolecular Condensates

At micromolar concentrations in cells, the hydrolysis of ATP phosphoanhydride bonds provides substantial free energy to fuel chemical processes such as post-translational modifications that may maintain fluid phases or facilitate phase separation by generating supersaturation gradients that can induce droplet segregation [13][15][31][32]. At higher physiological concentrations between 2 and 8 mM, ATP becomes a biological hydrotrope that can solubilize proteins to prevent abnormal aggregation and the formation of pathological amyloid fibrils often associated with neurodegenerative disorders such as Alzheimer’s disease (AD) [29]. Recent extensive all-atom molecular dynamics studies showed that at higher millimolar concentrations (150 mM), ATP prevented the aggregation of amyloid-beta peptide Aβ16−22 and disrupted prefibril formations [33], supporting earlier observations of decreased ATP levels in the brain and whole blood of AD transgenic mouse models [34]. Other experimental studies determined that mechanisms such as the suppressed fibrillation of disordered protein by the adenosine moiety of ATP leading to increased protein stability and reduced thermal aggregation may not be typical of hydrotrope-type reactions. Instead, ATP could be viewed as a kosmotropic anion [35] that can increase the solubility of the hydrophobic adenine part [36]; thus, the term “biological aggregation inhibitor” may be more appropriate [37].
Even though ATP is produced mainly in mitochondria, ATP levels in the mitochondrial matrix are significantly lower than those found in the cytoplasm and nucleus [38][39]. Voltage-dependent anion channels (VDACs) located in the mitochondrial outer membrane (MOM) [40] and adenine nucleotide translocators (ANTs) on the inner mitochondrial membranes (IMM) [41][42] facilitate the export of ATP into cytosol where ATP accumulation has been observed to be the highest [43]. The high physiological concentration of ATP in cytoplasm may be used to control the pathological aggregation of macromolecules that coacervate as a result of transient interactions during LLPS in the cytoplasm and nucleus [44][45]. A major hallmark of ALS/FTD is the presence of FUS inclusion in the cytoplasm. FUS are prosurvivor molecules that re-localize from the nucleus to cytoplasm under stress conditions to form reversible, survival-promoting stress granules via LLPS [46][47]. Stress granules contain important ATP-dependent RNA helicases that function as ATPases to hydrolyze ATP during assembly and disassembly [48]. Stress granules could not be formed without the presence of ATP, and the presence of ATP was required to maintain the liquid-like behavior of assembled droplets [31]. A recent in vitro study showed that aggregate disassembly is also an ATP-dependent process.
During metabolic stress such as nutrient deprivation that causes ATP depletion, cells compartmentalize and sequester misfolded proteins into stress granules to protect cellular fitness. Budding yeast subjected to 0.02% glucose starvation showed a 5-fold ATP decline to ~1.1 mM within 10 min, accompanied by a ~4.4-fold increase in median aggregate diameter, whereas the addition of glucose restored ATP levels, quickly reducing aggregate size and abundance back to control values [49]. Mutants with abolished ATP hydrolysis failed to dissolve aggregates even when placed back in 2% glucose solutions after starvation [49]. In the same manner, ATP has been shown to enhance the LLPS of FUS at low concentrations but dissolves FUS aggregates at higher concentrations [50]. Moreover, 8 mM of ATP complexed with Mg2+ ions prevented the LLPS of FUS and dissolved previously formed FUS condensates [29]. The presence of ATP facilitates the essential phase transition of FUS into stress granule droplets, yet prevents further transition into irreversible aggregation and the fibrillation of FUS to cause cytotoxicity by binding to the RNA-recognition motif (RRM) domain of FUS, kinetically inhibiting the fibrillization of FUS [51]. Similarly, through binding to arginine-containing domains in TDP-43, ATP altered physicochemical properties to induce LLPS, causing droplet formation at molar ratios as low as 1:100 (protein to ATP); by contrast, increasing ATP concentrations could reduce droplet formation, with TDP-43 droplets completely dissolving at a molar ratio of 1:1000 [52]. Nevertheless, in order to completely dissolve the amyloid-beta peptide Aβ-42 associated with AD, supraphysiological concentrations of ATP in excess of 100 mM were found to be necessary [29].
Tau is the major constituent of fibrillar tangles in AD. Phase-separated tau forms droplets that serve as intermediates toward aggregation [28]. Physiological concentrations of ATP at 0.1–10 mM enhanced the fibrillation of 10 μM tau K18 (equivalent to 10–1000-fold molar ratio) by accelerating aggregation in a concentration-dependent manner [53] through energy-independent binding to tau proteins [54]. It may seem paradoxical that ATP would enhance the formation of amyloids and prions that are associated with diseases. As a matter of fact, prion-like mechanisms are functional biological processes ubiquitously present from bacteria to humans [55]. The nucleation and growth of amyloid fibrils in FUS, TDP-43, tau and α-synuclein are dependent upon intermolecular interactions of intrinsically disordered regions (IDRs) and proteins (IDPs) such as prion-like domains and low-complexity sequence domains [56].
Proteins that undergo LLPS often contain long segments that are intrinsically disordered and lack well-defined three-dimensional structure [57]. The relatively low concentration of hydrophobic amino acids in IDPs enables the rapid exchange between multiple conformations where condensates form without altering the affinity of binding interactions during LLPS [58][59][60]. Although the formation of biomolecular condensates can potentially accelerate amyloid aggregation, condensates can also inhibit fibril formation by the sequestration of aggregation-prone, prion-like IDPs. Biomolecular condensates derived from proteins associated with the formation of processing bodies (P-bodies) prevented aberrant amyloid aggregation despite local increase in concentration of aggregate-prone proteins [61]. P-bodies are conserved eukaryotic cytoplasmic ribonucleoprotein (RNP) membraneless organelles that regulate protein homeostasis in non-stressed cells through LLPS involving messenger RNAs (mRNAs) and low-complexity sequence domains [62][63][64][65]. P-bodies respond to cellular stress, especially DNA replication stress, by increasing their sizes and numbers [66][67]. The disassembly of P-bodies in yeast is an ATP-dependent process involving ATP hydrolysis by DEAD-box ATPases [68]. Inhibition of DDX ATPase activity can disrupt the disassembly of physiological MLOs such as P-bodies and stress granules [68][69]. To remain in functional states, bimolecular condensates may require energy to support the continuous active restructuring and rearrangement of molecular components. Insufficient or the depletion of ATP can directly impact the physical and functional properties of biomolecular condensates [31][32][70][71].

2.1. Dimerized ATP Synthase/ATPase Require High-Curvature Lipid Domains

First isolated in 1960 [72][73], F1F0 ATP synthases are mostly localized in the inner membrane invaginations of mitochondria [74]. Eukaryotes and prokaryotes use four major types of ATPases localized in cell membranes to release energy during hydrolysis of ATP for the maintenance of critical transmembrane ionic electrochemical potential differences [75]. In the ubiquitous intracellular powerhouses of eukaryotes, F1F0 ATP synthase is complex V of the electron transport chain responsible for chemiosmotic oxidative phosphorylation (OXPHOS) that couples ATP synthesis to the inner membrane proton gradient [76][77][78]. The ATP synthases of mammalian mitochondria are usually arranged in rows of dimeric complexes of two identical monomers located at the highly curved apex of deep IMM invaginations known as cristae [79]. Dimerized ATP synthases are seven times more active than monomers [80]. Dimerization of ATP synthase may be a major determinant in cristae formation [81], because extreme cristae membrane curvature is shaped by the self-assembly of ATP monomers into dimerized rows [82]. Inability to form dimers resulted in reduced or deformed cristae invaginations [83] that impacted ATP production from decreased OXPHOS activity as a result of defective cristae morphology [84][85]. Experimentally purified ATP synthase reconstituted with membrane lipids revealed that dimerized rows of ATP synthases were formed only on curved surfaces and not on flat membrane areas [86]. Extracellular F1F0 ATP synthases have been observed to translocate from mitochondria to lipid raft domains of various cell types, including plasma membranes of gonadotropes [87], and the sarcolemma of muscle fibers [88].

2.2. Translocation of ATP Dimers to Lipid Rafts Are Cellular Responses to Stress and Stimuli

Biomolecular condensates adapt to changing endogenous or exogenous conditions [3] by continuously fine-tuning biochemical reactions, enriching or excluding biomolecules from their environment [7]. The rapid translocation of mitochondrial ATP synthase to lipid rafts may be integral to these adaptive responses because ATP functions not only as a biological hydrotrope [29][89], increasing the solubility of positively charged, intrinsically disordered proteins [90], but may act as a universal and specific regulator of intrinsically disordered regions (IDRs) capable of altering physicochemical properties, conformation dynamics, assembly, and aggregation [44], in addition to providing phosphates as an energy source to fuel post-translational modifications that regulate the fluctuation of biomolecule phase separation during condensate formation [70][89]. Failure to maintain nanoscopic lipid raft domains with appropriate line tension and membrane elasticity [91] to functionally host dimerized ATPase [92], ATP synthase [86] may contribute to aberrant phase separation, resulting in pathogenic protein aggregates in neurodegeneration [11] and cancer [10][12].

The ability of ATP synthase/ATPase to form dimerized rows on the IMM of mitochondria and other membrane surfaces may be highly dependent upon membrane lipid composition [93] and curvature [86]. Uncontrolled, excess oxidative stress can cause lipid peroxidation [94] which induces pathological changes to membrane lipid composition, including alterations of cardiolipin in IMMs [93][95], as well as changes in membrane curvature that prevent optimal dimerization and the subsequent functioning of ATP synthase/ATPase [96][97]. Insufficient or depletion of ATP can directly impact the physical and functional properties of biomolecular condensates [31][32][70][71]. ATP is not only a biological hydrotrope capable of inhibiting protein LLPS and aggregation at high mM concentrations; it has recently been observed to act as a universal and specific regulator of IDRs, altering their physicochemical properties, conformation dynamics, assembly, and aggregation [44].

3. Melatonin Is a Potent Ancient Antioxidant That Protects ATP Levels to Regulate the Formation and Dissolution of MLOs

Melatonin (N-acetyl-5-methoxytryptamine) is a mitochondria-targeted molecule found in cells of all tested eukarya and bacteria [98]. Effective distribution via horizontal gene transfers may explain the discovery of ancient homologs of arylalkylamine N-acetyltransferase (AANAT), the enzyme responsible for the rhythmic production and release of melatonin in bacteria, fungi, unicellular green algae, and chordates [99][100][101]. In present-day vertebrates, it is estimated that ~99% of melatonin is likely not produced in the pineal gland, nor released into circulation upon pineal production [102], but is mainly synthesized and localized in mitochondria [103][104]. Photosynthetic cyanobacteria responsible for filling the earth with oxygen that led to the extinction of obligate anaerobes produce melatonin [105][106]. The presence of melatonin in primitive unicellular organisms including Rhodospirillum rubrum and cyanobacteria, precursors to mitochondria and chloroplasts, respectively [99][107][108][109], may have conferred protection against endogenous and exogenous oxidative stress that could readily damage biomolecules and disrupt ATP production at plasma membranes [105][109][110][111]. This unique feature implies that melatonin may have an intrinsic modulatory effect over phase separation in early organisms.
As in all eukaryotic cells of plants and animals, LLPS is also believed to be the organizing principle behind the subcellular compartmentalization of membraneless organelles (MLOs) in prokaryotic bacteria [112][113], where condensate formation is tightly correlated with ATP levels. Impaired ATP hydrolysis from reduced ATPase activity in bacteria causes droplet formation by phase separation [114][115]. Cyanobacteria, the only known prokaryote capable of water oxidation [116], has recently been shown to exhibit circadian rhythm in the formation and dissolution of MLOs that remained soluble during daylight, but became reversible, insoluble condensates at night. The formation of aggregates allows cyanobacteria to conserve energy when metabolic activities and ATP levels are lowered at night [117][118][119][120]. It is therefore not unexpected that when ATP production was disrupted, insoluble aggregates could be induced to form in cyanobacteria even during daylight by suppressing F1F0-ATP synthase or uncoupling OXPHOS with mitochondrial proton gradient inhibitors [117].
The gene sequences of cyanobacteria ATP synthase subunits are extremely similar to those in chloroplasts [121]. Embedded in the thylakoid membrane, both ATP synthase in cyanobacteria and chloroplasts (CF0CF1) control transmembrane electrochemical proton gradients for the production of ATP [122][123][124]. Similar to CL, which is synthesized from phosphatidylglycerol (PG) in all organisms [125], PG is the primary phospholipid associated with photosystem complexes that carry out electron transport reactions during oxygenic photosynthesis [126]. Both CL and PG are essential for maintaining the proper lipid composition that supports electron transport and ATP production in eukarya and prokarya, although these lipids are easily subjected to damage via lipid peroxidation [95][127][128][129][130][131][132]. The antioxidant effects of melatonin and its metabolites become particularly meaningful when the prevention of CL peroxidation by hydroperoxyl in mitochondrial membranes can affect the formation and dissolution of biomolecular condensates (Figure 1).
Figure 1. Schematic illustrating the regulation of biomolecular condensates by melatonin represented through observations reported in antioxidant protection against lipid peroxidation to maintain membrane/lipid raft composition/stability that serves to maintain adequate ATP levels in all cellular compartments to fuel, support, and regulate post-translational/m6A modifications that may fine-tune RNA dynamics in the assembly and disassembly of MLOs to prevent pathological aggregations in neurodegenerative disorders. LLPS: liquid–liquid phase separation; DDX: Dead-box RNA helicase; m6A: N6-methyladenosine; MLO: membraneless organelle; RBP: RNA-binding protein; RNP: ribonucleoprotein; PTM: post-translational modification (See Abbreviations for additional acronyms).

3.1. Melatonin Metabolite 3-OHM Inhibits Lipid Peroxidation by Hydroperoxyl Radical

Melatonin and its secondary, tertiary, and quaternary metabolites actively scavenge potent free radicals [133][110][134] including hydroxyl radicals [135], singlet oxygen [136][137], hydrogen peroxide [138], nitric oxide [139][140][141], and peroxynitrite anions [142] via different antioxidant mechanisms such as direct radical trapping in Type I antioxidant reactions and inactivating hydroxyl radicals (OH) through the sequestration of metal ions and deactivating OH during Fenton-like reactions in Type II antioxidant reactions [143]. In addition, melatonin and its metabolites collectively preserve the chemical integrity of biomolecules from oxidative stress via Type III antioxidant cellular repair processes and Type IV antioxidant reactions that can enhance antioxidant enzymes and inhibit pro-oxidant enzymes [143].
A recent study that analyzed the mechanistic interactions between melatonin and OH employing density functional theory found that one molecule of melatonin effectively scavenged two OH radicals to produce the stable footprint metabolite, cyclic 3-hydroxymelatonin (3-OHM) [144], in perfect agreement with mechanisms reported in prior experimental and theoretical studies [135][145][146][147]. 3-OHM has been shown to react with hydroperoxyl radicals (OOH) at rates 98.4 times faster than Trolox in aqueous solution [146]. Trolox is a water-soluble, cell-permeable analog of vitamin E with high radical scavenging potential often used as a yardstick for measuring antioxidant capacities in vitro. Trolox resides mainly in the aqueous phase; therefore, it has been observed that Trolox and other water-soluble antioxidants exhibit reduced scavenging activity if radicals are produced within hydrophobic cores of lipid membranes [148]. Melatonin accumulates in all of the internal membranes of cells as well as other hydrophobic sites [149]; therefore, this antioxidant may be uniquely positioned for quenching lipid peroxidation by OOH and other free radicals that penetrate deep into lipid molecules.

3.2. Melatonin Is Preferentially Located at Hydrophilic/Hydrophobic Membrane Interfaces

All biological cell membranes comprise amphipathic lipid molecules with hydrophilic heads and hydrophobic tails that naturally form bilayers with headgroups oriented towards an aqueous environment and tails facing each other [150]. The melatonin molecule is uncharged in the entire pH range [151] and, accordingly, in laboratory environment, the “hydrophobic” molecule dissolved poorly in water [152] except when solubilized in pure aqueous medium by specific methodology that polarizes the pyrrole ring to facilitate hydrogen bonding of the N–H group [153]. The unique ability to form strong H-bonds with hydrophilic lipid headgroups allowed nonpolar melatonin to be preferentially located at hydrophilic/hydrophobic interfaces, with complete solubility observed at the interfaces between polar and lipophilic nanodomains in reversed micelles [154]. The presence of both hydrophilic and lipophilic moieties in melatonin facilitates the scavenging of both aqueous and lipophilic free radicals [155], especially OH [135] and OOH, the two most prevalent ROS responsible for the chain oxidation of unsaturated phospholipids [156][157] in the membranes of cells and mitochondria [158][159].

3.3. Melatonin Metabolite Free Radical Scavenging Cascades Rescue Cardiolipin from Hydroperoxyl Radicals (OOH)

Lipid peroxidation, a physiological process in all aerobic cells [160], is a cascading chain reaction that begins with the abstraction of allylic hydrogen from adjacent lipid molecules by free radicals such as OOH and OH and terminates with reactive aldehyde end products such as malondialdehyde (MDA) and 4-hydroxynonenal (HNE) [94][161][162][163][164]. Both OOH and OH are derived from ubiquitous superoxide radicals (O2•−) generated from the one-electron reduction of oxygen (O2) that may be catalyzed by nicotinamide adenine dinucleotide phosphate oxidase (NADPH oxidase) during respiratory bursts [165] and/or electron leakage during mitochondrial electron transport [166]. Due to its low rate constant values below ~102 L·mol−1·s−1 [167], O2•− behaves more similarly to an unimpressive reductant (E°′(O2/O2•−) = −0.33 V) than an oxidant (E°′(O2•−/H2O2) = 0.93 V) [163][168][169][170] which reacts at a much slower pace with the tested phospholipids compared to OOH [157][171]. Hydroperoxyl (OOH or HO2), also known as a perhydroxyl radical, is a chemically active, protonated form of superoxide radicals (O2•−) [172], engaged predominantly as intermediates for the disproportionation of O2•− into hydrogen peroxide (H2O2) which then can further be transformed via Fenton’s/Haber–Weiss reactions [173] into OH, possibly the most reactive and mobile species of oxygen that interacts with almost all molecules in cells [94][173]. Even though at neutral pH OOH exists primarily as the less reactive O2•−, where the ratio of protonated OOH to anionic O2•− is ~130:1 (less than 1%), OOH can be a potent initiator of lipid peroxidation [156][157].
When reacting with phospholipids, the advantageous free energy profile of −8.5 kJ/mol free energy minimum relative to the aqueous phase allowed OOH to accumulate at lipid headgroup membrane–water interface at concentration enhancement of over one order of magnitude [174]. Multi-level atomistic simulations for interactions of OH, OOH, and H2O2 with polar headgroups of phospholipid bilayer revealed that all three species traveled deep into the water layer to reach phospholipid biomolecules, oxidizing hydrophilic headgroups before hydrophobic tails [175], with OOH staying adsorbed for the longest duration at headgroup regions [174]. The headgroup of CL is fully ionized as a dianion in the physiological pH range [176], supporting its unique, optimal functionality as a “proton trap” that promotes mitochondrial respiratory enzyme activities [177].
The strong negative curvature of cristae in the IMM is primarily sustained by the distinct molecular geometry of CL with its smaller, elongated, conical-shaped, double-phosphate dianonic headgroups that increase lateral pressure within the acyl chain regions and stabilize cylindrically curved, tubular cristae structures [178][179][180]. In large unilamellar vesicles (LUVs) comprising similar lipid properties as the IMM, the addition of a typical concentration of 25% negatively charged, dianonic CL lowered pH at the membrane interface to ~3.9, compared to the bulk pH of 6.8 normally found in mitochondrial intermembrane space [181] and 7.7 in the matrix space [182]; in contrast, LUVs with mono-anionic lipids only reduced the pH to ~5.3 at the membrane interface [181]. The reduced pH at the membrane interface from CL, linearly associated with increased proton (H+) concentration (~700 to ~800) [181], is the reason why ATP production is doubled in mitochondrial models with cristae compared to those without [183]. At the same time, the increased H+ concentration at membrane surfaces may cause accumulation of OOH, the protonated form of O2•− [172].
OOH remains adsorbed at polar headgroups longer than other ROS tested [174]; therefore, a low pH at membrane interface that is favorable for enhanced ATP synthesis could also initiate peroxidation cascades. As such, even though the proper functioning of CL is prerequisite for optimal mitochondrial respiration and ATP production, peroxidation of CL in mitochondria is an inevitable, natural, physiological process that can deteriorate pathologically [184][185][186][187][188][189][190][191][192][193][194][195][196] unless properly counterbalanced by the continuous synthesis [104] and/or uptake of high levels of melatonin. Melatonin is known for its role in maintaining systemic energy homeostasis [197]. In the mitochondria of brown and beige adipose tissue, CL biosynthesis is robustly induced upon cold exposure [198][199] because CL can bind tightly to uncoupling protein 1 (UCP1), stabilizing its conformation and enhancing functionality [200]. The ability of melatonin to protect CL from peroxidation may account for the increased thermogenic response in Zücker diabetic fatty (ZDF) rats via the restoration of UCP1 mRNA expression, increased mitochondrial mass and brown adipose tissue (BAT) weight, as well as enhanced mitochondrial OXPHOS activities in complex I and IV [201].

3.4. Melatonin Antioxidant Cascades May Inhibit NLRP3 Prionoid-Like Aggregation in an ATP-Dependent Manner

Cardiolipin (CL) is a mitochondria signature lipid distinctly attracted to membrane lipid domains with strong negative curvatures, such as the apex of IMM cristae [202][203]. CL is often externalized to the outer mitochondrial membrane (OMM) upon mitochondrial distress from ROS attacks [204][205], whereas oxidized CL in OMM initiates apoptotic signaling processes [206] that can lead to opening of the mitochondrial permeability transition pore (mPTP) and the release of cytochrome c (Cyt c) [207][208]. Externalized CL, whether oxidized or not, becomes an essential signaling platform that binds and interacts with important mitophagic, autophagic, and inflammatory enzymes [205][209], including Beclin 1 [210], tBid, Bax [208][211], caspase-8 [212], and the NLR pyrin domain containing 3 (NLRP3) inflammasomes [213]. A major source of extremely inflammatory cytokines IL-1β and IL-18 [214], NLRP3 inflammasome is a phase-separated supramolecular complex that mediates immune responses upon the detection of cellular stress and dysfunction [215][216][217]. The activation of the NLRP3 inflammasome in macrophages is induced by oxidized phospholipids [218], whereas the docking of externalized CL to NLRP3 inflammasome primes its assembly and subsequent activation in mitochondria [213] as well as mitochondria-associated membranes (MAMs), a region comprising highly specialized proteins which is tethered to the endoplasmic reticulum (ER) [219][220].  

Melatonin is a potent antioxidant that has been shown to inhibit CL peroxidation in mitochondria, preventing mPTP opening and Cyt c release [221] by inhibiting peroxidation cascades initiated by specific ROS that accumulate in lipid headgroups at membrane–water interfaces [174] (Figure 1). The suppression of oxidative stress and lipid peroxidation may halt the externalization or oxidation of CL, effectively preventing potential pathological interactions with MLOs such as α-syn and the NLRP3 inflammasome. The interaction between pathological α-syn oligomers and externalized CL can result in increased ROS, lipid peroxidation, and mitochondrial dysfunction; therefore, it is not surprising that melatonin has been demonstrated to block α-syn fibril formation and oligomerization, decreasing cytotoxicity in primary neuronal cells [222], as well as rescuing impaired mitochondrial respiration induced by α-syn in Saccharomyces cerevisiae under ROS attack [223]. The NLRP3 inflammasome must be primed by externalized CL upon ROS stimulation before activation [204][213][219]. The regulation of the next phase where the NLRP3 inflammasome transitions into stable, prionoid-like complexes is mediated by DDX3X, one of the ATP-bound forms of DEAD-box RNA helicases responsible for the scaffolding of prionoid, self-oligomerizing specks known as apoptosis-associated speck-like protein containing a C-terminal caspase recruitment domain (ASC) which cannot be easily disassembled once they are formed [224][225][226] (Figure 2).
Figure 2. Overview of melatonin regulation of NLRP3 inflammasome (NLRP3) formation, assembly and activation: (A) Summary of melatonin and metabolite antioxidant cascade inhibiting the initiation and propagation of cardiolipin (CL) peroxidation, effectively terminating the CL peroxidation cascade; (B) Oxidized CL is externalized from the cristae/inner mitochondrial membrane (IMM) to the outer mitochondrial membrane (OMM) where it docks and primes NLRP3 inflammasome assembly prior to activation in mitochondria; (C) DDX3X, an ATP-dependent DEAD-box RNA helicase, is the mediator that selects the formation of “Pro-Survival” stress granules or the transition of the NLRP3 inflammasome into “Pro-Death”, stable, prionoid-like complexes. The successful formation of stress granules is also dependent upon the availability of ATP and RNA, both of which may be regulated by melatonin (See Abbreviations for additional acronyms).

ATP-dependent DEAD-box RNA helicases (DDXs) are ATPases that post-translationally regulate RNA-containing phase-separated organelles in prokaryotes and eukaryotes [227][228]. DDXs promote phase separation in their ATP-bound form, but can also release RNA and induce compartment turnover using ATP hydrolysis. Inhibition of DDX ATPase activity can disrupt the disassembly of physiological MLOs such as P-bodies and stress granules [68][69] (Figure 1). Phosphorylation is one of the most important PTMs that can control the assembly/disassembly of MLOs [229] as well as stabilize or destabilize MLOs including G bodies [230] and p53 [231]. Cells rely on phosphorylation as rapid, reversible responses to different stimuli by changing the physicochemical properties of proteins during phase separation multivalent interactions [70][232]. Phosphorylation establishes covalent bonds between phosphoryl and amino acid hydroxyl groups using the terminal phosphate group in ATP [233]. The ATP-dependent DEAD-box helicase [227] DDX3X responsible for initiating NLRP3 inflammasome aggregation is dependent upon phosphorylation-associated IFN promoter stimulation [224][234][235][236]. When the conserved, eukaryotic, integrated stress response (ISR) pathway is activated by external stress stimuli including hypoxia, nutrient deprivation, viral infections, as well as intrinsic ER stress [237], the phosphorylation of eukaryotic translation initiation factor 2 alpha (eIF2a) on Ser51 [238][239] triggers the formation of stress granules as adaptive homeostatic responses to promote survival and restore homeostasis [240][241][242][243].  It is presently unknown what prompts DDX3X to select the aggregation of pro-survival stress granules over pro-death NLRP3 inflammasomes or vice versa [224][234]. It would not be unreasonable to assume that an excessive oxidative local environment with pathological, enlarged lipid rafts (inflammarafts) [244][245] in membranes could exert a decisive influence over the selection process (Figure 2). 

The activation of the NLRP3 inflammasome is now associated with major neurodegenerative disorders such as AD, PD and ALS, where positive correlations have been found to exist between NLRP3 levels and abnormal protein aggregations such as Aβ and α-Syn, whereas the inhibition of the NLRP3 pathway attenuates pathological protein aggregations [246]. Melatonin inhibited NLRP3 inflammasome activation and reduced the aggregation of ASC specks in the mice hippocampus with major depressive disorder induced by inflammatory liposaccharides [247]; melatonin also inhibited the formation of hypoxia-induced inflammasome protein complexes and reduced the aggregation of ASC specks in macrophages of Sugen/hypoxia pulmonary arterial hypertension (PAH) mouse models [248]. Melatonin attenuated the progression of intervertebral disc degeneration in vitro and in vivo by reducing mitochondrial ROS products to inhibit NLRP3 inflammasome priming and activation, effectively terminating pro-inflammatory cytokine expression [249]. The ability of melatonin to prevent the opening of mPTP and release of Cyt c [221], inhibit NLRP3 inflammasome priming, activation, and ASC speck aggregation [247][248], block α-syn fibrillation [222], and improve mitochondrial respiration [223] could be directly related to its ability to stabilize nanoscopic lipid raft domains and suppress lipid peroxidation, which can alter the composition and molecular structures of lipid rafts.
During lipid peroxidation events, oxidized moieties were found to mainly reside close to the lipid headgroups forming hydrogen bonds with water. These oxidized lipids can perturb membrane bilayer structures and modify membrane properties, including decreasing the membrane fluidity [250][251][252][253]. The preferential location of melatonin in bilayer lipid headgroups allows dynamic interactions that lead to reductions in bilayer thickness and increased bilayer fluidity [254][255][256]. Eukaryotes and prokaryotes use ATPases localized in cell membranes and lipid raft domains to produce and release ATP energy [75][257][258][259]; therefore, increased ATPase activities from enhanced membrane fluidity [260][261] can impact how ATP interacts with phospholipids in bilayers [262] and modulate the LLPS of MLOs formed at membrane surfaces [44]. Moreover, lipid peroxidation is believed to be associated with the reduction in mitochondrial membrane fluidity during aging in animals [263]. Membranes themselves can affect local protein concentrations [264] where high-curvature lipids that form rafts may attract specific proteins that form aggregates to further enhance membrane curvature [265][266][267][268]. Increasingly, neurodegenerative diseases such as AD are viewed as membrane disorders [269]. The size of MLOs that aggregate at membrane surfaces can be tuned through PTMs such as phosphorylation, which is ATP-dependent [270]. The amount of ATP available at membrane surfaces and cytosol drives the formation, tuning, and dissolution of MLOs, and is regulated by oxidative-stress-sensitive ion channels that reside in lipid rafts (Figure 1).

3.5. Melatonin Maintains a High Cytosolic ATP:ADP Ratio through the Optimization of VDAC-CYB5R3 Redox Complexes in Lipid Rafts

Lipid rafts are phase-separated regions in lipid bilayers responsible for important biological functions including signal transduction [271][272] as well as the trafficking and sorting of proteins and lipids [273][274]. The fact that lipid rafts are also important redox signaling platforms that assemble, recruit, and activate redox regulatory multiprotein complex NADPH oxidase [275][276], and host the quintessential plasma membrane redox enzyme complex VDAC-CYB5R3 [277][278], emphasizes the relevance of melatonin as an antioxidant in the protection and stabilization of lipid raft domains.

Nanoscopic transient lipid raft domains in biological membranes are formed by phase separation in response to external stimuli [271][272][279]. Even though cells may alter lipid constituents to control the composition and size of lipid rafts [280], the propagation of molecular stress, lipid raft rattling dynamics and relaxation are some of the basic mechanisms underlying phase separation on the molecular level [281]. The presence of hydrophobic molecules such as melatonin can modulate viscoelastic dynamics through the accumulation and propagation of stress in lipid–lipid interactions [281][282]. Adding melatonin to membrane models led to a breakdown of out-of-phase membrane displacement patterns and the disruption of the vibrational landing platform of lipid biomolecules at the water–membrane interface, effectively slowing the permeation of ROS and other small molecules [281][133].

In 2005, melatonin was first observed to induce phase-separation in DPPC lipid bilayers [250]; recently, melatonin has been observed to modify lipid hydrocarbon chain order to promote phase separation in ternary membrane models [283]. Due to a preference to localize at membrane interfaces [154], melatonin can form strong hydrogen bonds with membrane lipid anionic headgroups that could significantly modulate lipid acyl chain flexibility and lipid dynamics [250]. Melatonin is able to directly interact with cholesterol [284] and displaced cholesterol due to competitive binding to lipid molecules, increasing disorder in the Ld phase to drive cholesterol into the ordered Lo phase [283]. These subtle changes in lipid nanodomains can profoundly affect amyloid processing at membrane sites. Aβ1–40 and Aβ1–42 peptides are known to interact strongly with negatively charged lipids by binding to anionic, negatively charged membranes [285][286][287][288][289]. Increasing cholesterol content lowered the surface charge of lipid membranes in saline solution from positive to negative [290]. Although cholesterol is an indispensable constituent of lipid rafts [271][291], its electrostatic properties altered interactions of charged or polar biomolecules on lipid membrane surfaces and attracted the targeted binding of Aβ deposits at lipid membranes [292][293][294][295].

Local variations in melatonin concentration also affected the re-ordering of lipids in membranes. At 0.5 mol% concentration, melatonin was documented to penetrate lipid bilayers to form fluid domains that enriched lipid membranes where melatonin molecules aligned parallel to phospholipid tails with the electron-dense regions slightly below hydrophilic headgroups; however, at 30 mol% concentration, melatonin molecules aligned parallel to the lipid bilayer, close to the headgroup regions where one melatonin molecule was associated with two lipid molecules to form an ordered, uniform, lateral membrane structure distributed evenly throughout the membrane model [255]. Variations in local concentration and conformational changes in melatonin molecules can directly impact the lipid phase transition, line tension, size, health, and functions of lipid rafts.

Present in all eukaryotes [296], CYB5R3 encodes for a NADH-cytochrome b5 reductase 3 flavoprotein that is engaged in the one-electron transfer from NADH to cytochrome b5 or plasma membrane coenzyme Q, producing NAD+ as a result [297][298]. The soluble isoform of CYB5R3 is exclusive to erythrocytes [299], whereas the membrane-bound isoform is anchored to MOM, ER, and plasma membrane lipid rafts [278][300][301]. Importantly, the OMM-bound CYB5R3 enzyme, ubiquitously expressed in all mammalian cells, is functionally attached to the voltage-dependent anion channel 1 (VDAC1), one of the most prevalent proteins located in the OMM [302][303].

Originally known as mitochondrial porin after its identification in yeast (1985) [304] and humans (1989) [305], VDAC was subsequently observed as a resident protein of lipid rafts in the plasma membranes of animal hearts, brains, and lungs [306] from different human cell lines, including epithelial cells, astrocytes, and neurons [307][308]. Aberrant lipid composition in neuronal lipid rafts disturbs physiological VDAC protein interactions that can affect the opening and closing of VDAC channels, resulting in oxidative stress and neuronal impairments prominent in most AD pathologies [307]. The force-from-lipid principle dictates that the opening and closing of membrane embedded channels can be propelled by the mechanical properties of surrounding lipids [309][310][311][312] and their composition. Changes to raft thickness, curvature and elasticity [313] as a result of lipid peroxidation can therefore affect physiological functions of the VDAC and CYB5R3 redox complex.

CYB5R3 enzymes form large redox centers in lipid rafts that enhance mitochondrial respiration rate and ATP production, albeit resulting in increased production of ROS [278][300][301]. Over stimulation and clustering of CYB5R3 induced oxidative stress-mediated apoptosis of cerebellar granule neurons [314]. Independent of respiratory chain activities, the ascorbate-dependent NADH: cytochrome c oxidoreductase oxidation of NADH at CYB5R3 centers in lipid rafts is also a major source of extracellular superoxide [303][315][316][317][318] that can initiate lipid peroxidation. In Wistar rats, the deregulation of CYB5R3 promptly triggers apoptosis due to the overproduction of superoxide anions at neuronal plasma membranes [278][315]. Excess NADH due to CYB5R3 redox dysfunction can close VDAC, suppressing OXPHOS and increasing glycolysis [303][319], whereas the opening of VDAC also elevates ROS from increased OXPHOS activities [40]. As the most abundant protein in the MOM, VDAC is regarded as a dynamic regulator of mitochondrial functions, interacting with over 100 proteins in health and disease [320]. VDAC opening is believed to globally control mitochondrial metabolism and ROS formation, modulating mitochondria and cellular bioenergetics [40][321]. Nevertheless, the question of whether apoptosis is associated with the opening [322] or closure [323][324] of VDAC has been highly debated [325], further emphasizing the important role of this protein in the regulation of cell life and death [320][326].

VDAC is the gatekeeper which controls the export of ATP out of mitochondria into cytosol and the import of essential respiratory substrates such as ADP and Pi into mitochondria [323][327]; therefore, VDAC opening may be instrumental in determining the fate of MLO formation, regulation, and dissolution. ATP is not only a biological hydrotrope capable of inhibiting protein LLPS and aggregation at high mM concentrations, but it has recently been observed to act as a universal and specific regulator of IDRs capable of altering physicochemical properties, conformation dynamics, assembly, and the aggregation of MLOs [44]. Not only is the preservation of lipid raft structure and composition essential for maintaining specific ion channel properties [307], the amount of cytosolic ATP is dependent upon mitochondrial synthesis and the integrity of CL enriched raft-like lipid domains in mitochondria [277][328][329][330].

The mitochondrial electron transport chain is a major ROS-generating site where complex III and mitochondrial glycerol 3-phosphate dehydrogenase can produce large amounts of redox signaling molecules such as superoxide and hydrogen peroxide to the external side of the IMM as well as the matrix [166][331]. Bis-allylic methylenes and abundant double-bonds in CL lipid chains are vulnerable targets of ROS attacks [184][186][332][333]; therefore, the lipid monolayer leaflets facing the crista lumen enriched in CL in mitochondria [203] may be subject to intense peroxidation events. Peroxidized CL could not support mitochondrial OXPHOS enzyme activities [184][334], leading to the depletion of ATP [183] that can potentiate and exacerbate the aggregation of pathological MLOs. 
Melatonin is an ancient, potent antioxidant that protects lipid nanodomains from peroxidation caused by excess oxidative stress. The addition of micromolar concentrations of melatonin to rat heart mitochondria dramatically inhibited CL oxidation by tert-Butylhydroperoxide (t-BuOOH), a peroxidation promoting peroxide, reversing cytochrome c release, matrix swelling, and proton motive force (ΔΨ) collapse in treated cells [221]. The melatonin molecule is uncharged in the entire pH range [151] and contains both hydrophilic and lipophilic moieties that support its easy accumulation in all internal membranes of cells as well as other hydrophobic sites [155][149]. The exogenous supplementation of melatonin in rodents results in dose-dependent increases in all subcellular compartments, with lipid membranes exhibiting 10-fold increases compared to mitochondria [335]. The presence of both hydrophilic and lipophilic moieties in melatonin not only facilitates the efficacious scavenging of both aqueous and lipophilic free radicals [155], but also places the molecule in a unique position during evolution to protect membrane lipids from oxidative damage and potentially regulate MLOs that form at membrane surfaces in an ATP-dependent manner (Figure 1).
Melatonin protects the functionality of the VDAC–CYB5R3 complex by reducing oxidative stress, lowering ROS that may induce lipid peroxidation, which can alter raft composition, thickness, curvature and elasticity [313] that may impact VDAC ion-channel opening/closure according to the force-from-lipid principle [309][310][311][312]. VDAC expressed in the plasma membranes of HT22 mouse hippocampal neuronal cells were quiescent under control conditions with normal ATP and an absence of apoptotic signals. Serum deprivation increased ROS and induced VDAC opening in the plasma membranes of hippocampal HT22 cells, resulting in mitochondrial dysfunction and increased apoptosis and autophagy. HT22 cells pre-loaded with 200 μM melatonin prior to serum deprivation did not exhibit VDAC activities. In the same manner, the addition of 4 mM ATP blocked the activation of VDAC channels [336], with the implication that melatonin was able to maintain optimal VDAC functioning in an ATP-dependent manner. 

3.6. Melatonin May Regulate Glycolytic G Bodies by Increasing ATP

The theoretical maximum of ATP calculated from simultaneous measurements of extracellular acidification and oxygen consumption indicated that OXPHOS ATP production was close to or more than 16 times above glycolysis, at 31.45 ATP/glucose (maximum total yield 33.45) and 2 ATP/glucose, respectively [337]. As early as 2002, melatonin was found to increase mitochondria OXPHOS activity and elevate the production of ATP [338]. Recent experimental and theoretical studies have presented different mechanisms explaining how melatonin may function as a glycolytic, such as stimulating the SIRT3/PDH axis in vitro to reverse the Warburg phenotype in lung cancer cells [339], converting cells to a healthy phenotype by inhibiting hypoxia-inducible factor-1α to encourage OXPHOS over glycolysis induced by hypoxic conditions [340], downregulating pyruvate dehydrogenase kinase (PDK) to increase acetyl CoA synthesis [341][342], or elevating α-ketoglutarate (α-KG) levels in macrophages to promote M2 polarization that favors OXPHOS over glycolysis [343][344].
Interestingly, in Saccharomyces cerevisiae and human hepatocarcinoma cells challenged with hypoxic stress, the non-canonical RNA-binding proteins in glycolytic enzymes have been observed to promote phase separation [345] that facilitate and maintain the assembly of glycolysis enzymes into cytoplasmic, membraneless glycolytic G bodies that increased glycolytic output during hypoxia [230]. Melatonin is able to increase ATP concentration in cells [201][338][339]; therefore, the switch between OXPHOS and glycolysis could possibly be part of the effect where high ATP concentration dissolves MLO aggregations. Molecular dynamics simulation experiments revealed that the propensity for self-aggregation enhanced the role of ATP as a hydrotrope, preferentially binding to polymers to unfold hydrophobic macromolecules and disrupting the aggregation process of hydrophobic assemblies via the introduction of charges to the macromolecules [346]. These results may explain previous observations where a high cytosolic ATP:ADP ratio readily suppressed glycolysis, whereas the closure of VDAC channels resulting in lower ATP:ADP ratios in cytosol activated glycolysis in vitro [347]. Alterations to the glycolytic pathways are often observed during the early stages of neurodegenerative diseases where mitochondrial dysfunction and reduced ATP levels may contribute to protein aggregation [348]. Increasingly, the pathogenic aggregation of MLOs such as stress granules, p53, FUS, TDP-43, and tau exhibiting dysregulated LLPS is believed to play a major part in the development of neurodegeneration and cancer [12][349][350][351].
The relationship between melatonin and ATP is likely an ancient one that might date as far back as ~4 billion years ago when a proposed gene duplication event at ~3.5 Ga involving CP43 and CP47, enzymes unique to photosystem II (PSII), marked the beginning of water oxidation [116]. Regulation of the synthesis and degradation of the evolutionarily conserved PSII D1 reaction center is mediated by post-translational RNA modulations [352][353][354] and the presence of ATP [355] in a light-dependent manner, where synthesis and/or degradation is induced by light but ceased in the dark. Unlike animals [356], melatonin in plants is increased by the presence of light [357][358], and treatment with melatonin enhanced the synthesis of PSII D1 reaction centers in tomato seedlings under salt stress [359]. Cyanobacteria, the only known prokaryote capable of water oxidation [116] which also produces melatonin [105][106], has recently been shown to exhibit circadian rhythm in the formation and dissolution of MLOs that remained soluble during daylight, but became reversible, insoluble condensates at night in an ATP-dependent manner [117]; therefore, it is not unreasonable to hypothesize that the relationship between melatonin, MLOs, and ATP was already in existence at ~3.5 Ga. The presence of melatonin in primitive unicellular organisms including Rhodospirillum rubrum and cyanobacteria, precursors to mitochondria and chloroplasts, respectively [99][107][108][109], may have conferred protection against endogenous and exogenous oxidative stress that could readily damage macromolecules and disrupt ATP production at membrane lipid domains [105][110][111]. This unique feature implies that melatonin may have an intrinsic modulatory effect over phase separation, not only in early but present-day organisms (Figure 1). 

4. Conclusions

The physiological and pathological functions of biomolecular condensates in health and disease may be shaped by powerful, complex, interdependent relationships between membraneless organelles, membranes/lipid rafts, ATP, and most of all, stress and its timely resolution. Melatonin’s intimate association with each of these decisive influencers may position the potent, ancient antioxidant as an important mediator of the phase separation of condensates in health and disease via principal ATP-dependent post-translational mechanisms and regulation of ATP levels in mitochondria and cytoplasm (Figure 1). This novel theoretical review highlights the important connections between melatonin and ATP in the regulation of biomolecular condensates with the intention to spur further research interest and exploration in the full, multi-faceted potential of melatonin that may provide solutions and answers to existing and future challenges and questions in this exciting and promising field of study.


  1. Feng, Z.; Chen, X.; Wu, X.; Zhang, M. Formation of Biological Condensates via Phase Separation: Characteristics, Analytical Methods, and Physiological Implications. J. Biol. Chem. 2019, 294, 14823–14835.
  2. Oparin, A.I.; Synge, A. The Origin of Life on the Earth/Translated from the Russian by Ann Synge; Elsevier Science Ltd.: Amsterdam, The Netherlands, 1957.
  3. Riback, J.A.; Katanski, C.D.; Kear-Scott, J.L.; Pilipenko, E.V.; Rojek, A.E.; Sosnick, T.R.; Drummond, D.A. Stress-Triggered Phase Separation is an Adaptive, Evolutionarily Tuned Response. Cell 2017, 168, 1028–1040.e19.
  4. Li, P.; Banjade, S.; Cheng, H.-C.; Kim, S.; Chen, B.; Guo, L.; Llaguno, M.; Hollingsworth, J.V.; King, D.S.; Banani, S.F.; et al. Phase Transitions in the Assembly of Multivalent Signalling Proteins. Nature 2012, 483, 336–340.
  5. Su, X.; Ditlev, J.A.; Hui, E.; Xing, W.; Banjade, S.; Okrut, J.; King, D.S.; Taunton, J.; Rosen, M.K.; Vale, R.D. Phase Separation of Signaling Molecules Promotes T Cell Receptor Signal Transduction. Science 2016, 352, 595–599.
  6. Laflamme, G.; Mekhail, K. Biomolecular Condensates as Arbiters of Biochemical Reactions inside the Nucleus. Commun. Biol. 2020, 3, 773.
  7. Ditlev, J.A.; Case, L.B.; Rosen, M.K. Who’s In and Who’s Out-Compositional Control of Biomolecular Condensates. J. Mol. Biol. 2018, 430, 4666–4684.
  8. Shin, Y.; Brangwynne, C.P. Liquid Phase Condensation in Cell Physiology and Disease. Science 2017, 357, eaaf4382.
  9. Alberti, S.; Dormann, D. Liquid-Liquid Phase Separation in Disease. Annu. Rev. Genet. 2019, 53, 171–194.
  10. Boija, A.; Klein, I.A.; Young, R.A. Biomolecular Condensates and Cancer. Cancer Cell 2021, 39, 174–192.
  11. Zbinden, A.; Pérez-Berlanga, M.; De Rossi, P.; Polymenidou, M. Phase Separation and Neurodegenerative Diseases: A Disturbance in the Force. Dev. Cell 2020, 55, 45–68.
  12. Taniue, K.; Akimitsu, N. Aberrant Phase Separation and Cancer. FEBS J. 2021.
  13. Hyman, A.A.; Weber, C.A.; Jülicher, F. Liquid-Liquid Phase Separation in Biology. Annu. Rev. Cell Dev. Biol. 2014, 30, 39–58.
  14. Ahlers, J.; Adams, E.M.; Bader, V.; Pezzotti, S.; Winklhofer, K.F.; Tatzelt, J.; Havenith, M. The Key Role of Solvent in Condensation: Mapping Water in Liquid-Liquid Phase-Separated FUS. Biophys. J. 2021, 120, 1266–1275.
  15. Lodish, H.; Berk, A.; Lawrence Zipursky, S.; Matsudaira, P.; Baltimore, D.; Darnell, J. Biochemical Energetics; W. H. Freeman: New York, NY, USA, 2000.
  16. Manchester, K.L. Free Energy ATP Hydrolysis and Phosphorylation Potential. Biochem. Educ. 1980, 8, 70–72.
  17. Peth, A.; Uchiki, T.; Goldberg, A.L. ATP-Dependent Steps in the Binding of Ubiquitin Conjugates to the 26S Proteasome That Commit to Degradation. Mol. Cell 2010, 40, 671–681.
  18. Callis, J. The Ubiquitination Machinery of the Ubiquitin System. Arab. Book 2014, 12, e0174.
  19. Zhao, X. SUMO-Mediated Regulation of Nuclear Functions and Signaling Processes. Mol. Cell 2018, 71, 409–418.
  20. Van Damme, E.; Laukens, K.; Dang, T.H.; Van Ostade, X. A Manually Curated Network of the PML Nuclear Body Interactome Reveals an Important Role for PML-NBs in SUMOylation Dynamics. Int. J. Biol. Sci. 2010, 6, 51–67.
  21. Hardenberg, M.; Horvath, A.; Ambrus, V.; Fuxreiter, M.; Vendruscolo, M. Widespread Occurrence of the Droplet State of Proteins in the Human Proteome. Proc. Natl. Acad. Sci. USA 2020, 117, 33254–33262.
  22. Hondele, M.; Heinrich, S.; De Los Rios, P.; Weis, K. Membraneless Organelles: Phasing out of Equilibrium. Emerg. Top Life Sci. 2020, 4, 331–342.
  23. Garcia-Jove Navarro, M.; Kashida, S.; Chouaib, R.; Souquere, S.; Pierron, G.; Weil, D.; Gueroui, Z. RNA Is a Critical Element for the Sizing and the Composition of Phase-Separated RNA-Protein Condensates. Nat. Commun. 2019, 10, 3230.
  24. Elbaum-Garfinkle, S.; Kim, Y.; Szczepaniak, K.; Chen, C.C.-H.; Eckmann, C.R.; Myong, S.; Brangwynne, C.P. The Disordered P Granule Protein LAF-1 Drives Phase Separation into Droplets with Tunable Viscosity and Dynamics. Proc. Natl. Acad. Sci. USA 2015, 112, 7189–7194.
  25. Patel, A.; Lee, H.O.; Jawerth, L.; Maharana, S.; Jahnel, M.; Hein, M.Y.; Stoynov, S.; Mahamid, J.; Saha, S.; Franzmann, T.M.; et al. A Liquid-to-Solid Phase Transition of the ALS Protein FUS Accelerated by Disease Mutation. Cell 2015, 162, 1066–1077.
  26. Niaki, A.G.; Sarkar, J.; Cai, X.; Rhine, K.; Vidaurre, V.; Guy, B.; Hurst, M.; Lee, J.C.; Koh, H.R.; Guo, L.; et al. Loss of Dynamic RNA Interaction and Aberrant Phase Separation Induced by Two Distinct Types of ALS/FTD-Linked FUS Mutations. Mol. Cell 2020, 77, 82–94.e4.
  27. Mann, J.R.; Gleixner, A.M.; Mauna, J.C.; Gomes, E.; DeChellis-Marks, M.R.; Needham, P.G.; Copley, K.E.; Hurtle, B.; Portz, B.; Pyles, N.J.; et al. RNA Binding Antagonizes Neurotoxic Phase Transitions of TDP-43. Neuron 2019, 102, 321–338.e8.
  28. Wegmann, S.; Eftekharzadeh, B.; Tepper, K.; Zoltowska, K.M.; Bennett, R.E.; Dujardin, S.; Laskowski, P.R.; MacKenzie, D.; Kamath, T.; Commins, C.; et al. Tau Protein Liquid-Liquid Phase Separation Can Initiate Tau Aggregation. EMBO J. 2018, 37, e98049.
  29. Patel, A.; Malinovska, L.; Saha, S.; Wang, J.; Alberti, S.; Krishnan, Y.; Hyman, A.A. ATP as a Biological Hydrotrope. Science 2017, 356, 753–756.
  30. Hatzopoulos, M.H.; Eastoe, J.; Dowding, P.J.; Rogers, S.E.; Heenan, R.; Dyer, R. Are Hydrotropes Distinct from Surfactants? Langmuir 2011, 27, 12346–12353.
  31. Jain, S.; Wheeler, J.R.; Walters, R.W.; Agrawal, A.; Barsic, A.; Parker, R. ATPase-Modulated Stress Granules Contain a Diverse Proteome and Substructure. Cell 2016, 164, 487–498.
  32. Brangwynne, C.P.; Mitchison, T.J.; Hyman, A.A. Active Liquid-like Behavior of Nucleoli Determines Their Size and Shape in Xenopus Laevis Oocytes. Proc. Natl. Acad. Sci. USA 2011, 108, 4334–4339.
  33. Pal, S.; Paul, S. ATP Controls the Aggregation of Aβ16-22 Peptides. J. Phys. Chem. B 2020, 124, 210–223.
  34. Zhang, C.; Rissman, R.A.; Feng, J. Characterization of ATP Alternations in an Alzheimer’s Disease Transgenic Mouse Model. J. Alzheimers. Dis. 2015, 44, 375–378.
  35. Salis, A.; Ninham, B.W. Models and Mechanisms of Hofmeister Effects in Electrolyte Solutions, and Colloid and Protein Systems Revisited. Chem. Soc. Rev. 2014, 43, 7358–7377.
  36. Mandl, I.; Grauer, A.; Neuberg, C. Solubilization of Insoluble Matter in Nature; I. The Part Played by Salts of Adenosinetriphosphate. Biochim. Biophys. Acta 1952, 8, 654–663.
  37. Mehringer, J.; Do, T.-M.; Touraud, D.; Hohenschutz, M.; Khoshsima, A.; Horinek, D.; Kunz, W. Hofmeister versus Neuberg: Is ATP Really a Biological Hydrotrope? Cell Rep. Phys. Science 2021, 2, 100343.
  38. Schwenke, W.D.; Soboll, S.; Seitz, H.J.; Sies, H. Mitochondrial and Cytosolic ATP/ADP Ratios in Rat Liver in Vivo. Biochem. J 1981, 200, 405–408.
  39. Imamura, H.; Nhat, K.P.H.; Togawa, H.; Saito, K.; Iino, R.; Kato-Yamada, Y.; Nagai, T.; Noji, H. Visualization of ATP Levels inside Single Living Cells with Fluorescence Resonance Energy Transfer-Based Genetically Encoded Indicators. Proc. Natl. Acad. Sci. USA 2009, 106, 15651–15656.
  40. Fang, D.; Maldonado, E.N. VDAC Regulation: A Mitochondrial Target to Stop Cell Proliferation. Adv. Cancer Res. 2018, 138, 41–69.
  41. Ruprecht, J.J.; King, M.S.; Zögg, T.; Aleksandrova, A.A.; Pardon, E.; Crichton, P.G.; Steyaert, J.; Kunji, E.R.S. The Molecular Mechanism of Transport by the Mitochondrial ADP/ATP Carrier. Cell 2019, 176, 435–447.e15.
  42. Liu, Y.; Chen, X.J. Adenine Nucleotide Translocase, Mitochondrial Stress, and Degenerative Cell Death. Oxid. Med. Cell. Longev. 2013, 2013, 146860.
  43. Depaoli, M.R.; Karsten, F.; Madreiter-Sokolowski, C.T.; Klec, C.; Gottschalk, B.; Bischof, H.; Eroglu, E.; Waldeck-Weiermair, M.; Simmen, T.; Graier, W.F.; et al. Real-Time Imaging of Mitochondrial ATP Dynamics Reveals the Metabolic Setting of Single Cells. Cell Rep. 2018, 25, 501–512.e3.
  44. Song, J. Adenosine Triphosphate Energy-Independently Controls Protein Homeostasis with Unique Structure and Diverse Mechanisms. Protein Sci. 2021, 30, 1277–1293.
  45. Takaine, M.; Imamura, H.; Yoshida, S. High and Stable ATP Levels Prevent Aberrant Intracellular Protein Aggregation. bioRxiv 2021, 2021, 801738.
  46. Sama, R.R.K.; Ward, C.L.; Kaushansky, L.J.; Lemay, N.; Ishigaki, S.; Urano, F.; Bosco, D.A. FUS/TLS Assembles into Stress Granules and Is a Prosurvival Factor during Hyperosmolar Stress. J. Cell. Physiol. 2013, 228, 2222–2231.
  47. Mahboubi, H.; Stochaj, U. Cytoplasmic Stress Granules: Dynamic Modulators of Cell Signaling and Disease. Biochim. Biophys. Acta Mol. Basis Dis. 2017, 1863, 884–895.
  48. Hilliker, A. Analysis of RNA Helicases in P-Bodies and Stress Granules. Methods Enzymol. 2012, 511, 323–346.
  49. Sathyanarayanan, U.; Musa, M.; Bou Dib, P.; Raimundo, N.; Milosevic, I.; Krisko, A. ATP Hydrolysis by Yeast Hsp104 Determines Protein Aggregate Dissolution and Size in Vivo. Nat. Commun. 2020, 11, 5226.
  50. Kang, J.; Lim, L.; Song, J. ATP Enhances at Low Concentrations but Dissolves at High Concentrations Liquid-Liquid Phase Separation (LLPS) of ALS/FTD-Causing FUS. Biochem. Biophys. Res. Commun. 2018, 504, 545–551.
  51. Kang, J.; Lim, L.; Song, J. ATP Binds and Inhibits the Neurodegeneration-Associated Fibrillization of the FUS RRM Domain. Commun. Biol. 2019, 2, 223.
  52. Dang, M.; Lim, L.; Kang, J.; Song, J. ATP Biphasically Modulates LLPS of TDP-43 PLD by Specifically Binding Arginine Residues. Commun. Biol. 2021, 4, 714.
  53. Heo, C.E.; Han, J.Y.; Lim, S.; Lee, J.; Im, D.; Lee, M.J.; Kim, Y.K.; Kim, H.I. ATP Kinetically Modulates Pathogenic Tau Fibrillations. ACS Chem. Neurosci. 2020, 11, 3144–3152.
  54. Farid, M.; Corbo, C.P.; Alonso, A.D.C. Tau Binds ATP and Induces Its Aggregation. Microsc. Res. Tech. 2014, 77, 133–137.
  55. Newby, G.A.; Lindquist, S. Blessings in Disguise: Biological Benefits of Prion-like Mechanisms. Trends Cell Biol. 2013, 23, 251–259.
  56. Li, L.; McGinnis, J.P.; Si, K. Translational Control by Prion-like Proteins. Trends Cell Biol. 2018, 28, 494–505.
  57. Schuster, B.S.; Dignon, G.L.; Tang, W.S.; Kelley, F.M.; Ranganath, A.K.; Jahnke, C.N.; Simpkins, A.G.; Regy, R.M.; Hammer, D.A.; Good, M.C.; et al. Identifying Sequence Perturbations to an Intrinsically Disordered Protein That Determine Its Phase-Separation Behavior. Proc. Natl. Acad. Sci. USA 2020, 117, 11421–11431.
  58. Harmon, T.S.; Holehouse, A.S.; Rosen, M.K.; Pappu, R.V. Intrinsically Disordered Linkers Determine the Interplay between Phase Separation and Gelation in Multivalent Proteins. Elife 2017, 6, e30294.
  59. Owen, I.; Shewmaker, F. The Role of Post-Translational Modifications in the Phase Transitions of Intrinsically Disordered Proteins. Int. J. Mol. Sci. 2019, 20, 5501.
  60. Van der Lee, R.; Buljan, M.; Lang, B.; Weatheritt, R.J.; Daughdrill, G.W.; Dunker, A.K.; Fuxreiter, M.; Gough, J.; Gsponer, J.; Jones, D.T.; et al. Classification of Intrinsically Disordered Regions and Proteins. Chem. Rev. 2014, 114, 6589–6631.
  61. Küffner, A.M.; Linsenmeier, M.; Grigolato, F.; Prodan, M.; Zuccarini, R.; Capasso Palmiero, U.; Faltova, L.; Arosio, P. Sequestration within Biomolecular Condensates Inhibits Aβ-42 Amyloid Formation. Chem. Sci. 2021, 12, 4373–4382.
  62. Luo, Y.; Na, Z.; Slavoff, S.A. P-Bodies: Composition, Properties, and Functions. Biochemistry 2018, 57, 2424–2431.
  63. Stoecklin, G.; Kedersha, N. Relationship of GW/P-Bodies with Stress Granules. Adv. Exp. Med. Biol. 2013, 768, 197–211.
  64. Kedersha, N.; Stoecklin, G.; Ayodele, M.; Yacono, P.; Lykke-Andersen, J.; Fritzler, M.J.; Scheuner, D.; Kaufman, R.J.; Golan, D.E.; Anderson, P. Stress Granules and Processing Bodies Are Dynamically Linked Sites of mRNP Remodeling. J. Cell Biol. 2005, 169, 871–884.
  65. Nostramo, R.; Xing, S.; Zhang, B.; Herman, P.K. Insights into the Role of P-Bodies and Stress Granules in Protein Quality Control. Genetics 2019, 213, 251–265.
  66. Teixeira, D.; Sheth, U.; Valencia-Sanchez, M.A.; Brengues, M.; Parker, R. Processing Bodies Require RNA for Assembly and Contain Nontranslating mRNAs. RNA 2005, 11, 371–382.
  67. Loll-Krippleber, R.; Brown, G.W. P-Body Proteins Regulate Transcriptional Rewiring to Promote DNA Replication Stress Resistance. Nat. Commun. 2017, 8, 558.
  68. Mugler, C.F.; Hondele, M.; Heinrich, S.; Sachdev, R.; Vallotton, P.; Koek, A.Y.; Chan, L.Y.; Weis, K. ATPase Activity of the DEAD-Box Protein Dhh1 Controls Processing Body Formation. Elife 2016, 5, e18746.
  69. Hondele, M.; Sachdev, R.; Heinrich, S.; Wang, J.; Vallotton, P.; Fontoura, B.M.A.; Weis, K. DEAD-Box ATPases Are Global Regulators of Phase-Separated Organelles. Nature 2019, 573, 144–148.
  70. Snead, W.T.; Gladfelter, A.S. The Control Centers of Biomolecular Phase Separation: How Membrane Surfaces, PTMs, and Active Processes Regulate Condensation. Mol. Cell 2019, 76, 295–305.
  71. Feric, M.; Vaidya, N.; Harmon, T.S.; Mitrea, D.M.; Zhu, L.; Richardson, T.M.; Kriwacki, R.W.; Pappu, R.V.; Brangwynne, C.P. Coexisting Liquid Phases Underlie Nucleolar Subcompartments. Cell 2016, 165, 1686–1697.
  72. Pullman, M.E.; Penefsky, H.S.; Datta, A.; Racker, E. Partial Resolution of the Enzymes Catalyzing Oxidative Phosphorylation. I. Purification and Properties of Soluble Dinitrophenol-Stimulated Adenosine Triphosphatase. J. Biol. Chem. 1960, 235, 3322–3329.
  73. Ernster, L.; Schatz, G. Mitochondria: A Historical Review. J. Cell Biol. 1981, 91, 227s–255s.
  74. Abrahams, J.P.; Leslie, A.G.; Lutter, R.; Walker, J.E. Structure at 2.8 A Resolution of F1-ATPase from Bovine Heart Mitochondria. Nature 1994, 370, 621–628.
  75. Grüber, G.; Wieczorek, H.; Harvey, W.R.; Müller, V. Structure–function Relationships of A-, F- and V-ATPases. J. Exp. Biol. 2001, 204, 2597–2605.
  76. Mitchell, P. Chemiosmotic Coupling in Oxidative and Photosynthetic Phosphorylation. Biol. Rev. Camb. Philos. Soc. 1966, 41, 445–502.
  77. Boyer, P.D. The ATP Synthase--a Splendid Molecular Machine. Annu. Rev. Biochem. 1997, 66, 717–749.
  78. Jonckheere, A.I.; Smeitink, J.A.M.; Rodenburg, R.J.T. Mitochondrial ATP Synthase: Architecture, Function and Pathology. J. Inherit. Metab. Dis. 2012, 35, 211–225.
  79. 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.
  80. 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.
  81. 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, e2021012118.
  82. Paumard, P.; Vaillier, J.; Coulary, B.; Schaeffer, J.; Soubannier, V.; Mueller, D.M.; Brèthes, D.; di Rago, J.-P.; Velours, J. The ATP Synthase Is Involved in Generating Mitochondrial Cristae Morphology. EMBO J. 2002, 21, 221–230.
  83. 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.
  84. Habersetzer, J.; Larrieu, I.; Priault, M.; Salin, B.; Rossignol, R.; Brèthes, D.; Paumard, P. Human F1F0 ATP Synthase, Mitochondrial Ultrastructure and OXPHOS Impairment: A (super-)complex Matter? PLoS ONE 2013, 8, e75429.
  85. Mannella, C.A. The Relevance of Mitochondrial Membrane Topology to Mitochondrial Function. Biochim. Biophys. Acta 2006, 1762, 140–147.
  86. Blum, T.B.; Hahn, A.; Meier, T.; Davies, K.M.; Kühlbrandt, W. Dimers of Mitochondrial ATP Synthase Induce Membrane Curvature and Self-Assemble into Rows. Proc. Natl. Acad. Sci. USA 2019, 116, 4250–4255.
  87. Allen-Worthington, K.; Xie, J.; Brown, J.L.; Edmunson, A.M.; Dowling, A.; Navratil, A.M.; Scavelli, K.; Yoon, H.; Kim, D.-G.; Bynoe, M.S.; et al. The F0F1 ATP Synthase Complex Localizes to Membrane Rafts in Gonadotrope Cells. Mol. Endocrinol. 2016, 30, 996–1011.
  88. Kim, B.-W.; Lee, J.-W.; Choo, H.-J.; Lee, C.S.; Jung, S.-Y.; Yi, J.-S.; Ham, Y.-M.; Lee, J.-H.; Hong, J.; Kang, M.-J.; et al. Mitochondrial Oxidative Phosphorylation System Is Recruited to Detergent-Resistant Lipid Rafts during Myogenesis. Proteomics 2010, 10, 2498–2515.
  89. Hayes, M.H.; Peuchen, E.H.; Dovichi, N.J.; Weeks, D.L. Dual Roles for ATP in the Regulation of Phase Separated Protein Aggregates in Xenopus Oocyte Nucleoli. Elife 2018, 7, e35224.
  90. Sridharan, S.; Kurzawa, N.; Werner, T.; Günthner, I.; Helm, D.; Huber, W.; Bantscheff, M.; Savitski, M.M. Proteome-Wide Solubility and Thermal Stability Profiling Reveals Distinct Regulatory Roles for ATP. Nat. Commun. 2019, 10, 1155.
  91. Kuzmin, P.I.; Akimov, S.A.; Chizmadzhev, Y.A.; Zimmerberg, J.; Cohen, F.S. Line Tension and Interaction Energies of Membrane Rafts Calculated from Lipid Splay and Tilt. Biophys. J. 2005, 88, 1120–1133.
  92. Blackwell, D.J.; Zak, T.J.; Robia, S.L. Cardiac Calcium ATPase Dimerization Measured by Cross-Linking and Fluorescence Energy Transfer. Biophys. J. 2016, 111, 1192–1202.
  93. Houtkooper, R.H.; Vaz, F.M. Cardiolipin, the Heart of Mitochondrial Metabolism. Cell. Mol. Life Sci. 2008, 65, 2493–2506.
  94. 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.
  95. 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.
  96. Sankhagowit, S.; Lee, E.Y.; Wong, G.C.L.; Malmstadt, N. Oxidation of Membrane Curvature-Regulating Phosphatidylethanolamine Lipid Results in Formation of Bilayer and Cubic Structures. Langmuir 2016, 32, 2450–2457.
  97. Acehan, D.; Malhotra, A.; Xu, Y.; Ren, M.; Stokes, D.L.; Schlame, M. Cardiolipin Affects the Supramolecular Organization of ATP Synthase in Mitochondria. Biophys. J. 2011, 100, 2184–2192.
  98. Reiter, R.J.; Rosales-Corral, S.; Tan, D.X.; Jou, M.J.; Galano, A.; Xu, B. Melatonin as a Mitochondria-Targeted Antioxidant: One of Evolution’s Best Ideas. Cell. Mol. Life Sci. 2017, 74, 3863–3881.
  99. Coon, S.L.; Klein, D.C. Evolution of Arylalkylamine N-Acetyltransferase: Emergence and Divergence. Mol. Cell. Endocrinol. 2006, 252, 2–10.
  100. Klein, D.C. Arylalkylamine N-Acetyltransferase: The Timezyme. J. Biol. Chem. 2007, 282, 4233–4237.
  101. Ganguly, S.; Weller, J.L.; Ho, A.; Chemineau, P.; Malpaux, B.; Klein, D.C. Melatonin Synthesis: 14-3-3-Dependent Activation and Inhibition of Arylalkylamine N-Acetyltransferase Mediated by Phosphoserine-205. Proc. Natl. Acad. Sci. USA 2005, 102, 1222–1227.
  102. Martín, M.; Macías, M.; Escames, G.; León, J.; Acuña-Castroviejo, D. Melatonin but Not Vitamins C and E Maintains Glutathione Homeostasis in T-Butyl Hydroperoxide-Induced Mitochondrial Oxidative Stress. FASEB J. 2000, 14, 1677–1679.
  103. Tan, D.-X.; Manchester, L.C.; Qin, L.; Reiter, R.J. Melatonin: A Mitochondrial Targeting Molecule Involving Mitochondrial Protection and Dynamics. Int. J. Mol. Sci. 2016, 17, 2124.
  104. Suofu, Y.; Li, W.; Jean-Alphonse, F.G.; Jia, J.; Khattar, N.K.; Li, J.; Baranov, S.V.; Leronni, D.; Mihalik, A.C.; He, Y.; et al. Dual Role of Mitochondria in Producing Melatonin and Driving GPCR Signaling to Block Cytochrome c Release. Proc. Natl. Acad. Sci. USA 2017, 114, E7997–E8006.
  105. Manchester, L.C.; Coto-Montes, A.; Boga, J.A.; Andersen, L.P.H.; Zhou, Z.; Galano, A.; Vriend, J.; Tan, D.-X.; Reiter, R.J. Melatonin: An Ancient Molecule That Makes Oxygen Metabolically Tolerable. J. Pineal. Res. 2015, 59, 403–419.
  106. Byeon, Y.; Lee, K.; Park, Y.-I.; Park, S.; Back, K. Molecular Cloning and Functional Analysis of Serotonin N-Acetyltransferase from the Cyanobacterium Synechocystis Sp. PCC 6803. J. Pineal. Res. 2013, 55, 371–376.
  107. Tan, D.-X.; Manchester, L.C.; Liu, X.; Rosales-Corral, S.A.; Acuna-Castroviejo, D.; Reiter, R.J. Mitochondria and Chloroplasts as the Original Sites of Melatonin Synthesis: A Hypothesis Related to Melatonin’s Primary Function and Evolution in Eukaryotes. J. Pineal. Res. 2013, 54, 127–138.
  108. Abhishek, A.; Bavishi, A.; Bavishi, A.; Choudhary, M. Bacterial Genome Chimaerism and the Origin of Mitochondria. Can. J. Microbiol. 2011, 57, 49–61.
  109. Raven, J.A.; Allen, J.F. Genomics and Chloroplast Evolution: What Did Cyanobacteria Do for Plants? Genome Biol. 2003, 4, 209.
  110. 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.
  111. Liu, L.-N. Distribution and Dynamics of Electron Transport Complexes in Cyanobacterial Thylakoid Membranes. Biochim. Biophys. Acta 2016, 1857, 256–265.
  112. Gomes, E.; Shorter, J. The Molecular Language of Membraneless Organelles. J. Biol. Chem. 2019, 294, 7115–7127.
  113. Azaldegui, C.A.; Vecchiarelli, A.G.; Biteen, J.S. The Emergence of Phase Separation as an Organizing Principle in Bacteria. Biophys. J. 2021, 120, 1123–1138.
  114. Guilhas, B.; Walter, J.-C.; Rech, J.; David, G.; Walliser, N.O.; Palmeri, J.; Mathieu-Demaziere, C.; Parmeggiani, A.; Bouet, J.-Y.; Le Gall, A.; et al. ATP-Driven Separation of Liquid Phase Condensates in Bacteria. Mol. Cell 2020, 79, 293–303.e4.
  115. Muthunayake, N.S.; Tomares, D.T.; Childers, W.S.; Schrader, J.M. Phase-Separated Bacterial Ribonucleoprotein Bodies Organize mRNA Decay. Wiley Interdiscip. Rev. RNA 2020, 11, e1599.
  116. Oliver, T.; Sánchez-Baracaldo, P.; Larkum, A.W.; Rutherford, A.W.; Cardona, T. Time-Resolved Comparative Molecular Evolution of Oxygenic Photosynthesis. Biochim. Biophys. Acta Bioenerg. 2021, 1862, 148400.
  117. Pattanayak, G.K.; Liao, Y.; Wallace, E.W.J.; Budnik, B.; Drummond, D.A.; Rust, M.J. Daily Cycles of Reversible Protein Condensation in Cyanobacteria. Cell Rep. 2020, 32, 108032.
  118. Bar Eyal, L.; Ranjbar Choubeh, R.; Cohen, E.; Eisenberg, I.; Tamburu, C.; Dorogi, M.; Ünnep, R.; Appavou, M.-S.; Nevo, R.; Raviv, U.; et al. Changes in Aggregation States of Light-Harvesting Complexes as a Mechanism for Modulating Energy Transfer in Desert Crust Cyanobacteria. Proc. Natl. Acad. Sci. USA 2017, 114, 9481–9486.
  119. Wang, H.; Yan, X.; Aigner, H.; Bracher, A.; Nguyen, N.D.; Hee, W.Y.; Long, B.M.; Price, G.D.; Hartl, F.U.; Hayer-Hartl, M. Rubisco Condensate Formation by CcmM in β-Carboxysome Biogenesis. Nature 2019, 566, 131–135.
  120. McKinney, D.W.; Buchanan, B.B.; Wolosiuk, R.A. Activation of Chloroplast ATPase by Reduced Thioredoxin. Phytochemistry 1978, 17, 794–795.
  121. Curtis, S.E. Structure, Organization and Expression of Cyanobacterial ATP Synthase Genes. Photosynth. Res. 1988, 18, 223–244.
  122. Pogoryelov, D.; Reichen, C.; Klyszejko, A.L.; Brunisholz, R.; Muller, D.J.; Dimroth, P.; Meier, T. The Oligomeric State of c Rings from Cyanobacterial F-ATP Synthases Varies from 13 to 15. J. Bacteriol. 2007, 189, 5895–5902.
  123. Walraven, H.S.; Bakels, R.H.A. Function, Structure and Regulation of Cyanobacterial and Chloroplast ATP Synthase. Physiol. Plant 1996, 96, 526–532.
  124. Buchert, F.E. Chapter Three—Chloroplast ATP Synthase from Green Microalgae. In Advances in Botanical Research; Hisabori, T., Ed.; Academic Press: Cambridge, MA, USA, 2020; Volume 96, pp. 75–118.
  125. Carman, G.M. An Unusual Phosphatidylethanolamine-Utilizing Cardiolipin Synthase Is Discovered in Bacteria. Proc. Natl. Acad. Sci. USA 2012, 109, 16402–16403.
  126. Kobayashi, K.; Endo, K.; Wada, H. Specific Distribution of Phosphatidylglycerol to Photosystem Complexes in the Thylakoid Membrane. Front. Plant Sci. 2017, 8, 1991.
  127. Basu Ball, W.; Neff, J.K.; Gohil, V.M. The Role of Nonbilayer Phospholipids in Mitochondrial Structure and Function. FEBS Lett. 2018, 592, 1273–1290.
  128. Shadyro, O.I.; Yurkova, I.L.; Kisel, M.A. Radiation-Induced Peroxidation and Fragmentation of Lipids in a Model Membrane. Int. J. Radiat. Biol. 2002, 78, 211–217.
  129. Althoff, T.; Mills, D.J.; Popot, J.-L.; Kühlbrandt, W. Arrangement of Electron Transport Chain Components in Bovine Mitochondrial Supercomplex I1III2IV1. EMBO J. 2011, 30, 4652–4664.
  130. Ostrander, D.B.; Zhang, M.; Mileykovskaya, E.; Rho, M.; Dowhan, W. Lack of Mitochondrial Anionic Phospholipids Causes an Inhibition of Translation of Protein Components of the Electron Transport Chain. A Yeast Genetic Model System for the Study of Anionic Phospholipid Function in Mitochondria. J. Biol. Chem. 2001, 276, 25262–25272.
  131. Yoshioka-Nishimura, M. Close Relationships Between the PSII Repair Cycle and Thylakoid Membrane Dynamics. Plant Cell Physiol. 2016, 57, 1115–1122.
  132. Megiatto, J.D.; Antoniuk-Pablant, A.; Sherman, B.D.; Kodis, G.; Gervaldo, M.; Moore, T.A.; Moore, A.L.; Gust, D. Mimicking the Electron Transfer Chain in Photosystem II with a Molecular Triad Thermodynamically Capable of Water Oxidation. Proc. Natl. Acad. Sci. USA 2012, 109, 15578–15583.
  133. Reiter, R.J. Melatonin: Lowering the High Price of Free Radicals. News Physiol. Sci. 2000, 15, 246–250.
  134. Reiter, R.J.; Tan, D.-X.; Terron, M.P.; Flores, L.J.; Czarnocki, Z. Melatonin and Its Metabolites: New Findings Regarding Their Production and Their Radical Scavenging Actions. Acta Biochim. Pol. 2007, 54, 1–9.
  135. 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.
  136. De Almeida, E.A.; Martinez, G.R.; Klitzke, C.F.; de Medeiros, M.H.G.; Di Mascio, P. Oxidation of Melatonin by Singlet Molecular Oxygen (O2(1deltag)) Produces N1-Acetyl-N2-Formyl-5-Methoxykynurenine. J. Pineal. Res. 2003, 35, 131–137.
  137. Matuszak, Z.; Bilska, M.A.; Reszka, K.J.; Chignell, C.F.; Bilski, P. Interaction of Singlet Molecular Oxygen with Melatonin and Related Indoles. Photochem. Photobiol. 2003, 78, 449–455.
  138. 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.
  139. Noda, Y.; Mori, A.; Liburdy, R.; Packer, L. Melatonin and Its Precursors Scavenge Nitric Oxide. J. Pineal. Res. 1999, 27, 159–163.
  140. Aydogan, S.; Yerer, M.B.; Goktas, A. Melatonin and Nitric Oxide. J. Endocrinol. Investig. 2006, 29, 281–287.
  141. Hardeland, R. Melatonin, Its Metabolites and Their Interference with Reactive Nitrogen Compounds. Molecules 2021, 26, 4105.
  142. Gilad, E.; Cuzzocrea, S.; Zingarelli, B.; Salzman, A.L.; Szabó, C. Melatonin Is a Scavenger of Peroxynitrite. Life Sci. 1997, 60, PL169–PL174.
  143. Galano, A.; Reiter, R.J. Melatonin and Its Metabolites vs. Oxidative Stress: From Individual Actions to Collective Protection. J. Pineal Res. 2018, 65, e12514.
  144. Purushothaman, A.; Sheeja, A.A.; Janardanan, D. Hydroxyl Radical Scavenging Activity of Melatonin and Its Related Indolamines. Free Radic. Res. 2020, 54, 373–383.
  145. Galano, A. On the Direct Scavenging Activity of Melatonin towards Hydroxyl and a Series of Peroxyl Radicals. Phys. Chem. Chem. Phys. 2011, 13, 7178–7188.
  146. 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.
  147. Galano, A.; Medina, M.E.; Tan, D.X.; Reiter, R.J. Melatonin and Its Metabolites as Copper Chelating Agents and Their Role in Inhibiting Oxidative Stress: A Physicochemical Analysis. J. Pineal. Res. 2015, 58, 107–116.
  148. Lúcio, M.; Nunes, C.; Gaspar, D.; Ferreira, H.; Lima, J.L.F.C.; Reis, S. Antioxidant Activity of Vitamin E and Trolox: Understanding of the Factors That Govern Lipid Peroxidation Studies In Vitro. Food Biophys. 2009, 4, 312–320.
  149. Yu, H.; Dickson, E.J.; Jung, S.-R.; Koh, D.-S.; Hille, B. High Membrane Permeability for Melatonin. J. Gen. Physiol. 2016, 147, 63–76.
  150. Watson, H. Biological Membranes. Essays Biochem. 2015, 59, 43–69.
  151. Römsing, S. Development and Validation of Bioanalytical Methods: Application to Melatonin and Selected Anti-Infective Drugs; Acta Universitatis Upsaliensis: Uppsala, Sweden, 2010.
  152. Zhang, J.; Yan, X.; Tian, Y.; Li, W.; Wang, H.; Li, Q.; Li, Y.; Li, Z.; Wu, T. Synthesis of a New Water-Soluble Melatonin Derivative with Low Toxicity and a Strong Effect on Sleep Aid. ACS Omega. 2020, 5, 6494–6499.
  153. Shida, C.S.; Castrucci, A.M.; Lamy-Freund, M.T. High Melatonin Solubility in Aqueous Medium. J. Pineal. Res. 1994, 16, 198–201.
  154. 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.
  155. 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.
  156. Aikens, J.; Dix, T.A. Perhydroxyl Radical (HOO.) Initiated Lipid Peroxidation. The Role of Fatty Acid Hydroperoxides. J. Biol. Chem. 1991, 266, 15091–15098.
  157. 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.
  158. Repetto, M.; Semprine, J.; Boveris, A. Lipid Peroxidation: Chemical Mechanism, Biological Implications and Analytical Determination. In Lipid Peroxidation; Catala, A., Ed.; IntechOpen: London, UK, 2012.
  159. 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.
  160. Niki, E. Lipid Peroxidation: Physiological Levels and Dual Biological Effects. Free Radic. Biol. Med. 2009, 47, 469–484.
  161. Esterbauer, H.; Schaur, R.J.; Zollner, H. Chemistry and Biochemistry of 4-Hydroxynonenal, Malonaldehyde and Related Aldehydes. Free Radic. Biol. Med. 1991, 11, 81–128.
  162. Kanner, J.; German, J.B.; Kinsella, J.E. Initiation of Lipid Peroxidation in Biological Systems. Crit. Rev. Food Sci. Nutr. 1987, 25, 317–364.
  163. Southorn, P.A.; Powis, G. Free Radicals in Medicine. I. Chemical Nature and Biologic Reactions. Mayo Clin. Proc. 1988, 63, 381–389.
  164. Yin, H.; Xu, L.; Porter, N.A. Free Radical Lipid Peroxidation: Mechanisms and Analysis. Chem. Rev. 2011, 111, 5944–5972.
  165. Wientjes, F.B.; Segal, A.W. NADPH Oxidase and the Respiratory Burst. Semin. Cell Biol. 1995, 6, 357–365.
  166. Wong, H.-S.; Dighe, P.A.; Mezera, V.; Monternier, P.-A.; Brand, M.D. Production of Superoxide and Hydrogen Peroxide from Specific Mitochondrial Sites under Different Bioenergetic Conditions. J. Biol. Chem. 2017, 292, 16804–16809.
  167. Bielski, B.H.J.; Cabelli, D.E.; Arudi, R.L.; Ross, A.B. Reactivity of HO2/O−2 Radicals in Aqueous Solution. J. Phys. Chem. Ref. Data 1985, 14, 1041–1100.
  168. Wardman, P. Reduction Potentials of One-Electron Couples Involving Free Radicals in Aqueous Solution. J. Phys. Chem. Ref. Data 1989, 18, 1637–1755.
  169. Hayyan, M.; Hashim, M.A.; AlNashef, I.M. Superoxide Ion: Generation and Chemical Implications. Chem. Rev. 2016, 116, 3029–3085.
  170. Collin, F. Chemical Basis of Reactive Oxygen Species Reactivity and Involvement in Neurodegenerative Diseases. Int. J. Mol. Sci. 2019, 20, 2407.
  171. Gebicki, J.M.; Bielski, B.H.J. Comparison of the Capacities of the Perhydroxyl and the Superoxide Radicals to Initiate Chain Oxidation of Linoleic Acid. J. Am. Chem. Soc. 1981, 103, 7020–7022.
  172. De Grey, A.D.N.J. HO2*: The Forgotten Radical. DNA Cell Biol. 2002, 21, 251–257.
  173. Halliwell, B.; Gutteridge, J.M. Oxygen Toxicity, Oxygen Radicals, Transition Metals and Disease. Biochem. J. 1984, 219, 1–14.
  174. Cordeiro, R.M. Reactive Oxygen Species at Phospholipid Bilayers: Distribution, Mobility and Permeation. Biochim. Biophys. Acta 2014, 1838, 438–444.
  175. Yusupov, M.; Wende, K.; Kupsch, S.; Neyts, E.C.; Reuter, S.; Bogaerts, A. Effect of Head Group and Lipid Tail Oxidation in the Cell Membrane Revealed through Integrated Simulations and Experiments. Sci. Rep. 2017, 7, 5761.
  176. Sathappa, M.; Alder, N.N. The Ionization Properties of Cardiolipin and Its Variants in Model Bilayers. Biochim. Biophys. Acta 2016, 1858, 1362–1372.
  177. Haines, T.H.; Dencher, N.A. Cardiolipin: A Proton Trap for Oxidative Phosphorylation. FEBS Lett. 2002, 528, 35–39.
  178. Horvath, S.E.; Daum, G. Lipids of Mitochondria. Prog. Lipid Res. 2013, 52, 590–614.
  179. Van den Brink-van der Laan, E.; Killian, J.A.; de Kruijff, B. Nonbilayer Lipids Affect Peripheral and Integral Membrane Proteins via Changes in the Lateral Pressure Profile. Biochim. Biophys. Acta 2004, 1666, 275–288.
  180. Khalifat, N.; Puff, N.; Bonneau, S.; Fournier, J.-B.; Angelova, M.I. Membrane Deformation under Local pH Gradient: Mimicking Mitochondrial Cristae Dynamics. Biophys. J. 2008, 95, 4924–4933.
  181. 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.
  182. Porcelli, A.M.; Ghelli, A.; Zanna, C.; Pinton, P.; Rizzuto, R.; Rugolo, M. pH Difference across the Outer Mitochondrial Membrane Measured with a Green Fluorescent Protein Mutant. Biochem. Biophys. Res. Commun. 2005, 326, 799–804.
  183. 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.
  184. Chicco, A.J.; Sparagna, G.C. Role of Cardiolipin Alterations in Mitochondrial Dysfunction and Disease. Am. J. Physiol. Cell Physiol. 2007, 292, C33–C44.
  185. Paradies, G.; Paradies, V.; Ruggiero, F.M.; Petrosillo, G. Oxidative Stress, Cardiolipin and Mitochondrial Dysfunction in Nonalcoholic Fatty Liver Disease. World J. Gastroenterol. 2014, 20, 14205–14218.
  186. Kim, J.; Minkler, P.E.; Salomon, R.G.; Anderson, V.E.; Hoppel, C.L. Cardiolipin: Characterization of Distinct Oxidized Molecular Species. J. Lipid Res. 2011, 52, 125–135.
  187. Paradies, G.; Ruggiero, F.M.; Petrosillo, G.; Quagliariello, E. Age-Dependent Decline in the Cytochrome c Oxidase Activity in Rat Heart Mitochondria: Role of Cardiolipin. FEBS Lett. 1997, 406, 136–138.
  188. Paradies, G.; Ruggiero, F.M.; Petrosillo, G.; Quagliariello, E. Peroxidative Damage to Cardiac Mitochondria: Cytochrome Oxidase and Cardiolipin Alterations. FEBS Lett. 1998, 424, 155–158.
  189. Paradies, G.; Petrosillo, G.; Pistolese, M.; Ruggiero, F.M. Reactive Oxygen Species Generated by the Mitochondrial Respiratory Chain Affect the Complex III Activity via Cardiolipin Peroxidation in Beef-Heart Submitochondrial Particles. Mitochondrion 2001, 1, 151–159.
  190. Petrosillo, G.; Ruggiero, F.M.; Pistolese, M.; Paradies, G. Reactive Oxygen Species Generated from the Mitochondrial Electron Transport Chain Induce Cytochrome c Dissociation from Beef-Heart Submitochondrial Particles via Cardiolipin Peroxidation. Possible Role in the Apoptosis. FEBS Lett. 2001, 509, 435–438.
  191. Paradies, G.; Petrosillo, G.; Pistolese, M.; Ruggiero, F.M. Reactive Oxygen Species Affect Mitochondrial Electron Transport Complex I Activity through Oxidative Cardiolipin Damage. Gene 2002, 286, 135–141.
  192. Paradies, G.; Petrosillo, G.; Pistolese, M.; Di Venosa, N.; Federici, A.; Ruggiero, F.M. Decrease in Mitochondrial Complex I Activity in Ischemic/reperfused Rat Heart: Involvement of Reactive Oxygen Species and Cardiolipin. Circ. Res. 2004, 94, 53–59.
  193. Paradies, G.; Petrosillo, G.; Paradies, V.; Ruggiero, F.M. Role of Cardiolipin Peroxidation and Ca2+ in Mitochondrial Dysfunction and Disease. Cell Calcium. 2009, 45, 643–650.
  194. Arnarez, C.; Mazat, J.-P.; Elezgaray, J.; Marrink, S.-J.; Periole, X. Evidence for Cardiolipin Binding Sites on the Membrane-Exposed Surface of the Cytochrome bc1. J. Am. Chem. Soc. 2013, 135, 3112–3120.
  195. Panov, A. Perhydroxyl Radical (HO2•) as Inducer of the Isoprostane Lipid Peroxidation in Mitochondria. Mol. Biol. 2018, 52, 295–305.
  196. Miranda, É.G.A.; Araujo-Chaves, J.C.; Kawai, C.; Brito, A.M.M.; Dias, I.W.R.; Arantes, J.T.; Nantes-Cardoso, I.L. Cardiolipin Structure and Oxidation Are Affected by Ca2+ at the Interface of Lipid Bilayers. Front. Chem. 2019, 7, 930.
  197. Cipolla-Neto, J.; Amaral, F.G.; Afeche, S.C.; Tan, D.X.; Reiter, R.J. Melatonin, Energy Metabolism, and Obesity: A Review. J. Pineal. Res. 2014, 56, 371–381.
  198. Sustarsic, E.G.; Ma, T.; Lynes, M.D.; Larsen, M.; Karavaeva, I.; Havelund, J.F.; Nielsen, C.H.; Jedrychowski, M.P.; Moreno-Torres, M.; Lundh, M.; et al. Cardiolipin Synthesis in Brown and Beige Fat Mitochondria Is Essential for Systemic Energy Homeostasis. Cell Metab. 2018, 28, 159–174.e11.
  199. Von Bank, H.; Hurtado-Thiele, M.; Oshimura, N.; Simcox, J. Mitochondrial Lipid Signaling and Adaptive Thermogenesis. Metabolites 2021, 11, 124.
  200. Lee, Y.; Willers, C.; Kunji, E.R.S.; Crichton, P.G. Uncoupling Protein 1 Binds One Nucleotide per Monomer and Is Stabilized by Tightly Bound Cardiolipin. Proc. Natl. Acad. Sci. USA 2015, 112, 6973–6978.
  201. Fernández Vázquez, G.; Reiter, R.J.; Agil, A. Melatonin Increases Brown Adipose Tissue Mass and Function in Zücker Diabetic Fatty Rats: Implications for Obesity Control. J. Pineal. Res. 2018, 64, e12472.
  202. 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.
  203. Ikon, N.; Ryan, R.O. Cardiolipin and Mitochondrial Cristae Organization. Biochim. Biophys. Acta Biomembr. 2017, 1859, 1156–1163.
  204. 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.
  205. Manganelli, V.; Capozzi, A.; Recalchi, S.; Riitano, G.; Mattei, V.; Longo, A.; Misasi, R.; Garofalo, T.; Sorice, M. The Role of Cardiolipin as a Scaffold Mitochondrial Phospholipid in Autophagosome Formation: In Vitro Evidence. Biomolecules 2021, 11, 222.
  206. Kagan, V.E.; Tyurin, V.A.; Jiang, J.; Tyurina, Y.Y.; Ritov, V.B.; Amoscato, A.A.; Osipov, A.N.; Belikova, N.A.; Kapralov, A.A.; Kini, V.; et al. Cytochrome c Acts as a Cardiolipin Oxygenase Required for Release of Proapoptotic Factors. Nat. Chem. Biol. 2005, 1, 223–232.
  207. Petrosillo, G.; Casanova, G.; Matera, M.; Ruggiero, F.M.; Paradies, G. Interaction of Peroxidized Cardiolipin with Rat-Heart Mitochondrial Membranes: Induction of Permeability Transition and Cytochrome c Release. FEBS Lett. 2006, 580, 6311–6316.
  208. Li, X.-X.; Tsoi, B.; Li, Y.-F.; Kurihara, H.; He, R.-R. Cardiolipin and Its Different Properties in Mitophagy and Apoptosis. J. Histochem. Cytochem. 2015, 63, 301–311.
  209. Dudek, J. Role of Cardiolipin in Mitochondrial Signaling Pathways. Front. Cell Dev. Biol. 2017, 5, 90.
  210. Huang, W.; Choi, W.; Hu, W.; Mi, N.; Guo, Q.; Ma, M.; Liu, M.; Tian, Y.; Lu, P.; Wang, F.-L.; et al. Crystal Structure and Biochemical Analyses Reveal Beclin 1 as a Novel Membrane Binding Protein. Cell Res. 2012, 22, 473–489.
  211. Gonzalvez, F.; Pariselli, F.; Dupaigne, P.; Budihardjo, I.; Lutter, M.; Antonsson, B.; Diolez, P.; Manon, S.; Martinou, J.-C.; Goubern, M.; et al. tBid Interaction with Cardiolipin Primarily Orchestrates Mitochondrial Dysfunctions and Subsequently Activates Bax and Bak. Cell Death Differ. 2005, 12, 614–626.
  212. Gonzalvez, F.; Schug, Z.T.; Houtkooper, R.H.; MacKenzie, E.D.; Brooks, D.G.; Wanders, R.J.A.; Petit, P.X.; Vaz, F.M.; Gottlieb, E. Cardiolipin Provides an Essential Activating Platform for Caspase-8 on Mitochondria. J. Cell Biol. 2008, 183, 681–696.
  213. Iyer, S.S.; He, Q.; Janczy, J.R.; Elliott, E.I.; Zhong, Z.; Olivier, A.K.; Sadler, J.J.; Knepper-Adrian, V.; Han, R.; Qiao, L.; et al. Mitochondrial Cardiolipin is Required for Nlrp3 Inflammasome Activation. Immunity 2013, 39, 311–323.
  214. De Zoete, M.R.; Palm, N.W.; Zhu, S.; Flavell, R.A. Inflammasomes. Cold Spring Harb. Perspect. Biol. 2014, 6, a016287.
  215. Sharif, H.; Wang, L.; Wang, W.L.; Magupalli, V.G.; Andreeva, L.; Qiao, Q.; Hauenstein, A.V.; Wu, Z.; Núñez, G.; Mao, Y.; et al. Structural Mechanism for NEK7-Licensed Activation of NLRP3 Inflammasome. Nature 2019, 570, 338–343.
  216. He, Y.; Hara, H.; Núñez, G. Mechanism and Regulation of NLRP3 Inflammasome Activation. Trends Biochem. Sci. 2016, 41, 1012–1021.
  217. 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.
  218. Yeon, S.H.; Yang, G.; Lee, H.E.; Lee, J.Y. Oxidized Phosphatidylcholine Induces the Activation of NLRP3 Inflammasome in Macrophages. J. Leukoc. Biol. 2017, 101, 205–215.
  219. 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.
  220. Raturi, A.; Simmen, T. Where the Endoplasmic Reticulum and the Mitochondrion Tie the Knot: The Mitochondria-Associated Membrane (MAM). Biochim. Biophys. Acta 2013, 1833, 213–224.
  221. 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.
  222. Ono, K.; Mochizuki, H.; Ikeda, T.; Nihira, T.; Takasaki, J.-I.; Teplow, D.B.; Yamada, M. Effect of Melatonin on α-Synuclein Self-Assembly and Cytotoxicity. Neurobiol. Aging. 2012, 33, 2172–2185.
  223. Zampol, M.A.; Barros, M.H. Melatonin Improves Survival and Respiratory Activity of Yeast Cells Challenged by Alpha-Synuclein and Menadione. Yeast 2018, 35, 281–290.
  224. 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.
  225. Compan, V.; Martín-Sánchez, F.; Baroja-Mazo, A.; López-Castejón, G.; Gomez, A.I.; Verkhratsky, A.; Brough, D.; Pelegrín, P. Apoptosis-Associated Speck-like Protein Containing a CARD Forms Specks but Does Not Activate Caspase-1 in the Absence of NLRP3 during Macrophage Swelling. J. Immunol. 2015, 194, 1261–1273.
  226. Stehlik, C.; Lee, S.H.; Dorfleutner, A.; Stassinopoulos, A.; Sagara, J.; Reed, J.C. Apoptosis-Associated Speck-like Protein Containing a Caspase Recruitment Domain Is a Regulator of Procaspase-1 Activation. J. Immunol. 2003, 171, 6154–6163.
  227. Ozgur, S.; Buchwald, G.; Falk, S.; Chakrabarti, S.; Prabu, J.R.; Conti, E. The Conformational Plasticity of Eukaryotic RNA-Dependent ATPases. FEBS J. 2015, 282, 850–863.
  228. Owttrim, G.W. RNA Helicases: Diverse Roles in Prokaryotic Response to Abiotic Stress. RNA Biol. 2013, 10, 96–110.
  229. Aumiller, W.M.; Keating, C.D. Phosphorylation-Mediated RNA/peptide Complex Coacervation as a Model for Intracellular Liquid Organelles. Nat. Chem. 2016, 8, 129–137.
  230. Jin, M.; Fuller, G.G.; Han, T.; Yao, Y.; Alessi, A.F.; Freeberg, M.A.; Roach, N.P.; Moresco, J.J.; Karnovsky, A.; Baba, M.; et al. Glycolytic Enzymes Coalesce in G Bodies under Hypoxic Stress. Cell Rep. 2017, 20, 895–908.
  231. Kamagata, K.; Kanbayashi, S.; Honda, M.; Itoh, Y.; Takahashi, H.; Kameda, T.; Nagatsugi, F.; Takahashi, S. Liquid-like Droplet Formation by Tumor Suppressor p53 Induced by Multivalent Electrostatic Interactions between Two Disordered Domains. Sci. Rep. 2020, 10, 580.
  232. Hofweber, M.; Dormann, D. Friend or Foe-Post-Translational Modifications as Regulators of Phase Separation and RNP Granule Dynamics. J. Biol. Chem. 2019, 294, 7137–7150.
  233. Ardito, F.; Giuliani, M.; Perrone, D.; Troiano, G.; Lo Muzio, L. The Crucial Role of Protein Phosphorylation in Cell Signaling and Its Use as Targeted Therapy (Review). Int. J. Mol. Med. 2017, 40, 271–280.
  234. Fox, D.; Man, S.M. DDX3X: Stressing the NLRP3 Inflammasome. Cell Res. 2019, 29, 969–970.
  235. Gustafson, E.A.; Wessel, G.M. DEAD-Box Helicases: Posttranslational Regulation and Function. Biochem. Biophys. Res. Commun. 2010, 395, 1–6.
  236. 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.
  237. Ron, D. Translational Control in the Endoplasmic Reticulum Stress Response. J. Clin. Investig. 2002, 110, 1383–1388.
  238. Wek, R.C.; Jiang, H.-Y.; Anthony, T.G. Coping with Stress: eIF2 Kinases and Translational Control. Biochem. Soc. Trans. 2006, 34, 7–11.
  239. Donnelly, N.; Gorman, A.M.; Gupta, S.; Samali, A. The eIF2α Kinases: Their Structures and Functions. Cell. Mol. Life Sci. 2013, 70, 3493–3511.
  240. Pakos-Zebrucka, K.; Koryga, I.; Mnich, K.; Ljujic, M.; Samali, A.; Gorman, A.M. The Integrated Stress Response. EMBO Rep. 2016, 17, 1374–1395.
  241. Sidrauski, C.; McGeachy, A.M.; Ingolia, N.T.; Walter, P. The Small Molecule ISRIB Reverses the Effects of eIF2α Phosphorylation on Translation and Stress Granule Assembly. Elife 2015, 4, e05033.
  242. Wheeler, J.R.; Matheny, T.; Jain, S.; Abrisch, R.; Parker, R. Distinct Stages in Stress Granule Assembly and Disassembly. Elife 2016, 5, e18413.
  243. Anderson, P.; Kedersha, N. Visibly Stressed: The Role of eIF2, TIA-1, and Stress Granules in Protein Translation. Cell Stress Chaperones 2002, 7, 213–221.
  244. Miller, Y.I.; Navia-Pelaez, J.M.; Corr, M.; Yaksh, T.L. Lipid Rafts in Glial Cells: Role in Neuroinflammation and Pain Processing: Thematic Review Series: Biology of Lipid Rafts. J. Lipid Res. 2020, 61, 655–666.
  245. Li, Y.C.; Park, M.J.; Ye, S.-K.; Kim, C.-W.; Kim, Y.-N. Elevated Levels of Cholesterol-Rich Lipid Rafts in Cancer Cells Are Correlated with Apoptosis Sensitivity Induced by Cholesterol-Depleting Agents. Am. J. Pathol. 2006, 168, 1107–1118; quiz 1404–1405.
  246. Guan, Y.; Han, F. Key Mechanisms and Potential Targets of the NLRP3 Inflammasome in Neurodegenerative Diseases. Front. Integr. Neurosci. 2020, 14, 37.
  247. Arioz, B.I.; Tastan, B.; Tarakcioglu, E.; Tufekci, K.U.; Olcum, M.; Ersoy, N.; Bagriyanik, A.; Genc, K.; Genc, S. Melatonin Attenuates LPS-Induced Acute Depressive-Like Behaviors and Microglial NLRP3 Inflammasome Activation Through the SIRT1/Nrf2 Pathway. Front. Immunol. 2019, 10, 1511.
  248. Zhang, J.; Lu, X.; Liu, M.; Fan, H.; Zheng, H.; Zhang, S.; Rahman, N.; Wołczyński, S.; Kretowski, A.; Li, X. Melatonin Inhibits Inflammasome-Associated Activation of Endothelium and Macrophages Attenuating Pulmonary Arterial Hypertension. Cardiovasc. Res. 2020, 116, 2156–2169.
  249. Chen, F.; Jiang, G.; Liu, H.; Li, Z.; Pei, Y.; Wang, H.; Pan, H.; Cui, H.; Long, J.; Wang, J.; et al. Melatonin Alleviates Intervertebral Disc Degeneration by Disrupting the IL-1β/NF-κB-NLRP3 Inflammasome Positive Feedback Loop. Bone Res. 2020, 8, 10.
  250. Severcan, F.; Sahin, I.; Kazanci, N. Melatonin Strongly Interacts with Zwitterionic Model Membranes--Evidence from Fourier Transform Infrared Spectroscopy and Differential Scanning Calorimetry. Biochim. Biophys. Acta 2005, 1668, 215–222.
  251. Wong-Ekkabut, J.; Xu, Z.; Triampo, W.; Tang, I.-M.; Tieleman, D.P.; Monticelli, L. Effect of Lipid Peroxidation on the Properties of Lipid Bilayers: A Molecular Dynamics Study. Biophys. J. 2007, 93, 4225–4236.
  252. Kaplán, P.; Racay, P.; Lehotský, J.; Mézesová, V. Change in Fluidity of Brain Endoplasmic Reticulum Membranes by Oxygen Free Radicals: A Protective Effect of Stobadine, Alpha-Tocopherol Acetate, and Butylated Hydroxytoluene. Neurochem. Res. 1995, 20, 815–820.
  253. Yu, B.P.; Suescun, E.A.; Yang, S.Y. Effect of Age-Related Lipid Peroxidation on Membrane Fluidity and Phospholipase A2: Modulation by Dietary Restriction. Mech. Ageing. Dev. 1992, 65, 17–33.
  254. Dies, H.; Toppozini, L.; Rheinstädter, M.C. The Interaction between Amyloid-β Peptides and Anionic Lipid Membranes Containing Cholesterol and Melatonin. PLoS ONE 2014, 9, e99124.
  255. Dies, H.; Cheung, B.; Tang, J.; Rheinstädter, M.C. The Organization of Melatonin in Lipid Membranes. Biochim. Biophys. Acta 2015, 1848, 1032–1040.
  256. Rudzite, V.; Jurika, E.; Jirgensons, J. Changes in Membrane Fluidity Induced by Tryptophan and Its Metabolites. In Tryptophan, Serotonin, and Melatonin: Basic Aspects and Applications; Huether, G., Kochen, W., Simat, T.J., Steinhart, H., Eds.; Springer: Boston, MA, USA, 1999.
  257. Beyenbach, K.W.; Wieczorek, H. The V-Type H+ ATPase: Molecular Structure and Function, Physiological Roles and Regulation. J. Exp. Biol. 2006, 209 Pt 4, 577–589.
  258. Bae, T.-J.; Kim, M.-S.; Kim, J.-W.; Kim, B.-W.; Choo, H.-J.; Lee, J.-W.; Kim, K.-B.; Lee, C.S.; Kim, J.-H.; Chang, S.Y.; et al. Lipid Raft Proteome Reveals ATP Synthase Complex in the Cell Surface. Proteomics 2004, 4, 3536–3548.
  259. Kim, B.-W.; Choo, H.-J.; Lee, J.-W.; Kim, J.-H.; Ko, Y.-G. Extracellular ATP Is Generated by ATP Synthase Complex in Adipocyte Lipid Rafts. Exp. Mol. Med. 2004, 36, 476–485.
  260. Alexandre, H.; Mathieu, B.; Charpentier, C. Alteration in Membrane Fluidity and Lipid Composition, and Modulation of H+-ATPase Activity in Saccharomyces Cerevisiae Caused by Decanoic Acid. Available online: (accessed on 27 March 2021).
  261. Keeffe, E.B.; Blankenship, N.M.; Scharschmidt, B.F. Alteration of Rat Liver Plasma Membrane Fluidity and ATPase Activity by Chlorpromazine Hydrochloride and Its Metabolites. Gastroenterology 1980, 79, 222–231.
  262. Garcia, A.; Pochinda, S.; Elgaard-Jørgensen, P.N.; Khandelia, H.; Clarke, R.J. Evidence for ATP Interaction with Phosphatidylcholine Bilayers. Langmuir 2019, 35, 9944–9953.
  263. Chen, J.J.; Yu, B.P. Alterations in Mitochondrial Membrane Fluidity by Lipid Peroxidation Products. Free Radic. Biol. Med. 1994, 17, 411–418.
  264. Kholodenko, B.N.; Hoek, J.B.; Westerhoff, H.V. Why Cytoplasmic Signalling Proteins Should Be Recruited to Cell Membranes. Trends Cell Biol. 2000, 10, 173–178.
  265. Botterbusch, S.; Baumgart, T. Interactions between Phase-Separated Liquids and Membrane Surfaces. NATO Adv. Sci. Inst. Ser. E Appl. Sci. 2021, 11, 1288.
  266. Alimohamadi, H.; Rangamani, P. Modeling Membrane Curvature Generation due to Membrane−Protein Interactions. Biomolecules 2018, 8, 120.
  267. Prévost, C.; Zhao, H.; Manzi, J.; Lemichez, E.; Lappalainen, P.; Callan-Jones, A.; Bassereau, P. IRSp53 Senses Negative Membrane Curvature and Phase Separates along Membrane Tubules. Nat. Commun. 2015, 6, 8529.
  268. Gallop, J.L.; McMahon, H.T. BAR Domains and Membrane Curvature: Bringing Your Curves to the BAR. Biochem. Soc. Symp. 2005, 72, 223–231.
  269. Fabiani, C.; Antollini, S.S. Alzheimer’s Disease as a Membrane Disorder: Spatial Cross-Talk Among Beta-Amyloid Peptides, Nicotinic Acetylcholine Receptors and Lipid Rafts. Front. Cell Neurosci. 2019, 13, 309.
  270. 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.
  271. Simons, K.; Ikonen, E. Functional Rafts in Cell Membranes. Nature 1997, 387, 569–572.
  272. Simons, K.; Toomre, D. Lipid Rafts and Signal Transduction. Nat. Rev. Mol. Cell Biol. 2000, 1, 31–39.
  273. Bagnat, M.; Keränen, S.; Shevchenko, A.; Shevchenko, A.; Simons, K. Lipid Rafts Function in Biosynthetic Delivery of Proteins to the Cell Surface in Yeast. Proc. Natl. Acad. Sci. USA 2000, 97, 3254–3259.
  274. Ikonen, E. Roles of Lipid Rafts in Membrane Transport. Curr. Opin. Cell Biol. 2001, 13, 470–477.
  275. Jin, S.; Zhou, F.; Katirai, F.; Li, P.-L. Lipid Raft Redox Signaling: Molecular Mechanisms in Health and Disease. Antioxid. Redox Signal. 2011, 15, 1043–1083.
  276. Jiang, F.; Zhang, Y.; Dusting, G.J. NADPH Oxidase-Mediated Redox Signaling: Roles in Cellular Stress Response, Stress Tolerance, and Tissue Repair. Pharmacol. Rev. 2011, 63, 218–242.
  277. Garofalo, T.; Manganelli, V.; Grasso, M.; Mattei, V.; Ferri, A.; Misasi, R.; Sorice, M. Role of Mitochondrial Raft-like Microdomains in the Regulation of Cell Apoptosis. Apoptosis 2015, 20, 621–634.
  278. Samhan-Arias, A.K.; Garcia-Bereguiain, M.A.; Martin-Romero, F.J.; Gutierrez-Merino, C. Clustering of Plasma Membrane-Bound Cytochrome b5 Reductase within “Lipid Raft” Microdomains of the Neuronal Plasma Membrane. Mol. Cell. Neurosci. 2009, 40, 14–26.
  279. Kinnun, J.J.; Bolmatov, D.; Lavrentovich, M.O.; Katsaras, J. Lateral Heterogeneity and Domain Formation in Cellular Membranes. Chem. Phys. Lipids 2020, 232, 104976.
  280. Heerklotz, H. Triton Promotes Domain Formation in Lipid Raft Mixtures. Biophys. J. 2002, 83, 2693–2701.
  281. Bolmatov, D.; Soloviov, D.; Zhernenkov, M.; Zav’yalov, D.; Mamontov, E.; Suvorov, A.; Cai, Y.Q.; Katsaras, J. Molecular Picture of the Transient Nature of Lipid Rafts. Langmuir 2020, 36, 4887–4896.
  282. Aponte-Santamaría, C.; Brunken, J.; Gräter, F. Stress Propagation through Biological Lipid Bilayers in Silico. J. Am. Chem. Soc. 2017, 139, 13588–13591.
  283. Mei, N.; Robinson, M.; Davis, J.H.; Leonenko, Z. Melatonin Alters Fluid Phase Co-Existence in POPC/DPPC/cholesterol Membranes. Biophys. J. 2020, 119, 2391–2402.
  284. Choi, Y.; Attwood, S.J.; Hoopes, M.I.; Drolle, E.; Karttunen, M.; Leonenko, Z. Melatonin Directly Interacts with Cholesterol and Alleviates Cholesterol Effects in Dipalmitoylphosphatidylcholine Monolayers. Soft Matter 2014, 10, 206–213.
  285. Del Mar Martínez-Senac, M.; Villalaín, J.; Gómez-Fernández, J.C. Structure of the Alzheimer Beta-Amyloid Peptide (25-35) and Its Interaction with Negatively Charged Phospholipid Vesicles. Eur. J. Biochem. 1999, 265, 744–753.
  286. Sani, M.-A.; Gehman, J.D.; Separovic, F. Lipid Matrix Plays a Role in Abeta Fibril Kinetics and Morphology. FEBS Lett. 2011, 585, 749–754.
  287. Hane, F.; Drolle, E.; Gaikwad, R.; Faught, E.; Leonenko, Z. Amyloid-β Aggregation on Model Lipid Membranes: An Atomic Force Microscopy Study. J. Alzheimers. Dis. 2011, 26, 485–494.
  288. Ahyayauch, H.; Raab, M.; Busto, J.V.; Andraka, N.; Arrondo, J.-L.R.; Masserini, M.; Tvaroska, I.; Goñi, F.M. Binding of β-Amyloid (1-42) Peptide to Negatively Charged Phospholipid Membranes in the Liquid-Ordered State: Modeling and Experimental Studies. Biophys. J. 2012, 103, 453–463.
  289. Ding, H.; Schauerte, J.A.; Steel, D.G.; Gafni, A. β-Amyloid (1-40) Peptide Interactions with Supported Phospholipid Membranes: A Single-Molecule Study. Biophys. J. 2012, 103, 1500–1509.
  290. Magarkar, A.; Dhawan, V.; Kallinteri, P.; Viitala, T.; Elmowafy, M.; Róg, T.; Bunker, A. Cholesterol Level Affects Surface Charge of Lipid Membranes in Saline Solution. Sci. Rep. 2014, 4, 5005.
  291. Chen, Z.; Rand, R.P. The Influence of Cholesterol on Phospholipid Membrane Curvature and Bending Elasticity. Biophys. J. 1997, 73, 267–276.
  292. Drolle, E.; Gaikwad, R.M.; Leonenko, Z. Nanoscale Electrostatic Domains in Cholesterol-Laden Lipid Membranes Create a Target for Amyloid Binding. Biophys. J. 2012, 103, L27–L29.
  293. Finot, E.; Leonenko, Y.; Moores, B.; Eng, L.; Amrein, M.; Leonenko, Z. Effect of Cholesterol on Electrostatics in Lipid-Protein Films of a Pulmonary Surfactant. Langmuir 2010, 26, 1929–1935.
  294. Eckert, G.P.; Kirsch, C.; Leutz, S.; Wood, W.G.; Müller, W.E. Cholesterol Modulates Amyloid Beta-Peptide’s Membrane Interactions. Pharmacopsychiatry 2003, 36 (Suppl. 2), S136–S143.
  295. Gibson Wood, W.; Eckert, G.P.; Igbavboa, U.; Müller, W.E. Amyloid Beta-Protein Interactions with Membranes and Cholesterol: Causes or Casualties of Alzheimer’s Disease. Biochim. Biophys. Acta 2003, 1610, 281–290.
  296. Gostincar, C.; Turk, M.; Gunde-Cimerman, N. The Evolution of Fatty Acid Desaturases and Cytochrome b5 in Eukaryotes. J. Membr. Biol. 2010, 233, 63–72.
  297. Ito, A.; Hayashi, S.; Yoshida, T. Participation of a Cytochrome b5-like Hemoprotein of Outer Mitochondrial Membrane (OM Cytochrome B) in NADH-Semidehydroascorbic Acid Reductase Activity of Rat Liver. Biochem. Biophys. Res. Commun. 1981, 101, 591–598.
  298. Navarro, F.; Villalba, J.M.; Crane, F.L.; Mackellar, W.C.; Navas, P. A Phospholipid-Dependent NADH-Coenzyme Q Reductase from Liver Plasma Membrane. Biochem. Biophys. Res. Commun. 1995, 212, 138–143.
  299. Percy, M.J.; Lappin, T.R. Recessive Congenital Methaemoglobinaemia: Cytochrome b(5) Reductase Deficiency. Br. J. Haematol. 2008, 141, 298–308.
  300. Siendones, E.; SantaCruz-Calvo, S.; Martín-Montalvo, A.; Cascajo, M.V.; Ariza, J.; López-Lluch, G.; Villalba, J.M.; Acquaviva-Bourdain, C.; Roze, E.; Bernier, M.; et al. Membrane-Bound CYB5R3 Is a Common Effector of Nutritional and Oxidative Stress Response through FOXO3a and Nrf2. Antioxid. Redox Signal. 2014, 21, 1708–1725.
  301. Marques-da-Silva, D.; Samhan-Arias, A.K.; Tiago, T.; Gutierrez-Merino, C. L-Type Calcium Channels and Cytochrome b5 Reductase Are Components of Protein Complexes Tightly Associated with Lipid Rafts Microdomains of the Neuronal Plasma Membrane. J. Proteom. 2010, 73, 1502–1510.
  302. Nikiforova, A.B.; Saris, N.-E.L.; Kruglov, A.G. External Mitochondrial NADH-Dependent Reductase of Redox Cyclers: VDAC1 or Cyb5R3? Free Radic. Biol. Med. 2014, 74, 74–84.
  303. Bakalova, R.; Zhelev, Z.; Miller, T.; Aoki, I.; Higashi, T. New Potential Biomarker for Stratification of Patients for Pharmacological Vitamin C in Adjuvant Settings of Cancer Therapy. Redox Biol. 2020, 28, 101357.
  304. Mihara, K.; Sato, R. Molecular Cloning and Sequencing of cDNA for Yeast Porin, an Outer Mitochondrial Membrane Protein: A Search for Targeting Signal in the Primary Structure. EMBO J. 1985, 4, 769–774.
  305. Thinnes, F.P.; Götz, H.; Kayser, H.; Benz, R.; Schmidt, W.E.; Kratzin, H.D.; Hilschmann, N. Identification of human porins. I. Purification of a porin from human B-lymphocytes (Porin 31HL) and the topochemical proof of its expression on the plasmalemma of the progenitor cell. Biol. Chem. Hoppe Seyler 1989, 370, 1253–1264.
  306. Bàthori, G.; Parolini, I.; Tombola, F.; Szabò, I.; Messina, A.; Oliva, M.; De Pinto, V.; Lisanti, M.; Sargiacomo, M.; Zoratti, M. Porin Is Present in the Plasma Membrane Where It Is Concentrated in Caveolae and Caveolae-Related Domains. J. Biol. Chem. 1999, 274, 29607–29612.
  307. Herrera, J.L.; Diaz, M.; Hernández-Fernaud, J.R.; Salido, E.; Alonso, R.; Fernández, C.; Morales, A.; Marin, R. Voltage-Dependent Anion Channel as a Resident Protein of Lipid Rafts: Post-Transductional Regulation by Estrogens and Involvement in Neuronal Preservation against Alzheimer’s Disease. J. Neurochem. 2011, 116, 820–827.
  308. De Pinto, V.; Messina, A.; Lane, D.J.R.; Lawen, A. Voltage-Dependent Anion-Selective Channel (VDAC) in the Plasma Membrane. FEBS Lett. 2010, 584, 1793–1799.
  309. Anishkin, A.; Loukin, S.H.; Teng, J.; Kung, C. Feeling the Hidden Mechanical Forces in Lipid Bilayer Is an Original Sense. Proc. Natl. Acad. Sci. USA 2014, 111, 7898–7905.
  310. Anishkin, A.; Kung, C. Stiffened Lipid Platforms at Molecular Force Foci. Proc. Natl. Acad. Sci. USA 2013, 110, 4886–4892.
  311. Martinac, B.; Adler, J.; Kung, C. Mechanosensitive Ion Channels of E. Coli Activated by Amphipaths. Nature 1990, 348, 261–263.
  312. Markin, V.S.; Martinac, B. Mechanosensitive Ion Channels as Reporters of Bilayer Expansion. A Theoretical Model. Biophys. J. 1991, 60, 1120–1127.
  313. Dart, C. Lipid Microdomains and the Regulation of Ion Channel Function. J. Physiol. 2010, 588 Pt 17, 3169–3178.
  314. Samhan-Arias, A.K.; Marques-da-Silva, D.; Yanamala, N.; Gutierrez-Merino, C. Stimulation and Clustering of Cytochrome b5 Reductase in Caveolin-Rich Lipid Microdomains Is an Early Event in Oxidative Stress-Mediated Apoptosis of Cerebellar Granule Neurons. J. Proteom. 2012, 75, 2934–2949.
  315. Samhan-Arias, A.K.; Fortalezas, S.; Cordas, C.M.; Moura, I.; Moura, J.J.G.; Gutierrez-Merino, C. Cytochrome b5 Reductase Is the Component from Neuronal Synaptic Plasma Membrane Vesicles That Generates Superoxide Anion upon Stimulation by Cytochrome c. Redox Biol. 2018, 15, 109–114.
  316. Samhan-Arias, A.K.; Gutierrez-Merino, C. Purified NADH-Cytochrome b5 Reductase Is a Novel Superoxide Anion Source Inhibited by Apocynin: Sensitivity to Nitric Oxide and Peroxynitrite. Free Radic. Biol. Med. 2014, 73, 174–189.
  317. La Piana, G.; Marzulli, D.; Gorgoglione, V.; Lofrumento, N.E. Porin and Cytochrome Oxidase Containing Contact Sites Involved in the Oxidation of Cytosolic NADH. Arch. Biochem. Biophys. 2005, 436, 91–100.
  318. Martín-Romero, F.J.; Gutiérrez-Martín, Y.; Henao, F.; Gutiérrez-Merino, C. The NADH Oxidase Activity of the Plasma Membrane of Synaptosomes Is a Major Source of Superoxide Anion and Is Inhibited by Peroxynitrite. J. Neurochem. 2002, 82, 604–614.
  319. Zizi, M.; Forte, M.; Blachly-Dyson, E.; Colombini, M. NADH Regulates the Gating of VDAC, the Mitochondrial Outer Membrane Channel. J. Biol. Chem. 1994, 269, 1614–1616.
  320. Shoshan-Barmatz, V.; Shteinfer-Kuzmine, A.; Verma, A. VDAC1 at the Intersection of Cell Metabolism, Apoptosis, and Diseases. Biomolecules 2020, 10, 1485.
  321. Lemasters, J.J. Evolution of Voltage-Dependent Anion Channel Function: From Molecular Sieve to Governator to Actuator of Ferroptosis. Front. Oncol. 2017, 7, 303.
  322. Elinder, F.; Akanda, N.; Tofighi, R.; Shimizu, S.; Tsujimoto, Y.; Orrenius, S.; Ceccatelli, S. Opening of Plasma Membrane Voltage-Dependent Anion Channels (VDAC) Precedes Caspase Activation in Neuronal Apoptosis Induced by Toxic Stimuli. Cell Death Differ. 2005, 12, 1134–1140.
  323. Rostovtseva, T.; Colombini, M. VDAC Channels Mediate and Gate the Flow of ATP: Implications for the Regulation of Mitochondrial Function. Biophys. J. 1997, 72, 1954–1962.
  324. Rostovtseva, T.K.; Tan, W.; Colombini, M. On the Role of VDAC in Apoptosis: Fact and Fiction. J. Bioenerg. Biomembr. 2005, 37, 129–142.
  325. McCommis, K.S.; Baines, C.P. The Role of VDAC in Cell Death: Friend or Foe? Biochim. Biophys. Acta 2012, 1818, 1444–1450.
  326. Shoshan-Barmatz, V.; Maldonado, E.N.; Krelin, Y. VDAC1 at the Crossroads of Cell Metabolism, Apoptosis and Cell Stress. Cell Stress Chaperones 2017, 1, 11–36.
  327. Camara, A.K.S.; Zhou, Y.; Wen, P.-C.; Tajkhorshid, E.; Kwok, W.-M. Mitochondrial VDAC1: A Key Gatekeeper as Potential Therapeutic Target. Front. Physiol. 2017, 8, 460.
  328. Sorice, M.; Manganelli, V.; Matarrese, P.; Tinari, A.; Misasi, R.; Malorni, W.; Garofalo, T. Cardiolipin-Enriched Raft-like Microdomains Are Essential Activating Platforms for Apoptotic Signals on Mitochondria. FEBS Lett. 2009, 583, 2447–2450.
  329. Sorice, M.; Mattei, V.; Matarrese, P.; Garofalo, T.; Tinari, A.; Gambardella, L.; Ciarlo, L.; Manganelli, V.; Tasciotti, V.; Misasi, R.; et al. Dynamics of Mitochondrial Raft-like Microdomains in Cell Life and Death. Commun. Integr. Biol. 2012, 5, 217–219.
  330. Malorni, W.; Giammarioli, A.M.; Garofalo, T.; Sorice, M. Dynamics of Lipid Raft Components during Lymphocyte Apoptosis: The Paradigmatic Role of GD3. Apoptosis 2007, 12, 941–949.
  331. Brand, M.D. Mitochondrial Generation of Superoxide and Hydrogen Peroxide as the Source of Mitochondrial Redox Signaling. Free Radic. Biol. Med. 2016, 100, 14–31.
  332. Schneider, C. An Update on Products and Mechanisms of Lipid Peroxidation. Mol. Nutr. Food Res. 2009, 53, 315–321.
  333. Ting, H.-C.; Chen, L.-T.; Chen, J.-Y.; Huang, Y.-L.; Xin, R.-C.; Chan, J.-F.; Hsu, Y.-H.H. Double Bonds of Unsaturated Fatty Acids Differentially Regulate Mitochondrial Cardiolipin Remodeling. Lipids Health Dis. 2019, 18, 53.
  334. Musatov, A. Contribution of Peroxidized Cardiolipin to Inactivation of Bovine Heart Cytochrome c Oxidase. Free Radic. Biol. Med. 2006, 41, 238–246.
  335. Venegas, C.; García, J.A.; Escames, G.; Ortiz, F.; López, A.; Doerrier, C.; García-Corzo, L.; López, L.C.; Reiter, R.J.; Acuña-Castroviejo, D. Extrapineal Melatonin: Analysis of Its Subcellular Distribution and Daily Fluctuations. J. Pineal Res. 2012, 52, 217–227.
  336. Cesarini, E.; Cerioni, L.; Canonico, B.; Di Sario, G.; Guidarelli, A.; Lattanzi, D.; Savelli, D.; Guescini, M.; Nasoni, M.G.; Bigini, N.; et al. Melatonin Protects Hippocampal HT22 Cells from the Effects of Serum Deprivation Specifically Targeting Mitochondria. PLoS ONE 2018, 13, e0203001.
  337. 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.
  338. 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.
  339. 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.
  340. 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.
  341. Reiter, R.J.; Sharma, R.; Ma, Q.; Rorsales-Corral, S.; de Almeida Chuffa, L.G. Melatonin Inhibits Warburg-Dependent Cancer by Redirecting Glucose Oxidation to the Mitochondria: A Mechanistic Hypothesis. Cell. Mol. Life Sci. 2020, 77, 2527–2542.
  342. Reiter, R.J.; Sharma, R.; Pires de Campos Zuccari, D.A.; de Almeida Chuffa, L.G.; Manucha, W.; Rodriguez, C. Melatonin Synthesis in and Uptake by Mitochondria: Implications for Diseased Cells with Dysfunctional Mitochondria. Future Med. Chem. 2021, 13, 335–339.
  343. Xia, Y.; Chen, S.; Zeng, S.; Zhao, Y.; Zhu, C.; Deng, B.; Zhu, G.; Yin, Y.; Wang, W.; Hardeland, R.; et al. Melatonin in Macrophage Biology: Current Understanding and Future Perspectives. J. Pineal. Res. 2019, 66, e12547.
  344. Reiter, R.J.; Sharma, R.; Ma, Q. Switching Diseased Cells from Cytosolic Aerobic Glycolysis to Mitochondrial Oxidative Phosphorylation: A Metabolic Rhythm Regulated by Melatonin? J. Pineal. Res. 2021, 70, e12677.
  345. Fuller, G.G.; Han, T.; Freeberg, M.A.; Moresco, J.J.; Ghanbari Niaki, A.; Roach, N.P.; Yates, J.R.; Myong, S.; Kim, J.K. RNA Promotes Phase Separation of Glycolysis Enzymes into Yeast G Bodies in Hypoxia. Elife 2020, 9, e48480.
  346. Sarkar, S.; Mondal, J. Mechanistic Insights on ATP’s Role as a Hydrotrope. J. Phys. Chem. B 2021, 125, 7717–7731.
  347. Maldonado, E.N.; Lemasters, J.J. ATP/ADP Ratio, the Missed Connection between Mitochondria and the Warburg Effect. Mitochondrion 2014, 19 Pt A, 78–84.
  348. Bell, S.M.; Burgess, T.; Lee, J.; Blackburn, D.J.; Allen, S.P.; Mortiboys, H. Peripheral Glycolysis in Neurodegenerative Diseases. Int. J. Mol. Sci. 2020, 21, 8924.
  349. Lu, J.; Qian, J.; Xu, Z.; Yin, S.; Zhou, L.; Zheng, S.; Zhang, W. Emerging Roles of Liquid-Liquid Phase Separation in Cancer: From Protein Aggregation to Immune-Associated Signaling. Front. Cell Dev. Biol. 2021, 9, 631486.
  350. Petronilho, E.C.; Pedrote, M.M.; Marques, M.A.; Passos, Y.M.; Mota, M.F.; Jakobus, B.; de Sousa, G.D.S.; Pereira da Costa, F.; Felix, A.L.; Ferretti, G.D.S.; et al. Phase Separation of p53 Precedes Aggregation and Is Affected by Oncogenic Mutations and Ligands. Chem. Sci. 2021, 12, 7334–7349.
  351. Gargini, R.; Segura-Collar, B.; Sánchez-Gómez, P. Novel Functions of the Neurodegenerative-Related Gene Tau in Cancer. Front. Aging Neurosci. 2019, 11, 231.
  352. Watkins, K.P.; Williams-Carrier, R.; Chotewutmontri, P.; Friso, G.; Teubner, M.; Belcher, S.; Ruwe, H.; Schmitz-Linneweber, C.; van Wijk, K.J.; Barkan, A. Exploring the Proteome Associated with the mRNA Encoding the D1 Reaction Center Protein of Photosystem II in Plant Chloroplasts. Plant J. 2020, 102, 369–382.
  353. Zoschke, R.; Bock, R. Chloroplast Translation: Structural and Functional Organization, Operational Control, and Regulation. Plant Cell 2018, 30, 745–770.
  354. Gawroński, P.; Enroth, C.; Kindgren, P.; Marquardt, S.; Karpiński, S.; Leister, D.; Jensen, P.E.; Vinther, J.; Scharff, L.B. Light-Dependent Translation Change of Arabidopsis psbA Correlates with RNA Structure Alterations at the Translation Initiation Region. Cells 2021, 10, 322.
  355. Preiss, S.; Schrader, S.; Johanningmeier, U. Rapid, ATP-Dependent Degradation of a Truncated D1 Protein in the Chloroplast. Eur. J. Biochem. 2001, 268, 4562–4569.
  356. Pévet, P. Melatonin in Animal Models. Dialogues Clin. Neurosci. 2003, 5, 343–352.
  357. Zheng, X.; Tan, D.X.; Allan, A.C.; Zuo, B.; Zhao, Y.; Reiter, R.J.; Wang, L.; Wang, Z.; Guo, Y.; Zhou, J.; et al. Chloroplastic Biosynthesis of Melatonin and Its Involvement in Protection of Plants from Salt Stress. Sci. Rep. 2017, 7, 41236.
  358. Hwang, O.J.; Kang, K.; Back, K. Effects of Light Quality and Phytochrome Form on Melatonin Biosynthesis in Rice. Biomolecules 2020, 10, 523.
  359. Zhou, X.; Zhao, H.; Cao, K.; Hu, L.; Du, T.; Baluška, F.; Zou, Z. Beneficial Roles of Melatonin on Redox Regulation of Photosynthetic Electron Transport and Synthesis of D1 Protein in Tomato Seedlings under Salt Stress. Front. Plant Sci. 2016, 7, 1823.
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