The Role Of PPAR Alpha in The Modulation Of Innate Immunity Part 2
Apr 21, 2023
5. PPARα’s Role in the Innate Immunity Effector Processes: ROS/RNS Production
An important component of innate immunity in animals is the generation of active forms of oxygen (mainly superoxide) and active forms of nitrogen, mainly nitric oxide and its derivatives [102]. The form of nitric oxide synthase (NOS) traditionally associated with inflammation is the so-called inducible nitric oxide synthase (iNOS or NOS 2). NOS 2 belongs to the enzymatic family of nitric oxide synthases (NOS), being the evolutionarily most distant member of the family. NOS 2 may be expressed in numerous types of cells and tissues [103].
The other two, NOS 1 and NOS 3, also called ‘constitutive’ or Ca2+-dependent enzymes, are present constitutively in many tissues and cells of the organism, mainly but not solely in some neurons (NOS 1), as well as endothelial cells (NOS 3) [104]. They generate a lower level of NO than NOS 2, despite their comparable enzymatic activity in vitro [102]. Importantly, under various conditions, all NOS enzymes are a source of active forms of nitrogen and oxygen; in the absence of L-arginine, they simply produce superoxide and may be an important source of oxidative/nitrosative stress [105].
There is a close relationship between enzyme activity and immunity. Enzymes are important protein molecules in living organisms. They can accelerate the speed of chemical reactions and play an important role in physiological processes. Many molecules in the immune system, including antibodies, cytokines, and enzymes, play a role in modulating the immune response. We should also pay attention to our immunity in daily life. Cistanche can enhance immunity. Cistanche is rich in various antioxidant substances, such as vitamin C, vitamin C, carotenoids, etc. These ingredients can scavenge free radicals, reduce oxidative stress, and improve immunity. system resistance.
PPARα agonists may downregulate NOS 2 [106,107], while they stimulate both NOS 3 [108], which plays a protective role in the cardiovascular system and NOS 1 (see [109,110]). NOS 2 is expressed de novo under the influence of proinflammatory factors [102], and, as it is not dependent on calcium, it can only be downregulated by inhibition of the enzymatic activity or proteolytic degradation of the enzyme. NOS activity also depends on the competition with the alternate substrate consumer arginase, which produces urea and L-ornithine instead of L-citrulline and nitric oxide [111,112].
The possibility of switching the main path of L-arginine metabolism from the generation of NO and citrulline to the generation of urea and ornithine is a basis for the functional diversification of M1 and M2 macrophages. M1 macrophages, unlike M2 macrophages, generate free radicals and are the proinflammatory type of these cells (as mentioned in Section 3). They contribute to the development of inflammation-driven tumors [107]. PPARα, as an attenuator of inflammation and free-radical production, acts in this case as an antitumor agent. Parallel to tumor progression and diversification of the tumor macrophage phenotype toward M2, the situation becomes more ambiguous and unpredictable. The actual effect of activation of PPARα depends on the type of tumor and its phase of development [108]. Indeed, fenofibrate inhibited the development of micrometastases of melanoma BHM in Syrian hamster lungs, but did not affect the kinetics of the primary tumor growth, nor the progression of macro-metastases [113]. It must be added that, recently, particular attention has been paid to the possibility of manipulation of NOS 2 activity by its selective inhibitors to achieve a desirable level of human monocyte physiological response [114].
The second mechanism of innate defense that involves the production of highly reactive small chemical molecules is a respiratory (or oxidative) burst carried out by phagocytes. PPARα agonists were shown to increase macrophage microbicidal activity through intensification of ROS production during respiratory burst. This was caused by PPARα-dependent elevated expression of crucial transmembrane (gp91phox) and cytosolic (p47phox and p67phox) components of NADPH oxidase [115]. Interestingly, increased ROS production led to the generation of oxidized low-density lipoproteins (ox-LDL), which further stimulated PPARα activation. Activated PPARα downregulated NO production via transrepression of iNOS [115]. This is an example of PPARα differently regulating various innate immunity effector molecules, in this case, ROS and RNS. An unexpectedly interesting transcriptional regulation occurs in the promoter of another gene crucial for the generation of reactive species during the respiratory burst, namely, myeloperoxidase (MPO). The human promoter of this gene contains primate-specific Alu elements that are repetitive DNA mobile fragments spread throughout the human genome in about 1 million copies [116]. The Alu fragment in the MPO gene promoter contains four hexamer sequences identical to or closely resembling canonical PPAR response elements (PPREs): AGGTCA, with 2 or 4 bp spacing between them [117].
The third and fourth hexamers serve as PPREs and accommodate PPARα/RXR or PPARγ/RXR heterodimers, which enables transcriptional regulation by PPAR ligands. Surprisingly, MPO expression is regulated by PPARα agonist GW9578 and PPARγ agonist MCC-555 in opposite directions in human macrophages, depending on the differentiation pathway; MPO is significantly downregulated in macrophages derived from MG-CSF-treated monocytes and upregulated in M-CSF differentiated cells [117]. The difference could probably be attributed to the differential utilization of nuclear co-repressors, such as NCoR or silencing mediator of retinoid and thyroid receptors (SMRT), in macrophages differentiated with GM- vs. M-DAMP [117]. Notably, such a mode of regulation is entirely human-specific, because mice do not possess Alu elements in their genome.
6. PPARα as an Immunomodulator during Infections
Truly immunomodulatory action does not lie in the unilateral inhibition or activation of all inflammatory processes but in selective influence on the chosen aspects of innate immunity. Such an immunomodulatory action of PPARα has been observed in parasitic or microbial infections. One example of such activity relates to the induction of M2 polarization in macrophages of patients infected with Trypanosoma cruzi, a parasitic euglenoid, which is responsible for Chagas disease development. The experiment carried out on the infected mice showed that PPARα agonist Wy-14643 elevated the expression of M2 macrophage markers, arginase-1, mannose receptor (CD206), Ym1, and TGFβ, and decreased the production of proinflammatory molecules characteristic of the M1 phenotype, such as iNOS, NO, IL-1β, IL-6 and TNFα [118]. However, this phenotypic switch was accompanied by a PPARα (but not PPARγ)-dependent increase in phagocytic capacity and efficiency of parasite phagocytosis [118]. These results indicate that PPARα activation might have therapeutic significance, because its immunomodulatory action, on the one hand, strengthens macrophage effector capacity, but, on the other hand, helps to alleviate severe chronic inflammation associated with Chagas disease, which is destructive to various organs.
Similar immunomodulatory activity of PPARα in the context of phagocytosis was described in primary peritoneal macrophage and microglia cultures treated with several PPARα agonists: endogenous cannabinomimetic (see below), PEA, fenofibrate, or palmitic acid [119]. These compounds, particularly PEA, significantly enhanced phagocytosis and intracellular killing of E. coli by macrophages and microglial cells. Although PEA pretreatment reduced the levels of proinflammatory cytokines (IL-1β, IL-6, and TNFα) and chemokines (CXCL1) in the tissues of mice subjected to intracerebellar or intraperitoneal E. coli infection, it induced a very effective bacterial clearance from blood, spleens, and cerebellum, which translated into improved survival of these animals [119]. These results suggest a prophylactic potential of PPARα activation in the case of bacterial infections.
Another example illustrating that the exaggerated inflammatory response is not beneficial for the host is tuberculosis infection. In this case, PPARα’s immunomodulatory and metabolic roles are connected, leading to a better outcome for wt mice infected with mycobacteria (Bacillus Calmette–Guerin or M. tuberculosis) in comparison with PPARα KO mice [120].
The absence of PPARα resulted in more rapidly increasing intracellular bacterial load in macrophages, heavier bacteremia in the lungs, spleen, and liver, and a significantly higher level of inflammatory cytokines TNFα and IL-6 in the lungs, as compared to wt PPARα mice. The exaggerated inflammatory response was associated with a higher number of granuloma lesions in the lungs of PPARα KO mice. Granuloma lesions are the manifestation of unsuccessful host defense against mycobacteria because they are full of dead leukocytes, damaged lung tissue multinucleated giant cells, and macrophages converted to foam cells, filled with lipid-containing vesicles, which create a favorable energy source for surviving and proliferating mycobacteria [121]. Pharmacological PPARα agonists, GW7647 and Wy-14643 induced phagosomal maturation through activation of transcription factor EB (TFEB) and significantly reduced the survival of intracellular bacteria, which resulted from increased fatty-acid β-oxidation and elimination of lipid-rich bodies [120]. This is an example of the interconnection between PPARα-mediated lipid catabolism and its immunomodulating effects, which support the effective antimicrobial innate defense.
Despite a large body of evidence documenting the beneficial outcomes of PPARα activation in various diseases with an inflammatory background, there are also certain conditions in which PPARα-mediated immunomodulation is hazardous. The illustrative example is a situation where, after a viral influenza infection, a subsequent bacterial (e.g., staphylococcal) superinfection occurs. Antibiotic-resistant Staphylococci are a frequent cause of life-threatening nosocomial infections in patients hospitalized due to viral pulmonary infections. Tam and colleagues [122] found out that the presence of PPARα was responsible for a more severe course of superinfection and higher mortality in wt mice as compared to PPARα KO mice. A viral infection that was induced before the challenge with S. aureus led to increased PPARα expression in the lungs. Moreover, the lipidomic analysis of bronchoalveolar lavage fluid from infected mice revealed that superinfection resulted in a significant enrichment of several inflammatory lipid mediators, such as LOX product LTE4 and CYP450 products 11,12-dihydroxyeicosatrienoic acid (11,12-diHETrE) and 14,15-diHETrE, as compared to single infection, whether viral or bacterial. 14,15-diHETre is a very potent PPARα agonist [123].
The inhibition of NF-κB signaling mediated by activated PPARα led to a blunted proinflammatory response to bacteria and loss of control over bacterial growth, which inflicted higher mortality [122]. Superinfection caused the decreased expression of macrophage inflammatory genes IL-1β, IL-6, CXCL5, and MMP-9, as well as a scavenger receptor Marco, which resulted in less efficient phagocytosis and heavier bacterial burden. Moreover, PPARα activation led to increased necroptosis (a programmed RIPK3 kinase-dependent lytic cell death), which was responsible for lung tissue damage and dramatically worsened the condition of infected animals [122].
The still scarce, but gradually emerging experimental data indicate that PPARα affects the innate host response to viral infections. Such involvement is beneficial in certain situations but could be detrimental in other conditions. The overexpression of PPARα homolog in a grouper fish (Epinephelus coioides, EcPPARα) blocked interferon- and NF-κBinduced cytokine expression during viral infections, which led to acute cytopathic injuries and heavier multiplicity of infection [124]. The topic of viral infection onset is currently very important due to its relationship with the ongoing COVID-19 pandemic. A study performed on primary human bronchial epithelial cells infected with SARS-CoV-2 revealed severe alterations in the gene transcription pattern that manifested endoplasmic reticular and mitochondrial stress, metabolic reprogramming toward intensive lipid synthesis and accumulation, impaired fatty acid oxidation, and upregulated aerobic glycolysis via activation of the NF-κB pathway [125].
Such a metabolic signature suggests that infection impairs PPARα signaling. Therefore, the restoration of PPARα activity could be beneficial through a reversal of these changes and metabolic ‘repair’. Indeed, the treatment of the infected cell cultures with PPARα ligand fenofibrate alleviated the dysregulation of lipid metabolism, blocked infection-induced phospholipid accumulation, and remarkably decreased viral load by 100-fold within 3 days and 1000-fold within 5 days [125]. These results seem to support the hypothesis that fenofibrate treatment could alleviate the acute infection symptoms during COVID-19 by supporting fatty-acid metabolism in alveolar epithelial cells, improving pulmonary endothelial cell function, and calming down the cytokine storm, leading to a better outcome for the patients [126].
7. Interplay between PPARα and the Endocannabinoid System: Implications for Inflamma-Tion, Neuroprotection, and Analgesia
7.1. Analgesic Lipid Mediators as PPARα Agonists
Mechanical tissue damage, hypersensitivity reactions, or local infection result in inflammation, which evokes a nociceptive response and pain. Pain signals are elicited by proalgesic lipid mediators, such as lysophospholipids and PDE2, or hydroxylated derivatives of linoleic acid (e.g., 13-hydroxy octadecanoic acid, 13-HODE), which increase the excitability of nociceptive neurons [127].
Nevertheless, another group of endogenous lipid mediators possesses the opposite, analgesic activity. Acting through cannabinoid receptors CB1 and/or CB2, they mitigate the excitability of sensory nociceptive neurons. This is a part of the so-called endocannabinoid system, which includes the ligands N-arachidonoylethanolamine (AEA, anandamide) and 2-arachidonoyl-glycerol (2-AG), which were first discovered, and their receptors, cannabinoid receptors CB1 and CB2 expressed in the CNS and immunocompetent cells, respectively, as well as TRPV1 and endocannabinoid-synthesizing and -degrading enzymes [128,129]. Later, other fatty-acid ethanolamides (FAEs), such as N-palmitoylethanolamide (PEA) and N-oleoyl ethanolamide (OEA), were detected in mammalian and invertebrate tissues [130–132]. OEA and PEA are biologically relevant and potent PPARα agonists, with EC50 values of 0.12 µM and 3 µM, respectively [44,133], which links PPARα with the endocannabinoid system. Numerous biological hormone-like functions of OEA and PEA are widely known, including analgesic and anti-nociceptive cannabinomimetic activities, although they are not bona fide CB1 or CB2 agonists [134]. Endocannabinoids and cannabimimetic are synthesized on demand from membrane phospholipids, but can also be accumulated intracellularly in lipid droplets [135,136]. They are abundantly present in the brain, leukocytes, gastrointestinal tract, and other tissues [137–139].
The most common FAE biosynthesis route involves the formation of N-acylphosphatidylethanolamine from phosphatidylethanolamine by calcium-dependentN-acyl-transferase and subsequent conversion to N-acyl-ethanolamine by N-acyl-phosphatidylethanolamine hydrolyzing phospholipase D (NAPE-PLD) [140]. Several other biosynthesis pathways that engage other phospholipases and glycerophosphodiesterases are also possible (for a review, see [128]). Endocannabinoids are absorbed by cells and metabolized by intracellular fatty-acid amide hydrolase (FAAH) or N-acylethanolamine-hydrolyzing acid amidase (NAAA) [141].
OEA and PEA exert analgesia and reduce nociception in various animal models of inflammatory pain [142,143]. PEA and synthetic PPARα ligands (GW7647, Wy-14634, perfluorooctanoic acid) produce analgesic effects and strongly reduce edema in chemically induced models of inflammation [142,144–146]. Although, in some cases, OEA acted independently of PPARα presence [143], PEA-induced nociception and anti-inflammatory actions were exerted through PPARα [142,145].
Importantly, PEA-mediated activation of PPARα in CNS through intracerebroventricular PEA application was able to reduce the peripheral inflammatory response (paw edema after carrageenan injection) [146]. This demonstrated a distant endocrine action of PEA, despite the molecular mechanism involving inhibition of the NF-κB signaling pathway in CNS tissue [146]. A PPARα involvement was also demonstrated in the experiments with a synthetic PPARα agonist GW7647, which induced synergistic enhancement of AEA analgesic properties in a chemically induced inflammatory pain model [145,147]. The antinociceptive action of GW7647 depended on the activity of large conductance potassium channels, which further supported the involvement of the endocannabinoid system [145,147]. The potentiation of endocannabinoid binding to CB1 and CB2 receptors by cognate molecules, which are not agonists themselves, was observed and named ‘the entourage effect’ [148].
In the case of AEA, PEA, and OEA, such an effect could be explained by FAAH engagement in PEA and OEA hydrolysis, sparing the large pool of AEA from degradation and allowing it to activate CB receptors. Indeed, the entourage effect has been described as an enhanced vasodilation activity of AEA through TRPV1 by PEA and OEA in the endothelium [149]. In summary, all these results indicate that PPARα signaling contributes to inflammatory pain control through cannabinomimetics OEA and PEA (Figure 3) [127].
7.2. PPARα Involvement in Resolution of Neuroinflammation
The presence of OEA and PEA in CNS implicates their activity in the physiology of neurons and glial cells. Both compounds were shown to exert beneficial effects by counteracting the glial inflammatory responses and by providing cytoprotection over neuronal cells and their activities in various neuropathic states. Neuroinflammation and exaggerated glial reactivity are associated with numerous neurodegenerative diseases, traumatic injuries, ischemia/reperfusion stress, and neuropathic pain [150–152]. The brain is regarded as ‘an immune-privileged organ, protected from peripheral proinflammatory stimuli by the blood–brain barrier, but microglia, astrocytes, and mast cells are capable of triggering neuroinflammation [153]. Aberrant or chronic activation of these cells in the CNS leads to increased expression of TLRs, cytokines (TNFα, IL-6), chemokines (CXCL6) metalloproteinases, ROS, and RNS, which results in the loss of calcium homeostasis, neuronal damage, or apoptosis [151–153]. The potential of lipid amides, called ALIAmides (autacoid local injury antagonists) to counteract neurogenic inflammation and mast-cell degranulation, was proposed by Rita Levi-Montalcini, a Nobel laureate (1988), for her discoveries in the field of neurobiology [154].
Indeed, numerous studies demonstrated that OEA and PEA, classified as ALIAmides, could provide neuroprotection via the downregulation of inflammatory responses in the brain through modulation of glial cell functions. Benito and colleagues discovered that N-fatty acyl ethanolamines (OEA, PEA, AEA) and synthetic agonists of PPARα (Wy-14643) and PPARγ (troglitazone) alleviate the inflammatory response induced by the treatment of astrocytes with β-amyloid peptide fragments [155]. The anti-inflammatory effects were mediated by PPARα, PPARγ, and TRPV1 activity, but not through CB1 or CB2 [155]. The neuroprotective action of PEA and an endocannabinoid 2-AG was observed in an excitatory model of neuronal damage in organotypic hippocampal slice cultures [156]. PEA and 2-AG rescued about 50% of neurons from NMDA-induced cell death, acting on microglial cells, albeit through different and mutually suppressing mechanisms. PEA blocked microglial inflammatory activities, such as NO production and the acquisition of ameboid morphology, characteristic of an activated condition [156]. These effects were associated with PPARα nuclear translocation, which suggests its involvement in the process.
7.3. PPARα-Mediated Regulation of Microglia and Macrophage Functions
The glia-directed activity of PEA was studied by Scuderi and coauthors, who, in a series of papers, demonstrated that PEA or synthetic PPARα agonists, in a PPARα-dependent manner, decreased markers of glial inflammation and improved neuronal viability in animal models of Alzheimer’s disease, as well as in mixed glial-neuronal cell cultures and organotypic neural cultures [157–159]. The immunomodulatory activity of PEA and the interplay between PPARα and the endocannabinoid system were also analyzed in primary microglial and macrophage cultures [160]. This study revealed that CB2 mRNA and protein levels were significantly increased by the treatment with PEA and a synthetic PPARα agonist GW7647, and this effect was evoked by the PPARα/RXR heterodimer binding to the promoter and transactivation of the gene encoding CB2 [160]. PEA-induced microglial effector functions in a PPARα-dependent manner and improved the phagocytosis and killing of Porphyromonas gingivalis by microglia and chemotaxis to 2-AG [160].
In addition to the modulation of antimicrobial phagocytosis-based defense, PEA can modulate regenerative functions of macrophages, such as efferocytosis (i.e., phagocytosis and clearance of apoptotic cells) [161]. PEA is produced endogenously by M2c-polarized but not M1-polarized macrophages [161]. Exogenous chronic administration of PEA limited early plaque formation, protected from the accumulation of the proinflammatory M1 macrophage within the plaque, and promoted efferocytosis by M2a- and M2c-polarized macrophages, which delayed the onset of arteriosclerosis [161]. These results show that endogenous PPARα ligand PEA is capable of modulating microglia and macrophage biological functions.
7.4. PPARα’s Role in the Restoration of Neural Function after Injury or Infection
Neuroprotective OEA activity was also demonstrated as an inhibition of so-called glial scar (i.e., zones enriched with reactive inflammatory astrocytes, microglia, fibroblasts, and accumulated extracellular matrix components) formation, after focal cerebral ischemia injury [162]. Glial scar is a natural physiological reaction to injury, but it impedes neurite formation, axon regrowth, and recovery after a brain stroke. OEA increased PPARα expression in the cerebral cortex and downregulated glial scar markers (S100B, glial fibrillary acidic protein GFAP, metalloproteinases MMP-2, MMP-9, and neurocan) in the ischemic region through a PPARα-dependent mechanism [162]. Importantly, these biological processes translated into a better recovery of motor function in mice after stroke [162]. OEA also decreases the inflammatory response of endothelial cells (such as IL-6, IL-8, ICAM-1, and VCAM expression) evoked by TNFα, in a PPARα- and CB2-dependent manner [163].
The biological activities of OEA and PEA seem similar and sometimes overlap, but are not always identical, as shown in different experimental settings. An intriguing difference between OEA and PEA actions was observed in a study that analyzed functional impairments of neurological functions in an animal model of neonatal anoxia/ischemia-induced brain injury [164]. PEA, but not OEA treatment was capable of limiting hippocampal astrogliosis markers (e.g., ionized calcium-binding adaptor protein Iba-1, GFAP) and restoring PPARα protein expression in anoxia/ischemia-affected brain regions [164]. These effects were associated with improved cognitive abilities and better recovery of spatial and recognition memory, as compared to control animals subjected to anoxia/ischemia [164]. Nevertheless, OEA was proved effective in ameliorating cognitive deficits and in supporting neurogenesis in ischemia-affected brain regions of rats subjected to middle cerebral artery occlusion [165].
An important immunomodulatory action of OEA and PEA involves TLR3 signaling during the innate response to viral infections. A recent report by Flannery et al. [166] demonstrated that intracerebroventricular administration of a TLR3 ligand, viral mimetic polyinosinic–polycytidylic acid (poly I: C), led to the induction of hypothalamic interferon and NF-κB-regulated pathways of proinflammatory gene expression and hyperthermia. The treatment with both OEA and PEA attenuated TLR3-mediated hyperthermia, but only OEA (not PEA) was effective in the downregulation of poly I: C-induced inflammatory gene expression, including TNFα, iNOS, IL-1β, COX-2, interferon gamma-induced protein 10 (IP-10), and interferon-regulated factor IRF7. The fact that the PPARα antagonist GW6471 attenuated these effects indicated the PPARα involvement in this regulation [166]. These results have important implications for the current pandemic of SARS-CoV-2 infections, which often cause complications within the CNS, manifested by neurological and mental disorders, such as impaired memory, attention, anxiety, depression, and dementia [167].
7.5. PPARα and Endocannabinoid Involvement in the Regulation of Mast-Cell Functions
Mast cells are important innate immune cells that, due to their rapid degranulation, can control the onset of inflammation in various tissues. PEA was shown to reduce local accumulation and the activation of mast cells in various inflammatory models: (i) after substance P injection to ear pinna [154], (ii) during chemically induced allergic dermatitis in mice [168], (iii) in myelin basic protein (MBP)-induced neuronal injury in a neuron–glia– mast cell coculture model of multiple sclerosis [169], (iv) in rat mast cell line RBL-2H3 [170], (v) after ischemia/reperfusion inflammatory injury of the intestine after splanchnic artery occlusion in mice [171], and (vi) during chemically induced colitis which serves as an animal model of inflammatory bowel disease [172].
In all these experimental models, PEA suppressed a variety of effector reactions produced by mast cells or other leukocytes, such as chemotaxis, degranulation, enzyme release, and induction of proinflammatory cytokines. This suppression of mast-cell activity led to the alleviation of inflammatory tissue damage and improved physiological tissue function. A common molecular mechanism could be involved in these effects, because, regardless of the model used, they were mediated, at least partially, by PPARα and CB2 activation [168–170], as well as, in some cases, by GPR55 and TRPV1 [172], which further supports the role of PPARα in the modulation of innate immunity and its connections with the endocannabinoid system.
However, a very intriguing recent discovery has shed new light on the connection among cannobinomimetics, mast cells, and metabolism, namely, ketogenesis. The publication from Daniele Piomelli’s group revealed the unexpected role of histamine secreted by mast cells as a mediator necessary to induce ketogenesis in the liver in the state of food deprivation [173]. The mode of metabolic regulation involves an OEA-mediated action on hepatocytes. Routinely, after feeding, OEA is produced in the small intestine from consumed dietary lipids and takes part in food intake control as a satiety mediator via PPARα activation [133,174]. However, during food deprivation, ketogenesis depends on liver-derived OEA. A crucial role in this process is played by a population of mast cells that reside in the gastrointestinal tract and release histamine in the fasting state. Histamine enters the liver through portal circulation and stimulates hepatocytes to OEA secretion via activation of histamine H1 receptors [173]. Furthermore, OEA binding to PPARα in hepatocytes activates the transcription of PPARα-target genes that control ketogenesis, including ACAT1, HMGSC2, and Fgf21 [173]. These results provide a novel link between mast cells as innate immunity effectors, cannabinomimetic PPARα ligand OEA, and PPARα-dependent ketogenesis as a metabolic response to fasting.
8. Evolutionary Aspects of PPARα-Mediated Immunomodulation
One of the crucially important features of the innate response is the speed and immediateness of the reaction to menacing invaders. In higher vertebrates, the accurate and prompt launching of the innate mechanisms buys time for the preparation of systemic adaptive immunity. In invertebrates, the effectiveness of innate immunity is a matter of life and death. The precise regulation of the innate responses is a multithreaded process that engages various signaling pathways, including the activity of nuclear receptors, such as PPARs. Such a regulation determines the success in coping with parasitic, viral, and bacterial infections, in addition to providing a hospitable environment for commensal microbiota and restricting inflammation-related tissue damage and injury.
PPARs and NOS serve as illustrative examples of how the elements of innate immunity and their regulatory mechanisms coevolved in the animal kingdom. On the one hand, NOS belongs to a large family of evolutionarily ancient enzymes that includes numerous pro- and eukaryotic flavodoxins [175,176]. There have been several hypotheses of their reciprocal relationship in invertebrates in the function of hemolymph homeostasis maintenance and the destruction of pathogens, i.e., probably unified in hemocyte NOS, as is the case for horseshoe crabs [175,177].
On the other hand, PPARs, despite their origin in the nuclear receptor family that emerged in metazoans, evolved in animals only as late as in the branch of Deuterostomata, whereas, in chordates, their presence dates from the evolution of Branchiostomata [178]. Consequently, they are present in all vertebrates, but (except for Branchiostomata) absent in invertebrates [178]. Their presence seems to correspond to the evolution of the immune system and adipose tissue, but their tissue specificity does not overlap with their functional diversification. The most basic branch of this family seems to be represented by PPARγ, and the evolution of the whole family comprised two duplications of the genes, the first moving PPARγ apart, and the other dividing the other group into the PPARβ and α subfamilies [179]. This must have taken place on the level of ancient, primitive Teleostei [178,179].
Meanwhile, the diversified NOS family tree must root as deeply as in some Protista, as present in a differentiated side-branch in slime molds, fungi, and practically all Eukaryota including (a loosely related variant) high plants (Arabidopsis thaliana [180]). This may explain the engagement of PPARs in the functioning of various NOS in vertebrates. Upon evolution, the diversification of the NOS family has been consistently appreciated, whereas the engagement of PPARs in various aspects of NOS functioning may have been more or less accidental (Figure 4).
9. Conclusions and Perspectives
PPARα as a transcription factor exerts a strong impact on cellular metabolism and intracellular signal transduction events, which alters the physiology and behavior of PPARα-expressing cells of both immune and nonimmune provenance. These physiological alterations underlie the immunomodulatory actions of PPARα presented in previous chapters. The broad spectrum of actions of endogenous and pharmacological PPARα agonists directed toward the immune system encourages the development of more commonly used therapeutic applications of PPARα-targeted solutions in various infectious diseases and disorders of immunological background. The currently ongoing SARS-CoV-2 pandemic has created a dire need to revise the canonical approaches to the treatment of viral infections and has opened an unexpected possibility for new attempts, such as applying PPARα agonists to calm down the destructive cytokine storm in severe COVID-19 cases.
Author Contributions:
Conceptualization, M.G.; literature survey and discussions on the topic, M.G., M.P., P.M.P., and P.P.; writing—original draft preparation, M.G., M.P., P.M.P., and P.P.; writing—review and editing, M.G., M.P., P.M.P., and P.P.; figure preparation, M.G. All authors have read and agreed to the published version of the manuscript.
Funding:
This research was funded by N43/DBS/000158 to P.P.
Conflicts of Interest:
The authors declare no conflict of interest.
Abbreviations
2-AG, 2-arachidonoyl-glycerol; ACAT1, acetoacetyl-CoA thiolase 1; AEA, N-arachidonoylethanolamine; AMPs, antimicrobial peptides; AP-1, activation protein 1; CB, cannabinoid receptors; CLRs, C-type lectin receptors; COX, cyclooxygenase; CSF, colony-stimulating factor; DAMPs, damage-associated molecular patterns; DOPA, dihydroxyphenylalanine; FAAH, fatty-acid amide hydrolase; FAEs, fatty-acid ethanolamides; FAO, fatty-acid oxidation; FGF21, fibroblast growth factor 21; FREPs, fibrinogen-related proteins; HETE, hydroxy eicosatetraenoic acid; HMGCS2, 3-hydroxy-3-methylglytaryl-CoA synthetase 2; HPETE, hydroperoxy eicosatetraenoic acid; IDO, indoleamine-2,3-dioxygenase; IL, interleukin; ILCs, innate lymphoid cells; IRF, interferon-regulated factor; JAK, Janus activated kinase; JNK, c-Jun N-terminal kinase; KO, knockout; LOX, lipoxygenase; LPS, lipopolysaccharide; LT, leukotriene; MAMPs, microbial-associated molecular patterns; MBP, myelin basic protein; MCP1, monocyte chemoattractant protein 1; MDSCs, myeloid-derived suppressor cells; MMP-9, matrix metalloproteinase 9; NAAA, N-acylethanolaminehydrolyzing acid amidase; NAPE-PLD, N-acyl-phosphatidylethanolamine-hydrolyzing phospholipase D; NCoR, nuclear receptor co-repressor; NF-κB, nuclear factor κB; NLR, nucleotide-binding oligomerization domain (NOD)–leucin-rich repeat (LRR)-containing receptors; NO, nitric oxide; NOD, nucleotide-binding oligomerization domain; NOS, nitric oxide synthase; OEA, oleylethanolamide; PAMPs, pathogen-associated molecular patterns; PEA, palmitoylethanolamide; PG, prostaglandin; PPAR, peroxisome proliferator-activated receptor; PPRE, peroxisome proliferator response element; PRRs, pattern-recognition receptors; RIG1, retinoic acid inducible gene 1; RLR, retinoic acid inducible gene 1 (RIG1)-like receptors; RNS, reactive nitrogen species; ROR, retinoid orphan receptor; ROS, reactive oxygen species; RXR, retinoid X receptor; SAPK, stress-activated protein kinase; SMRT, silencing mediator of retinoid and thyroid receptors; STAT, signal transducer and activator of transcription; TF, tissue factor; TFEB, transcription factor EB; TGF, transforming growth factor; TLR, Toll-like receptors; TNF, tumor necrosis factor; TRPV1, transient receptor potential cation channel vanilloid subfamily member 1; TXNIP, thioredoxin-interacting protein.
References
1. Boraschi, D.; Italiani, P. Innate immune memory: Time for adopting a correct terminology. Front. Immunol. 2018, 9, 799. [CrossRef]
2. Netea, M.G.; Quintin, J.; van der Meer, J.W. Trained immunity: A memory for innate host defense. Cell Host Microbe 2011, 9, 355–361. [CrossRef]
3. Moret, Y.; Siva-Jothy, M.T. Adaptive innate immunity? Responsive-mode prophylaxis in the mealworm beetle, Tenebrio molitor. Proc. Biol. Sci. 2003, 270, 2475–2480. [CrossRef]
4. Torre, C.; Abnave, P.; Tsoumtsa, L.L.; Mottola, G.; Lepolard, C.; Trouplin, V.; Gimenez, G.; Desrousseaux, J.; Gempp, S.; Levasseur, A.; et al. Staphylococcus aureus promotes Smed-PGRP-2/Smed-setd8-1 methyltransferase signaling in planarian neoblasts to sensitize anti-bacterial gene responses during re-infection. EBioMedicine 2017, 20, 150–160. [CrossRef] [PubMed]
5. Zhang, T.; Qiu, L.; Sun, Z.; Wang, L.; Zhou, Z.; Liu, R.; Yue, F.; Sun, R.; Song, L. The specifically enhanced cellular immune responses in Pacific oysters (Crassostrea gigas) against secondary challenge with Vibrio splendidus. Dev. Comp. Immunol. 2014, 45, 141–150. [CrossRef]
6. Wang, X.; Zhao, Q.; Christensen, B.M. Identification and characterization of the fibrinogen-like domain of fibrinogen-related proteins in the mosquito, Anopheles gambiae, and the fruitfly, Drosophila melanogaster, genomes. BMC Genom. 2005, 6, 114. [CrossRef] [PubMed]
7. Melillo, D.; Marino, R.; Italiani, P.; Boraschi, D. Innate immune memory in invertebrate metazoans: A Critical appraisal. Front. Immunol. 2018, 9, 1915. [CrossRef]
8. Cerenius, L.; Soderhall, K. Immune properties of invertebrate phenoloxidases. Dev. Comp. Immunol. 2021, 122, 104098. [CrossRef]
9. Coates, C.J.; Nairn, J. Diverse immune functions of hemocyanins. Dev. Comp. Immunol. 2014, 45, 43–55. [CrossRef] [PubMed]
10. Yang, D.; Han, Z.; Oppenheim, J.J. Alarmins and immunity. Immunol. Rev. 2017, 280, 41–56. [CrossRef]
11. Amarante-Mendes, G.P.; Adjemian, S.; Branco, L.M.; Zanetti, L.C.; Weinlich, R.; Bortoluci, K.R. Pattern Recognition Receptors, and the Host Cell Death Molecular Machinery. Front. Immunol. 2018, 9, 2379. [CrossRef] [PubMed]
12. Olive, C. Pattern recognition receptors: Sentinels in innate immunity and targets of new vaccine adjuvants. Expert Rev. Vaccines 2012, 11, 237–256. [CrossRef]
13. Rabinovitch, M. Professional, and non-professional phagocytes: An introduction. Trends Cell Biol. 1995, 5, 85–87. [CrossRef]
14. Lim, J.J.; Grinstein, S.; Roth, Z. Diversity and versatility of phagocytosis: Roles in innate immunity, tissue remodeling, and homeostasis. Front. Cell Infect. Microbiol. 2017, 7, 191. [CrossRef]
15. Witko-Sarsat, V.; Descamps-Latscha, B. Neutrophil-derived Oxidants and proteinases as immunomodulatory mediators in inflammation. Mediat. Inflamm. 1994, 3, 257–273. [CrossRef] [PubMed]
16. Nguyen, G.T.; Green, E.R.; Mecsas, J. Neutrophils to the ROScue: Mechanisms of NADPH oxidase activation and bacterial resistance. Front. Cell Infect. Microbiol. 2017, 7, 373. [CrossRef] [PubMed]
17. Kettle, A.J.; Winterbourn, C.C. Myeloperoxidase: A key regulator of neutrophil oxidant production. Redox Rep. 1997, 3, 3–15. [CrossRef]
18. Prolo, C.; Alvarez, M.N.; Radi, R. Peroxynitrite, a potent macrophage-derived oxidizing cytotoxin to combat invading pathogens. Biofactors 2014, 40, 215–225. [CrossRef]
19. Yamasaki, R.; Lu, H.; Butovsky, O.; Ohno, N.; Rietsch, A.M.; Cialic, R.; Wu, P.M.; Doykan, C.E.; Lin, J.; Cotleur, A.C.; et al. Differential roles of microglia and monocytes in the inflamed central nervous system. J. Exp. Med. 2014, 211, 1533–1549. [CrossRef]
20. Schulz, D.; Severin, Y.; Zanotelli, V.R.T.; Bodenmiller, B. In-Depth characterization of monocyte-derived macrophages using a mass cytometry-based phagocytosis assay. Sci. Rep. 2019, 9, 1925. [CrossRef]
21. Gerner, E.W.; Meyskens, F.L., Jr. Polyamines, and cancer: Old molecules, new understanding. Nat. Rev. Cancer 2004, 4, 781–792. [CrossRef] [PubMed]
22. Luk, G.D. Essential role of polyamine metabolism in hepatic regeneration. Inhibition of deoxyribonucleic acid and protein synthesis and tissue regeneration by difluoromethylornithine in the rat. Gastroenterology 1986, 90, 1261–1267. [CrossRef]
23. Murray, P.J. Macrophage polarization. Annu. Rev. Physiol. 2017, 79, 541–566. [CrossRef]
24. Locati, M.; Curtale, G.; Mantovani, A. Diversity, Mechanisms, and significance of macrophage plasticity. Annu. Rev. Pathol. 2020, 15, 123–147. [CrossRef] [PubMed]
25. Gasteiger, G.; D’Osualdo, A.; Schubert, D.A.; Weber, A.; Bruscia, E.M.; Hartl, D. Cellular innate immunity: An old game with new players. J. Innate. Immun. 2017, 9, 111–125. [CrossRef]
26. Erb, K.J.; Holloway, J.W.; Le Gros, G. Mast cells in the front line. Innate immunity. Curr. Biol. 1996, 6, 941–942. [CrossRef]
27. Sendo, S.; Saegusa, J.; Morinobu, A. Myeloid-derived suppressor cells in non-neoplastic inflamed organs. Inflamm. Regen. 2018, 38, 19. [CrossRef]
28. Killig, M.; Glatzer, T.; Romagnani, C. Recognition strategies of group 3 innate lymphoid cells. Front. Immunol. 2014, 5, 142. [CrossRef]
29. Trabanelli, S.; Gomez-Cadena, A.; Salome, B.; Michaud, K.; Mavilio, D.; Landis, B.N.; Jandus, P.; Jandus, C. Human innate lymphoid cells (ILCs): Toward a uniform immune-phenotyping. Cytom. B Clin. Cytom. 2018, 94, 392–399. [CrossRef]
30. Manoharan, I.; Suryawanshi, A.; Hong, Y.; Ranganathan, P.; Shanmugam, A.; Ahmad, S.; Swafford, D.; Manicassamy, B.; Ramesh, G.; Koni, P.A.; et al. Homeostatic PPARalpha signaling limits inflammatory responses to commensal microbiota in the intestine. J. Immunol. 2016, 196, 4739–4749. [CrossRef]
31. Sagebiel, A.F.; Steinert, F.; Lunemann, S.; Korner, C.; Schreurs, R.; Altfeld, M.; Perez, D.; Reinshagen, K.; Bunders, M.J. tissue-resident Eomes(+) NK cells are the major innate lymphoid cell population in the human infant intestine. Nat. Commun. 2019, 10, 975. [CrossRef]
32. Gordon, S.M.; Chaix, J.; Rupp, L.J.; Wu, J.; Madera, S.; Sun, J.C.; Lindsten, T.; Reiner, S.L. The transcription factors T-bet and Eomes control key checkpoints of natural killer cell maturation. Immunity 2012, 36, 55–67. [CrossRef] [PubMed]
33. Herbert, D.R.; Douglas, B.; Zullo, K. Group 2 innate lymphoid cells (ILC2): Type 2 immunity and helminth immunity. Int. J. Mol. Sci. 2019, 20, 2276. [CrossRef]
34. Hepworth, M.R.; Fung, T.C.; Masur, S.H.; Kelsen, J.R.; McConnell, F.M.; Dubrot, J.; Withers, D.R.; Hugues, S.; Farrar, M.A.; Reith, W.; et al. Immune tolerance. Group 3 innate lymphoid cells mediate intestinal selection of commensal bacteria-specific CD4(+) T cells. Science 2015, 348, 1031–1035. [CrossRef] [PubMed]
35. Withers, D.R.; Hepworth, M.R. Group 3 innate lymphoid cells: Communications hubs of the intestinal immune system. Front. Immunol. 2017, 8, 1298. [CrossRef]
36. Celsus, A.C. De Medicina, 1st ed.; Nicolaus Laurentii: Florence, Italy, 1478.
37. Frias, B.; Merighi, A. Capsaicin, nociception and pain. Molecules 2016, 21, 797. [CrossRef]
38. Nilius, B.; Owsianik, G. The transient receptor potential family of ion channels. Genome Biol. 2011, 12, 218. [CrossRef] [PubMed]
39. Desvergne, B.; Wahli, W. Peroxisome proliferator-activated receptors: Nuclear control of metabolism. Endocrinol. Rev. 1999, 20, 649–688. [CrossRef]
40. Sertznig, P.; Seifert, M.; Tilgen, W.; Reichrath, J. Present concepts and future outlook: Function of peroxisome proliferator-activated receptors (PPARs) for pathogenesis, progression, and therapy of cancer. J. Cell Physiol. 2007, 212, 1–12. [CrossRef]
41. Kamata, S.; Oyama, T.; Saito, K.; Honda, A.; Yamamoto, Y.; Suda, K.; Ishikawa, R.; Itoh, T.; Watanabe, Y.; Shibata, T.; et al. PPARalpha ligand-binding domain structures with endogenous fatty acids and fibrates. iScience 2020, 23, 101727. [CrossRef]
42. Wahli, W.; Michalik, L. PPARs at the crossroads of lipid signaling and inflammation. Trends Endocrinol. Metab. 2012, 23, 351–363. [CrossRef]
43. Lo Verme, J.; Fu, J.; Astarita, G.; La Rana, G.; Russo, R.; Calignano, A.; Piomelli, D. The nuclear receptor peroxisome proliferator-activated receptor-alpha mediates the anti-inflammatory actions of palmitoylethanolamide. Mol. Pharmacol. 2005, 67, 15–19. [CrossRef]
44. LoVerme, J.; La Rana, G.; Russo, R.; Calignano, A.; Piomelli, D. The search for the palmitoylethanolamide receptor. Life Sci. 2005, 77, 1685–1698. [CrossRef]
45. de Lera, A.R.; Krezel, W.; Ruhl, R. An endogenous mammalian retinoid X receptor ligand, at last! ChemMedChem 2016, 11, 1027–1037. [CrossRef]
46. Ruhl, R.; Krzyzosiak, A.; Niewiadomska-Cimicka, A.; Rochel, N.; Szeles, L.; Vaz, B.; Wietrzych-Schindler, M.; Alvarez, S.; Szklenar, M.; Nagy, L.; et al. 9-cis-13,14-dihydro retinoic acid is an endogenous retinoid acting as RXR ligand in mice. PLoS Genet. 2015, 11, e1005213. [CrossRef] [PubMed]
47. Najib, J. Fenofibrate in the treatment of dyslipidemia: A review of the data as they relate to the new supra bioavailable tablet formulation. Clin. Ther. 2002, 24, 2022–2050. [CrossRef]
48. Blais, J.E.; Tong, G.K.Y.; Pathadka, S.; Mok, M.; Wong, I.C.K.; Chan, E.W. Comparative efficacy and safety of statin and fibrate monotherapy: A systematic review and meta-analysis of head-to-head randomized controlled trials. PLoS ONE 2021, 16, e0246480. [CrossRef] [PubMed]
49. Alsheikh-Ali, A.A.; Kuvin, J.T.; Karas, R.H. Risk of adverse events with fibrates. Am. J. Cardiol. 2004, 94, 935–938. [CrossRef] [PubMed]
50. Kaipainen, A.; Kieran, M.W.; Huang, S.; Butterfield, C.; Bielenberg, D.; Mostoslavsky, G.; Mulligan, R.; Folkman, J.; Panigrahy, D. PPARalpha deficiency in inflammatory cells suppresses tumor growth. PLoS ONE 2007, 2, e260. [CrossRef] [PubMed]
51. Fujimura, Y.; Tachibana, H.; Yamada, K. Peroxisome proliferator-activated receptor ligands negatively regulate the expression of the high-affinity IgE receptor Fc epsilon RI in human basophilic KU812 cells. Biochem. Biophys. Res. Commun. 2002, 297, 193–201. [CrossRef]
52. Woerly, G.; Honda, K.; Loyens, M.; Papin, J.P.; Auwerx, J.; Staels, B.; Capron, M.; Dombrowicz, D. Peroxisome proliferator-activated receptors alpha and gamma down-regulate allergic inflammation and eosinophil activation. J. Exp. Med. 2003, 198, 411–421. [CrossRef] [PubMed]
53. Chinetti, G.; Fruchart, J.C.; Staels, B. Peroxisome proliferator-activated receptors: New targets for the pharmacological modulation of macrophage gene expression and function. Curr. Opin. Lipidol. 2003, 14, 459–468. [CrossRef]
54. Babaev, V.R.; Ishiguro, H.; Ding, L.; Yancey, P.G.; Dove, D.E.; Kovacs, W.J.; Semenkovich, C.F.; Fazio, S.; Linton, M.F. Macrophage expression of peroxisome proliferator-activated receptor-alpha reduces atherosclerosis in low-density lipoprotein receptor-deficient mice. Circulation 2007, 116, 1404–1412. [CrossRef] [PubMed]
55. Wu, L.; Zhang, X.; Zheng, L.; Zhao, H.; Yan, G.; Zhang, Q.; Zhou, Y.; Lei, J.; Zhang, J.; Wang, J.; et al. RIPK3 orchestrates fatty acid metabolism in tumor-associated macrophages and hepatocarcinogenesis. Cancer Immunol. Res. 2020, 8, 710–721. [CrossRef] [PubMed]
56. Brocker, C.N.; Yue, J.; Kim, D.; Qu, A.; Bonzo, J.A.; Gonzalez, F.J. Hepatocyte-specific PPARA expression exclusively promotes agonist-induced cell proliferation without influence from nonparenchymal cells. Am. J. Physiol. Gastrointest. Liver Physiol. 2017, 312, G283–G299. [CrossRef]
57. Dubrac, S.; Stoitzner, P.; Pirkebner, D.; Elentner, A.; Schoonjans, K.; Auwerx, J.; Saeland, S.; Hengster, P.; Fritsch, P.; Romani, N.; et al. Peroxisome proliferator-activated receptor-alpha activation inhibits Langerhans cell function. J. Immunol. 2007, 178, 4362–4372. [CrossRef]
58. Poulsen, R.C.; Moughan, P.J.; Kruger, M.C. Long-chain polyunsaturated fatty acids and the regulation of bone metabolism. Exp. Biol. Med. 2007, 232, 1275–1288. [CrossRef]
59. Warden, A.; Truitt, J.; Merriman, M.; Ponomareva, O.; Jameson, K.; Ferguson, L.B.; Mayfield, R.D.; Harris, R.A. Localization of PPAR isotypes in the adult mouse and human brain. Sci. Rep. 2016, 6, 27618. [CrossRef]
60. Kroetz, D.L.; Yook, P.; Costet, P.; Bianchi, P.; Pineau, T. Peroxisome proliferator-activated receptor alpha controls the hepatic CYP4A induction adaptive response to starvation and diabetes. J. Biol. Chem. 1998, 273, 31581–31589. [CrossRef]
61. Hashimoto, T.; Fujita, T.; Usuda, N.; Cook, W.; Qi, C.; Peters, J.M.; Gonzalez, F.J.; Yeldandi, A.V.; Rao, M.S.; Reddy, J.K. Peroxisomal and mitochondrial fatty acid beta-oxidation in mice nullizygous for both peroxisome proliferator-activated receptor alpha and peroxisomal fatty acyl-CoA oxidase. Genotype correlation with fatty liver phenotype. J. Biol. Chem. 1999, 274, 19228–19236. [CrossRef]
62. Vila-Brau, A.; De Sousa-Coelho, A.L.; Mayordomo, C.; Haro, D.; Marrero, P.F. Human HMGCS2 regulates mitochondrial fatty acid oxidation and FGF21 expression in HepG2 cell line. J. Biol. Chem. 2011, 286, 20423–20430. [CrossRef]
63. Daynes, R.A.; Jones, D.C. Emerging roles of PPARs in inflammation and immunity. Nat. Rev. Immunol. 2002, 2, 748–759. [CrossRef]
64. Ammazzalorso, A.; Bruno, I.; Florio, R.; De Lellis, L.; Laghezza, A.; Cerchia, C.; De Filippis, B.; Fantacuzzi, M.; Giampietro, L.; Maccallini, C.; et al. Sulfonimide and amide derivatives as novel PPARalpha antagonists: Synthesis, antiproliferative activity, and docking studies. ACS Med. Chem. Lett. 2020, 11, 624–632. [CrossRef]
65. Delerive, P.; De Bosscher, K.; Besnard, S.; Vanden Berghe, W.; Peters, J.M.; Gonzalez, F.J.; Fruchart, J.C.; Tedgui, A.; Haegeman, G.; Staels, B. Peroxisome proliferator-activated receptor alpha negatively regulates the vascular inflammatory gene response by negative cross-talk with transcription factors NF-kappaB and AP-1. J. Biol. Chem. 1999, 274, 32048–32054. [CrossRef] [PubMed]
66. Planavila, A.; Iglesias, R.; Giralt, M.; Villarroya, F. Sirt1 acts in association with PPARalpha to protect the heart from hypertrophy, metabolic dysregulation, and inflammation. Cardiovasc. Res. 2011, 90, 276–284. [CrossRef]
67. Rothgiesser, K.M.; Fey, M.; Hottiger, M.O. Acetylation of p65 at lysine 314 is important for late NF-kappaB-dependent gene expression. BMC Genom. 2010, 11, 22. [CrossRef] [PubMed]
68. Zhang, N.; Chu, E.S.; Zhang, J.; Li, X.; Liang, Q.; Chen, J.; Chen, M.; Teoh, N.; Farrell, G.; Sung, J.J.; et al. Peroxisome proliferator-activated receptor alpha inhibits hepatocarcinogenesis by mediating the NF-kappaB signaling pathway. Oncotarget 2014, 5, 8330–8340. [CrossRef] [PubMed]
69. Jacobs, M.D.; Harrison, S.C. Structure of an IkappaBalpha/NF-kappaB complex. Cell 1998, 95, 749–758. [CrossRef]
70. Chen, L.F.; Greene, W.C. Shaping the nuclear action of NF-kappaB. Nat. Rev. Mol. Cell Biol. 2004, 5, 392–401. [CrossRef]
71. Murakami, K.; Bujo, H.; Unoki, H.; Saito, Y. Effect of PPARalpha activation of macrophages on the secretion of inflammatory cytokines in cultured adipocytes. Eur. J. Pharmacol. 2007, 561, 206–213. [CrossRef]
72. Marx, N.; Mackman, N.; Schonbeck, U.; Yilmaz, N.; Hombach, V.; Libby, P.; Plutzky, J. PPARalpha activators inhibit tissue factor expression and activity in human monocytes. Circulation 2001, 103, 213–219. [CrossRef]
73. Neve, B.P.; Corseaux, D.; Chinetti, G.; Zawadzki, C.; Fruchart, J.C.; Duriez, P.; Staels, B.; Jude, B. PPARalpha agonists inhibit tissue factor expression in human monocytes and macrophages. Circulation 2001, 103, 207–212. [CrossRef] 74. Haque, S.J.; Sharma, P. Interleukins and STAT signaling. Vitam. Horm. 2006,
74, 165–206. [CrossRef]
75. Shipley, J.M.; Waxman, D.J. Down-regulation of STAT5b transcriptional activity by ligand-activated peroxisome proliferator-activated receptor (PPAR) alpha and PPARgamma. Mol. Pharmacol. 2003, 64, 355–364. [CrossRef] [PubMed]
76. Shipley, J.M.; Waxman, D.J. Simultaneous, bidirectional inhibitory crosstalk between PPAR and STAT5b. Toxicol. Appl. Pharmacol. 2004, 199, 275–284. [CrossRef] [PubMed]
77. Zhou, Y.C.; Waxman, D.J. STAT5b down-regulates peroxisome proliferator-activated receptor alpha transcription by inhibition of ligand-independent activation function region-1 trans-activation domain. J. Biol. Chem. 1999, 274, 29874–29882. [CrossRef] [PubMed]
78. Lin, J.X.; Leonard, W.J. The role of Stat5a and Stat5b in signaling by IL-2 family cytokines. Oncogene 2000, 19, 2566–2576. [CrossRef] [PubMed]
79. Bendickova, K.; Fric, J. Roles of IL-2 in bridging adaptive and innate immunity, and as a tool for cellular immunotherapy. J. Leukoc. Biol. 2020, 108, 427–437. [CrossRef] 80. Devchand, P.R.; Keller, H.; Peters, J.M.; Vazquez, M.; Gonzalez, F.J.; Wahli, W. The PPARalpha-leukotriene B4 pathway to inflammation control. Nature 1996, 384, 39–43. [CrossRef]
81. Dahlen, S.E.; Bjork, J.; Hedqvist, P.; Arfors, K.E.; Hammarstrom, S.; Lindgren, J.A.; Samuelsson, B. Leukotrienes promote plasma leakage and leukocyte adhesion in postcapillary venules: In vivo effects with relevance to the acute inflammatory response. Proc. Natl. Acad. Sci. USA 1981, 78, 3887–3891. [CrossRef]
82. Lindbom, L.; Hedqvist, P.; Dahlen, S.E.; Lindgren, J.A.; Arfors, K.E. Leukotriene B4 induces extravasation and migration of polymorphonuclear leukocytes in vivo. Acta Physiol. Scand. 1982, 116, 105–108. [CrossRef] [PubMed]
83. Marleau, S.; Fruteau de Laclos, B.; Sanchez, A.B.; Poubelle, P.E.; Borgeat, P. Role of 5-lipoxygenase products in the local accumulation of neutrophils in dermal inflammation in the rabbit. J. Immunol. 1999, 163, 3449–3458.
84. Yu, K.; Bayona, W.; Kallen, C.B.; Harding, H.P.; Ravera, C.P.; McMahon, G.; Brown, M.; Lazar, M.A. Differential activation of peroxisome proliferator-activated receptors by eicosanoids. J. Biol. Chem. 1995, 270, 23975–23983. [CrossRef] [PubMed]
85. Christofides, A.; Konstantinidou, E.; Jani, C.; Boussiotis, V.A. The role of peroxisome proliferator-activated receptors (PPAR) in immune responses. Metabolism 2021, 114, 154338. [CrossRef]
86. Wolf, A.J.; Reyes, C.N.; Liang, W.; Becker, C.; Shimada, K.; Wheeler, M.L.; Cho, H.C.; Popescu, N.I.; Coggeshall, K.M.; Arditi, M.; et al. Hexokinase Is an innate immune receptor for the detection of bacterial peptidoglycan. Cell 2016, 166, 624–636. [CrossRef]
87. Frosali, S.; Pagliari, D.; Gambassi, G.; Landolfi, R.; Pandolfi, F.; Cianci, R. How the intricate interaction among toll-like receptors, microbiota, and intestinal immunity can influence gastrointestinal pathology. J. Immunol. Res. 2015, 2015, 489821. [CrossRef]
88. Wang, Y.; Yin, Y.; Chen, X.; Zhao, Y.; Wu, Y.; Li, Y.; Wang, X.; Chen, H.; Xiang, C. Induction of intestinal Th17 cells by flagellins from segmented filamentous bacteria. Front. Immunol. 2019, 10, 2750. [CrossRef] [PubMed]
89. Park, J.H.; Jeong, S.Y.; Choi, A.J.; Kim, S.J. Lipopolysaccharide directly stimulates Th17 differentiation in vitro modulating phosphorylation of RelB and NF-kappaB1. Immunol. Lett. 2015, 165, 10–19. [CrossRef]
90. Silveira, L.S.; Pimentel, G.D.; Souza, C.O.; Biondo, L.A.; Teixeira, A.A.S.; Lima, E.A.; Batatinha, H.A.P.; Rosa Neto, J.C.; Lira, F.S. Effect of an acute moderate-exercise session on the metabolic and inflammatory profile of PPAR-alpha knockout mice. Cell Biochem. Funct. 2017, 35, 510–517. [CrossRef]
91. Becker, J.; Delayre-Orthez, C.; Frossard, N.; Pons, F. Regulation of peroxisome proliferator-activated receptor-alpha expression during lung inflammation. Pulm. Pharmacol. Ther. 2008, 21, 324–330. [CrossRef]
92. Chistyakov, D.V.; Aleshin, S.E.; Astakhova, A.A.; Sergeeva, M.G.; Reiser, G. Regulation of peroxisome proliferator-activated receptors (PPAR) alpha and -gamma of rat brain astrocytes in the course of activation by toll-like receptor agonists. J. Neurochem. 2015, 134, 113–124. [CrossRef]
93. Dana, N.; Javanmard, H.S.; Vaseghi, G. The effect of fenofibrate, a PPARalpha activator on toll-like receptor-4 signal transduction in melanoma both in vitro and in vivo. Clin. Transl. Oncol. 2020, 22, 486–494. [CrossRef]
94. Shen, W.; Gao, Y.; Lu, B.; Zhang, Q.; Hu, Y.; Chen, Y. Negatively regulating TLR4/NF-kappaB signaling via PPARalpha in endotoxin-induced uveitis. Biochim. Biophys. Acta 2014, 1842, 1109–1120. [CrossRef] [PubMed]
95. Lamkanfi, M.; Dixit, V.M. Mechanisms and functions of inflammasomes. Cell 2014, 157, 1013–1022. [CrossRef] [PubMed]
96. Zheng, D.; Liwinski, T.; Elinav, E. Inflammasome activation and regulation: Toward a better understanding of complex mechanisms. Cell Discov. 2020, 6, 36. [CrossRef] [PubMed]
97. Seok, J.K.; Kang, H.C.; Cho, Y.Y.; Lee, H.S.; Lee, J.Y. Regulation of the NLRP3 inflammasome by post-translational modifications and small molecules. Front. Immunol. 2020, 11, 618231. [CrossRef]
98. Gugliandolo, E.; Fusco, R.; Ginestra, G.; D’Amico, R.; Bisignano, C.; Mandalari, G.; Cuzzocrea, S.; Di Paola, R. Involvement of TLR4 and PPAR-alpha receptors in host response and NLRP3 Inflammasome activation, against pulmonary infection with Pseudomonas aeruginosa. Shock 2019, 51, 221–227. [CrossRef] [PubMed]
99. Brocker, C.N.; Kim, D.; Melia, T.; Karri, K.; Velenosi, T.J.; Takahashi, S.; Aibara, D.; Bonzo, J.A.; Levi, M.; Waxman, D.J.; et al. Long non-coding RNA Gm15441 attenuates hepatic inflammasome activation in response to PPARA agonism and fasting. Nat. Commun. 2020, 11, 5847. [CrossRef]
100. Hu, J.; Zhu, Z.; Ying, H.; Yao, J.; Ma, H.; Li, L.; Zhao, Y. Oleoylethanolamide protects against acute liver injury by regulating Nrf-2/HO-1 and NLRP3 pathways in mice. Front. Pharmacol. 2020, 11, 605065. [CrossRef]
101. Lama, A.; Provensi, G.; Amoriello, R.; Pirozzi, C.; Rani, B.; Mollica, M.P.; Raso, G.M.; Ballerini, C.; Meli, R.; Passani, M.B. The anti-inflammatory and immune-modulatory effects of OEA limit DSS-induced colitis in mice. Biomed. Pharmacother. 2020, 129, 110368. [CrossRef]
102. Alderton, W.K.; Cooper, C.E.; Knowles, R.G. Nitric oxide synthases: Structure, function, and inhibition. Biochem. J. 2001, 357, 593–615. [CrossRef]
103. Kleinert, H.; Schwarz, P.M.; Forstermann, U. Regulation of the expression of inducible nitric oxide synthase. Biol. Chem. 2003, 384, 1343–1364. [CrossRef] [PubMed] 104. Forstermann, U.; Boissel, J.P.; Kleinert, H. Expressional control of the ‘constitutive’ isoforms of nitric oxide synthase (NOS I and NOS III). FASEB J. 1998, 12, 773–790. [CrossRef] [PubMed]
105. Pou, S.; Keaton, L.; Surichamorn, W.; Rosen, G.M. Mechanism of superoxide generation by neuronal nitric-oxide synthase. J. Biol. Chem. 1999, 274, 9573–9580. [CrossRef] [PubMed]
106. Paukkeri, E.L.; Leppanen, T.; Sareila, O.; Vuolteenaho, K.; Kankaanranta, H.; Moilanen, E. PPARalpha agonists inhibit nitric oxide production by enhancing iNOS degradation in LPS-treated macrophages. Br. J. Pharmacol. 2007, 152, 1081–1091. [CrossRef] [PubMed]
107. Van Ginderachter, J.A.; Movahedi, K.; Van den Bossche, J.; De Baetselier, P. Macrophages, PPARs, and cancer. PPAR Res. 2008, 2008, 169414. [CrossRef]
108. Maccallini, C.; Mollica, A.; Amoroso, R. The positive regulation of eNOS signaling by PPAR agonists in cardiovascular diseases. Am. J. Cardiovasc. Drugs 2017, 17, 273–281. [CrossRef]
109. Tanaka, S.; Hosogi, S.; Sawabe, Y.; Shimamoto, C.; Matsumura, H.; Inui, T.; Marunaka, Y.; Nakahari, T. PPARalpha induced NOS1 phosphorylation via PI3K/Akt in guinea pig antral mucous cells: NO-enhancement in Ca(2+)-regulated exocytosis. Biomed. Res. 2016, 37, 167–178. [CrossRef]
110. Tanaka, S.; Sugiyama, N.; Takahashi, Y.; Mantoku, D.; Sawabe, Y.; Kuwabara, H.; Nakano, T.; Shimamoto, C.; Matsumura, H.; Marunaka, Y.; et al. PPARalpha autocrine regulation of Ca(2)(+)-regulated exocytosis in guinea pig antral mucous cells: NO and cGMP accumulation. Am. J. Physiol. Gastrointest. Liver Physiol. 2014, 307, G1169–G1179. [CrossRef]
111. Bune, A.J.; Shergill, J.K.; Cammack, R.; Cook, H.T. L-arginine depletion by arginase reduces nitric oxide production in endotoxic shock: An electron paramagnetic resonance study. FEBS Lett. 1995, 366, 127–130. [CrossRef]
112. Mills, C.D.; Kincaid, K.; Alt, J.M.; Heilman, M.J.; Hill, A.M. M-1/M-2 macrophages and the Th1/Th2 paradigm. J. Immunol. 2000, 164, 6166–6173. [CrossRef] [PubMed] 113. Grabacka, M.; Placha, W.; Plonka, P.M.; Pajak, S.; Urbanska, K.; Laidler, P.; Slominski, A. Inhibition of melanoma metastases by fenofibrate. Arch. Dermatol. Res. 2004, 296, 54–58. [CrossRef]
114. Gallorini, M.; Rapino, M.; Schweikl, H.; Cataldi, A.; Amoroso, R.; Maccallini, C. Selective inhibitors of the inducible nitric oxide synthase as modulators of cell responses in LPS-stimulated human monocytes. Molecules 2021, 26, 4419. [CrossRef]
115. Teissier, E.; Nohara, A.; Chinetti, G.; Paumelle, R.; Cariou, B.; Fruchart, J.C.; Brandes, R.P.; Shah, A.; Staels, B. Peroxisome proliferator-activated receptor alpha induces NADPH oxidase activity in macrophages, leading to the generation of LDL with PPAR-alpha activation properties. Circ. Res. 2004, 95, 1174–1182. [CrossRef]
116. Deininger, P. Alu elements: Know the SINEs. Genome Biol. 2011, 12, 236. [CrossRef] [PubMed]
117. Reynolds, W.F.; Kumar, A.P.; Piedrafita, F.J. The human myeloperoxidase gene is regulated by LXR and PPARalpha ligands. Biochem. Biophys. Res. Commun. 2006, 349, 846–854. [CrossRef]
118. Penas, F.; Mirkin, G.A.; Vera, M.; Cevey, A.; Gonzalez, C.D.; Gomez, M.I.; Sales, M.E.; Goren, N.B. Treatment in vitro with PPARalpha and PPARgamma ligands drive M1-to-M2 polarization of macrophages from T. cruzi-infected mice. Biochim. Biophys. Acta 2015, 1852, 893–904. [CrossRef] [PubMed]
119. Redlich, S.; Ribes, S.; Schutze, S.; Nau, R. Palmitoylethanolamide stimulates phagocytosis of Escherichia coli K1 by macrophages and increases the resistance of mice against infections. J. Neuroinflamm. 2014, 11, 108. [CrossRef]
120. Kim, Y.S.; Lee, H.M.; Kim, J.K.; Yang, C.S.; Kim, T.S.; Jung, M.; Jin, H.S.; Kim, S.; Jang, J.; Oh, G.T.; et al. PPAR-alpha activation mediates innate host defense through the induction of TFEB and lipid catabolism. J. Immunol. 2017, 198, 3283–3295. [CrossRef]
121. Peyron, P.; Vaubourgeix, J.; Poquet, Y.; Levillain, F.; Botanch, C.; Bardou, F.; Daffe, M.; Emile, J.F.; Marchou, B.; Cardona, P.J.; et al. Foamy macrophages from tuberculous patients’ granulomas constitute a nutrient-rich reservoir for M. tuberculosis persistence. PLoS Pathog. 2008, 4, e1000204. [CrossRef]
122. Tam, V.C.; Suen, R.; Treuting, P.M.; Armando, A.; Lucarelli, R.; Gorrochotegui-Escalante, N.; Diercks, A.H.; Quehenberger, O.; Dennis, E.A.; Aderem, A.; et al. PPARalpha exacerbates necroptosis, leading to increased mortality in post-influenza bacterial superinfection. Proc. Natl. Acad. Sci. USA 2020, 117, 15789–15798. [CrossRef] [PubMed]
123. Ng, V.Y.; Huang, Y.; Reddy, L.M.; Falck, J.R.; Lin, E.T.; Kroetz, D.L. Cytochrome P450 eicosanoids are activators of peroxisome proliferator-activated receptor alpha. Drug Metab. Dispos. 2007, 35, 1126–1134. [CrossRef]
124. Yang, M.; Wang, Y.; Chen, J.; Wang, Q.; Wei, S.; Wang, S.; Qin, Q. Functional analysis of Epinephelus coioides peroxisome proliferative-activated receptor alpha (PPARalpha): Involvement in response to viral infection. Fish. Shellfish Immunol. 2020, 102, 257–266. [CrossRef]
125. Ehrlich, A.; Uhl, S.; Ioannidis, K.; Hofree, M.; tenOever, B.R.; Nahmias, Y. The SARS-CoV-2 Transcriptional Metabolic Signature in Lung Epithelium. Available online: https://ssrn.com/abstract=3650499 (accessed on 28 September 2021).
126. Heffernan, K.S.; Ranadive, S.M.; Jae, S.Y. Exercise as medicine for COVID-19: On PPAR with emerging pharmacotherapy. Med. Hypotheses 2020, 143, 110197. [CrossRef]
127. Piomelli, D.; Hohmann, A.G.; Seybold, V.; Hammock, B.D. A lipid gate for the peripheral control of pain. J. Neurosci. 2014, 34, 15184–15191. [CrossRef]
128. Chiurchiu, V.; Battistini, L.; Maccarrone, M. Endocannabinoid signaling in innate and adaptive immunity. Immunology 2015, 144, 352–364. [CrossRef]
129. Guindon, J.; Hohmann, A.G. The endocannabinoid system and pain. CNS Neurol. Disord. Drug Targets 2009, 8, 403–421. [CrossRef] [PubMed]
130. Bisogno, T.; Ventriglia, M.; Milone, A.; Mosca, M.; Cimino, G.; Di Marzo, V. Occurrence and metabolism of anandamide and related acyl-ethanolamides in ovaries of the sea urchin Paracentrotus lividus. Biochim. Biophys. Acta 1997, 1345, 338–348. [CrossRef]
131. Sepe, N.; De Petrocellis, L.; Montanaro, F.; Cimino, G.; Di Marzo, V. Bioactive long chain N-acyl ethanolamines in five species of edible bivalve mollusks. Possible implications for mollusk physiology and seafood industry. Biochim. Biophys. Acta 1998, 1389, 101–111. [CrossRef]
132. Matias, I.; Bisogno, T.; Melck, D.; Vandenbulcke, F.; Verger-Bocquet, M.; De Petrocellis, L.; Sergheraert, C.; Breton, C.; Di Marzo, V.; Salzet, M. Evidence for an endocannabinoid system in the central nervous system of the leech Hirudo medicinalis. Brain Res. Mol. Brain Res. 2001, 87, 145–159. [CrossRef]
133. Fu, J.; Gaetani, S.; Oveisi, F.; Lo Verme, J.; Serrano, A.; Rodriguez De Fonseca, F.; Rosengarth, A.; Luecke, H.; Di Giacomo, B.; Tarzia, G.; et al. Oleylethanolamide regulates feeding and body weight through the activation of the nuclear receptor PPAR-alpha. Nature 2003, 425, 90–93. [CrossRef]
134. Bradshaw, H.B.; Walker, J.M. The expanding field of cannabimimetic and related lipid mediators. Br. J. Pharmacol. 2005, 144, 459–465. [CrossRef]
135. Oddi, S.; Fezza, F.; Pasquariello, N.; De Simone, C.; Rapino, C.; Dainese, E.; Finazzi-Agro, A.; Maccarrone, M. Evidence for the intracellular accumulation of anandamide in adiposomes. Cell Mol. Life Sci. 2008, 65, 840–850. [CrossRef]
136. Kaczocha, M.; Glaser, S.T.; Chae, J.; Brown, D.A.; Deutsch, D.G. Lipid droplets are novel sites of N-acylethanolamine inactivation by fatty acid amide hydrolase-2. J. Biol. Chem. 2010, 285, 2796–2806. [CrossRef] [PubMed]
137. Bisogno, T.; Maurelli, S.; Melck, D.; De Petrocellis, L.; Di Marzo, V. Biosynthesis, uptake, and degradation of anandamide and palmitoylethanolamide in leukocytes. J. Biol. Chem. 1997, 272, 3315–3323. [CrossRef] [PubMed]
138. Tsuboi, K.; Ikematsu, N.; Uyama, T.; Deutsch, D.G.; Tokumura, A.; Ueda, N. Biosynthetic pathways of bioactive Nacylethanolamines in the brain. CNS Neurol. Disord. Drug Targets 2013, 12, 7–16. [CrossRef]
139. Fegley, D.; Gaetani, S.; Duranti, A.; Tontini, A.; Mor, M.; Tarzia, G.; Piomelli, D. Characterization of the fatty acid amide hydrolase inhibitor cyclohexyl carbamic acid 30 -carbamoyl-biphenyl-3-yl ester (URB597): Effects on anandamide and oleoylethanolamide deactivation. J. Pharmacol. Exp. Ther. 2005, 313, 352–358. [CrossRef]
140. Ogura, Y.; Parsons, W.H.; Kamat, S.S.; Cravatt, B.F. A calcium-dependent acyltransferase that produces N-acyl phosphatidylethanolamines. Nat. Chem. Biol. 2016, 12, 669–671. [CrossRef]
141. Ueda, N.; Tsuboi, K.; Uyama, T. Metabolism of endocannabinoids and related N-acyl ethanolamines: Canonical and alternative pathways. FEBS J. 2013, 280, 1874–1894. [CrossRef] [PubMed]
142. LoVerme, J.; Russo, R.; La Rana, G.; Fu, J.; Farthing, J.; Mattace-Raso, G.; Meli, R.; Hohmann, A.; Calignano, A.; Piomelli, D. Rapid broad-spectrum analgesia through activation of peroxisome proliferator-activated receptor-alpha. J. Pharmacol. Exp. Ther. 2006, 319, 1051–1061. [CrossRef]
143. Suardiaz, M.; Estivill-Torrus, G.; Goicoechea, C.; Bilbao, A.; de Fonseca, R.F. Analgesic properties of oleoyl ethanolamide (OEA) in visceral and inflammatory pain. Pain 2007, 133, 99–110. [CrossRef] [PubMed]
144. Taylor, B.K.; Dadia, N.; Yang, C.B.; Krishnan, S.; Badr, M. Peroxisome proliferator-activated receptor agonists inhibit inflammatory edema and hyperalgesia. Inflammation 2002, 26, 121–127. [CrossRef]
145. Russo, R.; LoVerme, J.; La Rana, G.; D’Agostino, G.; Sasso, O.; Calignano, A.; Piomelli, D. Synergistic antinociception by the cannabinoid receptor agonist anandamide and the PPAR-alpha receptor agonist GW7647. Eur. J. Pharmacol. 2007, 566, 117–119. [CrossRef]
146. D’Agostino, G.; La Rana, G.; Russo, R.; Sasso, O.; Iacono, A.; Esposito, E.; Raso, G.M.; Cuzzocrea, S.; Lo Verme, J.; Piomelli, D.; et al. Acute intracerebroventricular administration of palmitoylethanolamide, an endogenous peroxisome proliferator-activated receptor-alpha agonist, modulates carrageenan-induced paw edema in mice. J. Pharmacol. Exp. Ther. 2007, 322, 1137–1143. [CrossRef] [PubMed]
147. Russo, R.; Loverme, J.; La Rana, G.; Compton, T.R.; Parrott, J.; Duranti, A.; Tontini, A.; Mor, M.; Tarzia, G.; Calignano, A.; et al. The fatty acid amide hydrolase inhibitor URB597 (cyclohexyl carbamic acid 30 -carbamoylbiphenyl-3-yl ester) reduces neuropathic pain after oral administration in mice. J. Pharmacol. Exp. Ther. 2007, 322, 236–242. [CrossRef]
148. Ben-Shabat, S.; Fride, E.; Sheskin, T.; Tamiri, T.; Rhee, M.H.; Vogel, Z.; Bisogno, T.; De Petrocellis, L.; Di Marzo, V.; Mechoulam, R. An entourage effect: Inactive endogenous fatty acid glycerol esters enhance 2-arachidonoyl-glycerol cannabinoid activity. Eur. J. Pharmacol. 1998, 353, 23–31. [CrossRef]
149. Ho, W.S.; Barrett, D.A.; Randall, M.D. ‘Entourage’ effects of N-palmitoylethanolamide and N-oleoyl ethanolamide on vasorelaxation to anandamide occur through TRPV1 receptors. Br. J. Pharmacol. 2008, 155, 837–846. [CrossRef]
150. Steardo, L., Jr.; Bronzuoli, M.R.; Iacomino, A.; Esposito, G.; Steardo, L.; Scuderi, C. Does neuroinflammation turn on the flame in Alzheimer’s disease? Focus on astrocytes. Front. Neurosci. 2015, 9, 259. [CrossRef]
151. Skaper, S.D.; Facci, L.; Giusti, P. Glia and mast cells as targets for palmitoylethanolamide, an anti-inflammatory and neuroprotective lipid mediator. Mol. Neurobiol. 2013, 48, 340–352. [CrossRef] [PubMed]
152. Amor, S.; Puentes, F.; Baker, D.; van der Valk, P. Inflammation in neurodegenerative diseases. Immunology 2010, 129, 154–169. [CrossRef] [PubMed]
153. Colombo, E.; Farina, C. Astrocytes: Key regulators of neuroinflammation. Trends Immunol. 2016, 37, 608–620. [CrossRef] [PubMed]
154. Aloe, L.; Leon, A.; Levi-Montalcini, R. A proposed autacoid mechanism controlling mastocyte behavior. Agents Actions 1993, 39, C145–C147. [CrossRef] 155. Benito, C.; Tolon, R.M.; Castillo, A.I.; Ruiz-Valdepenas, L.; Martinez-Orgado, J.A.; Fernandez-Sanchez, F.J.; Vazquez, C.; Cravatt, B.F.; Romero, J. beta-Amyloid exacerbates inflammation in astrocytes lacking fatty acid amide hydrolase through a mechanism involving PPAR-alpha, PPAR-gamma, and TRPV1, but not CB(1) or CB(2) receptors. Br. J. Pharmacol. 2012, 166, 1474–1489. [CrossRef]
156. Hohmann, U.; Pelzer, M.; Kleine, J.; Hohmann, T.; Ghadban, C.; Dehghani, F. Opposite effects of neuroprotective cannabinoids, palmitoylethanolamide, and 2-arachidonoylglycerol on function and morphology of microglia. Front. Neurosci. 2019, 13, 1180. [CrossRef] [PubMed]
157. Bronzuoli, M.R.; Facchinetti, R.; Steardo, L., Jr.; Romano, A.; Stecca, C.; Passarella, S.; Steardo, L.; Cassano, T.; Scuderi, C. Palmitoylethanolamide dampens reactive astrogliosis and improves neuronal trophic support in a triple transgenic model of Alzheimer’s disease: In vitro and in vivo evidence. Oxid. Med. Cell. Longev. 2018, 2018, 4720532. [CrossRef]
158. Scuderi, C.; Stecca, C.; Valenza, M.; Ratano, P.; Bronzuoli, M.R.; Bartoli, S.; Steardo, L.; Pompili, E.; Fumagalli, L.; Campolongo, P.; et al. Palmitoylethanolamide controls reactive gliosis and exerts neuroprotective functions in a rat model of Alzheimer’s disease. Cell Death Dis. 2014, 5, e1419. [CrossRef] [PubMed]
159. Scuderi, C.; Valenza, M.; Stecca, C.; Esposito, G.; Carratu, M.R.; Steardo, L. Palmitoylethanolamide exerts neuroprotective effects in mixed neuroglial cultures and organotypic hippocampal slices via peroxisome proliferator-activated receptor-alpha. J. Neuroinflamm. 2012, 9, 49. [CrossRef]
160. Guida, F.; Luongo, L.; Boccella, S.; Giordano, M.E.; Romano, R.; Bellini, G.; Manzo, I.; Furiano, A.; Rizzo, A.; Imperatore, R.; et al. Palmitoylethanolamide induces microglia changes associated with increased migration and phagocytic activity: Involvement of the CB2 receptor. Sci. Rep. 2017, 7, 375. [CrossRef]
161. Rinne, P.; Guillamat-Prats, R.; Rami, M.; Bindila, L.; Ring, L.; Lyytikainen, L.P.; Raitoharju, E.; Oksala, N.; Lehtimaki, T.; Weber, C.; et al. Palmitoylethanolamide promotes a pro-resolving macrophage phenotype and attenuates atherosclerotic plaque formation. Arterioscler. Thromb. Vasc. Biol. 2018, 38, 2562–2575. [CrossRef]
162. Luo, D.; Zhang, Y.; Yuan, X.; Pan, Y.; Yang, L.; Zhao, Y.; Zhuo, R.; Chen, C.; Peng, L.; Li, W.; et al. Oleoylethanolamide inhibits glial activation via modulating PPARalpha and promotes motor function recovery after brain ischemia. Pharmacol. Res. 2019, 141, 530–540. [CrossRef]
163. Xu, X.; Guo, H.; Jing, Z.; Yang, L.; Chen, C.; Peng, L.; Wang, X.; Yan, L.; Ye, R.; Jin, X.; et al. N-oleoyl ethanolamine reduces inflammatory cytokines and adhesion molecules in TNF-alpha-induced human umbilical vein endothelial cells by activating CB2 and PPAR-alpha. J. Cardiovasc. Pharmacol. 2016, 68, 280–291. [CrossRef]
164. Holubiec, M.I.; Romero, J.I.; Suarez, J.; Portavella, M.; Fernandez-Espejo, E.; Blanco, E.; Galeano, P.; de Fonseca, F.R. Palmitoylethanolamide prevents neuroinflammation, reduces astrogliosis and preserves recognition and spatial memory following induction of neonatal anoxia-ischemia. Psychopharmacology 2018, 235, 2929–2945. [CrossRef] [PubMed]
165. Yang, L.C.; Guo, H.; Zhou, H.; Suo, D.Q.; Li, W.J.; Zhou, Y.; Zhao, Y.; Yang, W.S.; Jin, X. Chronic oleoyl ethanolamide treatment improves spatial cognitive deficits through enhancing hippocampal neurogenesis after transient focal cerebral ischemia. Biochem. Pharmacol. 2015, 94, 270–281. [CrossRef]
166. Flannery, L.E.; Kerr, D.M.; Hughes, E.M.; Kelly, C.; Costello, J.; Thornton, A.M.; Humphrey, R.M.; Finn, D.P.; Roche, M. Nacylethanolamine regulation of TLR3-induced hyperthermia and neuroinflammatory gene expression: A role for PPARalpha. J. Neuroimmunol. 2021, 358, 577654. [CrossRef] [PubMed]
167. Taquet, M.; Geddes, J.R.; Husain, M.; Luciano, S.; Harrison, P.J. 6-month neurological and psychiatric outcomes in 236 379 survivors of COVID-19: A retrospective cohort study using electronic health records. Lancet Psychiatry 2021, 8, 416–427. [CrossRef]
168. Vaia, M.; Petrosino, S.; De Filippis, D.; Negro, L.; Guarino, A.; Carnuccio, R.; Di Marzo, V.; Iuvone, T. Palmitoylethanolamide reduces inflammation and itch in a mouse model of contact allergic dermatitis. Eur. J. Pharmacol. 2016, 791, 669–674. [CrossRef] [PubMed]
169. Skaper, S.D.; Facci, L.; Romanello, S.; Leon, A. Mast cell activation causes delayed neurodegeneration in mixed hippocampal cultures via the nitric oxide pathway. J. Neurochem. 1996, 66, 1157–1166. [CrossRef]
170. Facci, L.; Dal Toso, R.; Romanello, S.; Buriani, A.; Skaper, S.D.; Leon, A. Mast cells express a peripheral cannabinoid receptor with differential sensitivity to anandamide and palmitoylethanolamide. Proc. Natl. Acad. Sci. USA 1995, 92, 3376–3380. [CrossRef]
171. Di Paola, R.; Impellizzeri, D.; Torre, A.; Mazzon, E.; Cappellani, A.; Faggio, C.; Esposito, E.; Trischitta, F.; Cuzzocrea, S. Effects of palmitoylethanolamide on intestinal injury and inflammation caused by ischemia-reperfusion in mice. J. Leukoc. Biol. 2012, 91, 911–920. [CrossRef] [PubMed]
172. Borrelli, F.; Romano, B.; Petrosino, S.; Pagano, E.; Capasso, R.; Coppola, D.; Battista, G.; Orlando, P.; Di Marzo, V.; Izzo, A.A. Palmitoylethanolamide, a naturally occurring lipid, is an orally effective intestinal anti-inflammatory agent. Br. J. Pharmacol. 2015, 172, 142–158. [CrossRef]
173. Misto, A.; Provensi, G.; Vozella, V.; Passani, M.B.; Piomelli, D. Mast cell-derived histamine regulates liver ketogenesis via oleoyl ethanolamide signaling. Cell Metab. 2019, 29, 91–102e5. [CrossRef] [PubMed]
174. Fu, J.; Kim, J.; Oveisi, F.; Astarita, G.; Piomelli, D. Targeted enhancement of oleoylethanolamide production in proximal small intestine induces across-meal satiety in rats. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2008, 295, R45–R50. [CrossRef]
175. Golderer, G.; Werner, E.R.; Leitner, S.; Grobner, P.; Werner-Felmayer, G. Nitric oxide synthase is induced in sporulation of Physarum polycephalum. Genes Dev. 2001, 15, 1299–1309. [CrossRef]
176. Hall, D.A.; Vander Kooi, C.W.; Stasik, C.N.; Stevens, S.Y.; Zuiderweg, E.R.; Matthews, R.G. Mapping the interactions between flavodoxin and its physiological partner's flavodoxin reductase and cobalamin-dependent methionine synthase. Proc. Natl. Acad. Sci. USA 2001, 98, 9521–9526. [CrossRef] [PubMed]
177. Radomski, M.W.; Palmer, R.M.; Moncada, S. Modulation of platelet aggregation by an L-arginine-nitric oxide pathway. Trends Pharmacol. Sci. 1991, 12, 87–88. [CrossRef]
178. Escriva, H.; Safi, R.; Hanni, C.; Langlois, M.C.; Saumitou-Laprade, P.; Stehelin, D.; Capron, A.; Pierce, R.; Laudet, V. Ligand binding was acquired during evolution of nuclear receptors. Proc. Natl. Acad. Sci. USA 1997, 94, 6803–6808. [CrossRef]
179. Zhou, T.; Yan, X.; Wang, G.; Liu, H.; Gan, X.; Zhang, T.; Wang, J.; Li, L. Evolutionary pattern and regulation analysis to support why diversity functions existed within PPAR gene family members. Biomed. Res. Int. 2015, 2015, 613910. [CrossRef]
180. Crawford, N.M. Mechanisms for nitric oxide synthesis in plants. J. Exp. Bot. 2006, 57, 471–478. [CrossRef] 181. Dohrmann, M.; Worheide, G. Dating early animal evolution using phylogenomic data. Sci. Rep. 2017, 7, 3599. [CrossRef]
For more information:1950477648nn@gmail.com
