The Octadecanoids: Synthesis and Bioactivity of 18-Carbon Oxygenated Fatty Acids in Mammals, Bacteria, and Fungi

Johanna Revol-Cavalier; Alessandro Quaranta; John W Newman; Alan R Brash; Mats Hamberg; Craig E Wheelock

doi:10.1021/acs.chemrev.3c00520

. 2024 Dec 16;125(1):1–90. doi: 10.1021/acs.chemrev.3c00520

The Octadecanoids: Synthesis and Bioactivity of 18-Carbon Oxygenated Fatty Acids in Mammals, Bacteria, and Fungi

Johanna Revol-Cavalier ^†,^‡, Alessandro Quaranta ^†, John W Newman ^§,^#,^∇, Alan R Brash ^∥, Mats Hamberg ^†,^‡, Craig E Wheelock ^†,^⊥,^*

PMCID: PMC11719350 PMID: 39680864

Abstract

graphic file with name cr3c00520_0098.jpg

The octadecanoids are a broad class of lipids consisting of the oxygenated products of 18-carbon fatty acids. Originally referring to production of the phytohormone jasmonic acid, the octadecanoid pathway has been expanded to include products of all 18-carbon fatty acids. Octadecanoids are formed biosynthetically in mammals via cyclooxygenase (COX), lipoxygenase (LOX), and cytochrome P450 (CYP) activity, as well as nonenzymatically by photo- and autoxidation mechanisms. While octadecanoids are well-known mediators in plants, their role in the regulation of mammalian biological processes has been generally neglected. However, there have been significant advancements in recognizing the importance of these compounds in mammals and their involvement in the mediation of inflammation, nociception, and cell proliferation, as well as in immuno- and tissue modulation, coagulation processes, hormone regulation, and skin barrier formation. More recently, the gut microbiome has been shown to be a significant source of octadecanoid biosynthesis, providing additional biosynthetic routes including hydratase activity (e.g., CLA-HY, FA-HY1, FA-HY2). In this review, we summarize the current field of octadecanoids, propose standardized nomenclature, provide details of octadecanoid preparation and measurement, summarize the phase-I metabolic pathway of octadecanoid formation in mammals, bacteria, and fungi, and describe their biological activity in relation to mammalian pathophysiology as well as their potential use as biomarkers of health and disease.

1. Introduction

1.1. Fatty Acid Biosynthesis

Fatty acids were described by Michel Eugène Chevreul in 1813.¹ They have since been demonstrated to be important sources of energy and membrane constituents as well as act as signaling molecules that are involved in multiple biological activities including intracellular signaling, regulation of transcription, protein modification and other cellular processes.²⁻⁴ The early work of George and Mildred Burr established that the C18 polyunsaturated long-chain fatty acids (PUFAs) linoleic acid (LA, 18:2 ω6) and α-linolenic acid (ALA, 18:3 ω3) are dietary essential fatty acids (EFAs)⁵ because humans and most mammals lack the desaturase enzymes that can introduce an ω6 (n-6) or ω3 (n-3) double bond beyond carbons 9 and 10.⁶ Ralph Holman then formally established the ω6 and ω3 families of PUFAs in 1963.⁷ PUFA biochemistry has now been extensively mapped and we have a solid understanding of the enzymatic processes that form the full complement of these compounds in vivo (Scheme 1). Fatty acid biosynthesis involves a series of CH₂ extensions and desaturations on the α-side of the fatty acid chain to convert EFAs to longer chain PUFAs.⁸ Both LA and ALA are converted by desaturases (fatty acid desaturases, FADS) and elongases (elongation of very long chain fatty acids, ELOVL) into downstream PUFAs. γ-linolenic acid (GLA, 18:3 ω6) and stearidonic acid (SDA, 18:4 ω3) are formed from LA and ALA, respectively, via the action of FADS2, and further converted into C20-PUFAs (dihomo-γ-linolenic acid (DGLA) and eicosatetraenoic acid (ETA), respectively) by ELOVL5. The enzyme FADS1 then desaturates these species to form arachidonic acid (AA) and eicosapentaenoic acid (EPA), which can then be converted into C22-PUFAs (adrenic acid (AdA) and docosapentaenoic acid (DPAn-3)) by ELOVL2. Finally, FADS2 activity forms DPAn-6 and docosahexaenoic acid (DHA), respectively.⁹ As our knowledge of the fatty acid biosynthesis pathways has increased, so has our understanding of their importance in biochemistry, physiology and health, with dysregulation in fatty acid biosynthesis associated with multiple pathologies.¹⁰

Scheme 1 — FADS = fatty acid desaturase. ELOVL = elongase. PUFA nomenclature is as in the text.

1.2. Oxylipins

The term oxylipin was defined by Hamberg and colleagues in 1991 to constitute a large group of oxygenated lipids formed from fatty acids by reaction(s) involving at least one step of mono- or dioxygenase-catalyzed oxygenation.¹¹ The term was introduced to address the need to describe oxygenated lipids produced from fatty acids besides arachidonic acid (i.e., noneicosanoids). This definition has since been expanded to generally encompass all fatty acid-derived oxygenated lipids including products of enzymatic and nonenzymatic free radical-catalyzed oxidation and nonradical photosensitized oxidation of monounsaturated fatty acids (MUFAs) and PUFAs.¹² Selected oxylipins exert potent mediator functions in a multitude of biological processes, including inflammation, immune activation, cellular development, ion transport, airway smooth muscle contraction, and thrombosis.¹³⁻¹⁶ These lipid mediators can also play a pivotal role in numerous diseases, including diabetes,¹⁷ obesity,¹⁸ cancer,^19,20 pulmonary,²¹ psychiatric,²² metabolic,^23,24 cardiovascular,²⁵ and autoimmune diseases.²⁶

Oxylipins can be classified according to the number of carbon atoms in the backbone of the fatty acid from which they are produced. In mammalian systems, the most well studied oxylipins are the eicosanoids produced from C20-PUFAs including the ω6 fatty acids AA and DGLA, the ω3 fatty acid EPA, and the ω9 fatty acid Mead acid (MA).^27,28 More recently, the docosanoids produced from C22-PUFAs, including the ω6 fatty acid AdA and the ω3 fatty acids DPAn-3 and DHA, have been investigated.^29,30 The oxygenation of C20-fatty acids from both ω6 and ω3 PUFAs produces a wide array of eicosanoid lipid mediators, including the well-studied prostaglandins (and their autoxidation analogues, the isoprostanes), thromboxanes, hydroxy-eicosatetraenoic acids (HETEs) and leukotrienes.³¹ The 2-series prostaglandins are produced from AA (e.g., PGD₂, TxB₂); however, analogous series can be produced from other C20-PUFAs including DGLA (e.g., PGD₁, TxB₁) and EPA (e.g., PGD₃, TxB₃). The critical role of eicosanoids in the mediation of a plethora of biological processes has been thoroughly studied and reviewed in the literature³¹⁻³⁴ and will not be further discussed. Analogously, the oxygenation of long chain ω3 PUFAs produces the docosanoids including the resolvins, maresins and protectins, collectively referred to as specialized pro-resolving mediators (SPMs).³⁵ The growing interest in SPMs has resulted in several recent reviews of their synthesis and biological activity, and interested readers are referred to those publications.^12,36−39 There is increasing interest in this important class of compounds and the biosynthesis of nonmammalian oxylipins has been reviewed in detail.⁴⁰ Collectively, oxylipins constitute a broad class of lipids that include multiple potent mediators of fundamental biological processes, demonstrating the importance of investigating these molecules.⁴¹

1.3. Octadecanoids

The term octadecanoid refers to the oxygenated products of C18-FAs (including saturated FAs, MUFAs and PUFAs, Scheme 2).³² While the canonical use of the term octadecanoid has referred to the products of the jasmonate pathway,⁴²⁻⁴⁴ the definition can be broadened to be employed in the same fashion as for the eicosanoids and docosanoids. Many C18-FAs are present in the diet, resulting in high endogenous concentrations. The C18-PUFAs are found in high concentrations in triglyceride pools, cholesteryl esters, and membrane phospholipids³⁶ to which they impart important physicochemical properties.^45,46 A significant amount of C18-PUFAs are released from the membranes by phospholipases⁴⁷⁻⁴⁹ and are oxidized by the same enzymatic systems responsible for the formation of eicosanoids and docosanoids to produce the octadecanoids. It should be highlighted that phospholipase activity releases fatty acids from the nuclear and mitochondrial membranes.⁵⁰ Given the high abundance of C18-FAs in the human diet,⁵¹ constitutive levels of many octadecanoids are generally higher than the eicosanoids and docosanoids, with LA- and ALA-derived octadecanoids constituting over 50% of the oxylipins present in tissues.³² However, the role of octadecanoids in human pathophysiology has not been closely investigated. The vast majority of studies related to octadecanoid oxylipins have focused on their role as phytohormones in plant systems, where they occur ubiquitously and regulate biotic and abiotic stress signaling, as well as plant growth and developmental processes, especially via the jasmonate pathway.⁵² The formation and function of octadecanoids in plants have been extensively studied, with a number of thorough reviews and will not be further discussed here.⁵²⁻⁵⁴

Only a few canonical octadecanoids have been investigated in-depth in humans, primarily the monohydroxy products, epoxides and diols produced from LA. In the 1980s, the pioneering work of Ozawa and colleagues identified potent toxic effects exerted by LA-derived epoxides and diols, linking them to the progression and severity of acute respiratory distress syndrome (ARDS).⁵⁵ For these reasons, and because the epoxides are synthesized by leukocytes, the 9(10)- and 12(13)-epoxide regioisomers derived from LA were named leukotoxin and isoleukotoxin, respectively. These efforts were continued by Hammock and colleagues, who have extensively investigated the biology of the LA-derived epoxides and corresponding vicinal diols.⁵⁶⁻⁵⁸ The discovery of the potent bioactivity of these compounds sourced growing interest in LA-derived octadecanoids. In recent years, an increasing number of studies have started to expand our knowledge of how octadecanoids impact human pathophysiology, highlighting their ability to interact with various receptors and their involvement in the correct formation of the skin-water barrier,⁵⁹ pain mediation,⁶⁰⁻⁶² thermogenesis and heat regulation, cell proliferation,^65,66 as well as in inflammation⁶⁷⁻⁶⁹ and immunomodulation.^70,71

1.4. Scope

Given the lack of systematic reviews dedicated to octadecanoids outside of plant systems, the purpose of this review is to provide a detailed compilation of the primary octadecanoids formed from C18-FAs and their biosynthetic mechanisms, and to present a general overview of their biological functions as well as their suitability as biomarkers of diseases. In addition, an overview of the standard synthetic methods for preparation of octadecanoids is provided as well as a summary of the analytical methods used to measure octadecanoids. There is divergent terminology in the literature used for octadecanoids (e.g., terms like OXLAMs and oxiOMEs have been used to describe subclasses of LA metabolites)⁶¹ and a portion of this review addresses the need to standardize the nomenclature with a unifying principle applied to both oxygenated and nitrated fatty acids. The primary focus is on octadecanoids produced from oleic acid (OA), LA, ALA, GLA, and SDA. While octadecapentaenoic acid (ODPA) can be observed in some alga (e.g., dinoflagellates),⁷² it is generally not observed in mammals and ODPA-derived octadecanoids will not be discussed here. In addition, we discuss the conjugated linoleic acid (CLA) and conjugated linolenic (CLnA)-derived nitro-FAs. The less studied octadecanoids produced from C18-FAs possessing conjugated and/or trans-double bonds (e.g., conjugated linoleic acid) as well as minor C18-FAs (e.g., sebaleic acid, the major PUFA in human sebum and skin surface lipids) are discussed if the literature is available (Scheme 2). This review focuses on phase-I mammalian metabolism and includes bacterial and fungal metabolites produced by the symbiotic human microbiome. Nonenzymatic production of octadecanoids is considered independently from the organisms in which it takes place. Nonenzymatic oxidation by reactive oxygen species (ROS) and reactive nitrogen species (RNS) are discussed here; however, they are also broadly reviewed elsewhere.⁷³⁻⁷⁶

2. Proposed Octadecanoid Nomenclature

There is currently no agreed common nomenclature for octadecanoids, with nonstandardized abbreviations often used to describe the same compound (e.g., 12(Z)-10-HOME,⁷⁷ 10-OHODA,⁷⁸ or HYA⁷⁹ are used to name 10-hydroxy-12(Z)-octadecenoic acid, and 9-oxo-OTrE,⁸⁰ 9-oxo-OTA,⁸¹ and 9-oxo-OTE⁸² are used to designate 9-oxo-10(E),12(Z),15(Z)-octadecatrienoic acid). This unclarity complicates the literature as well as proposes challenges for systematic lipid naming and curation approaches (e.g., LIPID MAPS,^83,84 HMDB⁸⁵). We therefore propose a system of nomenclature that is based upon the IUPAC system used for linear eicosanoids, which is well established and broadly used in the literature.⁸⁶ This nomenclature was first proposed by our group in a previous paper,⁷⁷ and it will be expanded and further developed here. The name of each compound starts by numbering the position of the oxygenated function carried by the carbon chain. If relevant (i.e., in the case of enantiopure compounds), the chirality of the carbon carrying the oxygenated function is indicated just after the position number of the function. The number, followed by chirality indication if appropriate, is separated by a dash from the abbreviation of the oxygenated function: “Hp”, hydroperoxy; “H”, monohydroxy; “DiH”, dihydroxy; “TriH”, trihydroxy; “oxo”, ketone; “Ep”, epoxide; “DiEp”, diepoxide, “N”, nitro. This prefix is followed by a capital “O”, indicating that these compounds have an 18-carbon backbone (i.e., octadecanoid), akin to the use of “E” for the 20-carbon backbone (i.e., eicosanoid). Finally, the number of unsaturations (i.e., “enes”) in the carbon chain is indicated: “ME”, monoene; “DE”, dienes; “TrE”, trienes; “TE”, tetraenes. For saturated metabolites, the suffix “DA” is used in the abbreviation to indicate that the compound is saturated (e.g., 9-HODA, 9-HydroxyOctaDecAnoic). In this system, 9-HOTrE will be 9-Hydroxy-OctadecaTriEnoic acid, and 9,10-DiHODA will be 9,10-DiHydroxy OctaDecAnoic acid (Scheme 3). Of note, the prefix “oxo” is recommended for ketone functionality, which is in accordance with IUPAC nomenclature. While the letter “K” has been used extensively (e.g., 9-KODE, 9-KOTrE), “oxo” is the correct terminology (e.g., 9-oxo-ODE, 9-oxo-OTrE).

Isomeric octadecanoids derived from ALA and GLA, which only differ in the position of the double bonds (9(Z),12(Z),15(Z) and 6(Z),12(Z),15(Z) (for ALA and GLA, respectively), are discriminated by the addition of a suffix after their name. Given the lower abundance and minor number of reported octadecanoids produced by GLA, the suffix γ will be expressed only for GLA-octadecanoids. Scheme 4 shows an example for the metabolites produced by hydroxylation of C-9 in ALA and GLA.

This nomenclature does not indicate the position of the double bonds, and several isomers possessing different double bond configuration or position would be named in the same fashion. To address this issue, we propose to use this system of nomenclature for the products generated by canonical bis-allylic attack of the 1,4-pentadiene (i.e., giving products at the first or fifth positions), and to explicitly indicate the position of double bonds for any other metabolites (e.g., 10(E),12(Z)-9-HODE would be named simply 9-HODE, because it results from the bis-allylic attack of the 1,4-pentadiene of LA), whereas the position of the unsaturations is indicated for any other isomer. For example, 10-hydroxy-9(Z),12(Z)-octadecadienoic acid would be expressed as 9(Z),12(Z)-10-HODE. For cis-epoxides and diols, the positions of the double bonds can be neglected for the products of epoxidation of a cis-double bond of the parent fatty acid (and for the diol resulting from the hydrolysis of these epoxides). Thus, 9(10)-EpOME and 9,10-DiHOME are used only for 12(Z)-9(10)-EpOME and 12(Z)-9,10-DiHOME respectively, while the position of the double bonds is indicated for any other isomers. For the triols, this system of nomenclature is used for the products generated by bis-allylic attack followed by a rearrangement described in Scheme 12. Since the mechanisms of formation of triols identified so far only produce compounds with unsaturations as described in Scheme 5, the nomenclature will encompass most triols identified to date. For other regioisomers, the position of the double bonds is indicated (e.g., 9(Z)-12,13,17-TriHOME). If new metabolites with a different system of unsaturations are discovered, the positions will have to be specified in the name.

Scheme 5 — The dashed lines indicate potential position of unsaturations that would need to be explicitly indicated in the naming scheme.

For metabolites possessing multiple oxygenated moieties, the functions are defined in the following order: hydroperoxys > hydroxys > ketones > epoxides > nitro. The position of the first function is indicated, followed by a dash and then indication of the nature of the function: “Hp” for hydroperoxides “OH” for hydroxys, “DiOH” for diols, “TriOH” for triols, “oxo” for ketones, “Ep” for epoxides and “N” for nitro. Then a dash is added before indicating the position of the next function (e.g., 12(Z)-9-OH-10-oxo-OME, 11(E)-13-oxo-9(10)-EpOME, 10(E)-9-OH-12(13)-EpOME: see Scheme 6). The position of the double bonds is always indicated for metabolites presenting multiple functions.

Scheme 6 — Nomenclature is as described in Scheme 3.

This system of nomenclature is not novel, is currently used by multiple research groups in the oxylipin field and is based upon the accepted IUPAC system. The necessity to define it unequivocally arises from the common use of ambiguous abbreviations that can result in unclear identification of compounds. A relevant example is the well-known octadecanoid EKODE, which is a nonenzymatic keto-epoxy metabolite of LA. EKODE is abbreviated as a compound possessing two unsaturations when it is in reality a monounsaturated metabolite. In our proposed system, this compound would be named oxo-EpOME, as illustrated in Scheme 6. As interest in octadecanoids continues to increase, it is important for the field that a common nomenclature system is used when reporting structures. This approach will greatly enhance the ability of lipid curation databases such as LIPID MAPS to include these compounds. The nomenclature adopted in this review for the phytoprostane^87,88 and phytofuran^88,89 species is well established and follows the rules determined by Taber and colleagues for isoprostane species.

3. Sources of Octadecanoids

3.1. Metabolism Overview

In plants, there are four main biosynthetic pathways for PUFA oxidation: the CYP74 enzymes, including allene oxide synthases (AOS),⁹⁰ hydroperoxide lyases (hemiacetal synthases),⁹¹ divinyl ether synthases,⁹² and epoxy-alcohol synthases (EAS).⁹³ First, ALA is oxidized by 15-LOX-1 to form a hydroperoxide on the 13-position (13-HpOTrE), which can be rearranged by divinyl ether synthase to form an ether⁹² or by hemiacetal synthase to form a hemiacetal that is spontaneously cleaved into 9(Z)-12-oxo-9-dodecenoic acid and 3(Z)-hexenal.⁹¹ EAS transforms 13-HpOTrE into 11-OH-12(13)-EpODE.⁹³ Finally, AOS transforms the hydroperoxide into 9(Z),11(E),15(Z)-12(13)-EpOTrE. This transformation is the first step of the jasmonate pathway, originally described as the “octadecanoid pathway”, which produces the phytohormone jasmonic acid. The epoxide is then cyclized by allene oxide cyclase (AOC) into cis-OPDA. The intracyclic double bond is reduced by OPDA reductase and several steps of β-oxidation to give 7-iso-jasmonic acid.⁹⁰ Finally, 9(Z),11(Z),15(Z)-12(13)-EpOTrE can spontaneously rearrange into 9(Z),15(Z)-13-OH-12-oxo-ODE (Scheme 7).

Scheme 7 — This pathway includes the canonical octadecanoid pathway, which leads to the production of the phytohormone jasmonic acid. The synthetic route is referenced if known.

Plants, like animals, produce monooxygenated products (hydroxy and epoxy acids) as well as desaturated products and products possessing conjugated double bonds. Well-studied conversions here are those catalyzed by various nonheme di-iron enzymes.⁹⁴ Action by such enzymes results in the formation of ricinoleic acid, crepenynic acid, and vernolic acid as well as calendic acid and other conjugated fatty acids present in certain seed oils. These products occur in plants only, and to the best of our knowledge there are no parallel pathways in animal tissue. Dimorphecolic acid, a plant-specific product related to 9-HODE, is also produced by a member of this group of enzymes. However, again there is no overlap between plant and animal products since in dimorphecolic acid the double bonds have the 10(E),12(E) configuration whereas they are 10(E),12(Z) in the animal lipoxygenase product 9-HODE.

In mammals, the main enzymes responsible for the formation of eicosanoids and docosanoids, namely lipoxygenases (LOX), cyclooxygenases (COX), and cytochrome P450s (CYP), also utilize C18-FAs as substrates.⁹⁵⁻⁹⁸ The affinity of these enzymes for C18-FAs is generally lower compared to their longer chain analogues; however, lower octadecanoid production can be compensated by the higher substrate abundance.⁹⁹ Enzymatic oxidation is performed with determined stereospecificity, where the produced stereoisomers depend on the specific enzyme and substrate. The octadecanoids produced by enzymatic biosynthesis include alcohols, diols, triols, ketones, and epoxides, or combinations of these functional groups (e.g., hydroxy-epoxides and keto-epoxides). The introduction of multiple functional groups in C18-FA metabolism is generally the result of sequential metabolism. For example, the ALOXE3 gene in human skin encodes epidermal lipoxygenase 3 (eLOX3), which possesses hydroperoxide isomerase activity that is able to transform 12(R)-LOX-dependent LA metabolites to the corresponding hydroxy-epoxides.¹⁰⁰ Similarly, the 12-LOX-derived ALA C-9 monohydroxy products can be further targeted by 15-LOX to produce a nonvicinal diol.¹⁰¹ Nonenzymatic conversion is effected by radical or small reactive molecules such as singlet oxygen. As opposed to enzymatic oxidation, the stereochemistry of these processes is not controlled and nonenzymatic octadecanoids are produced as racemic mixtures. In addition to the same linear structures described for the enzymatic oxidation, nonenzymatic oxidation can produce cyclic compounds like endoperoxides and, for PUFAs possessing at least three double bonds, compounds with 5-atom rings (e.g., phytoprostanes and phytofurans^102,103). Finally, the interaction of conjugated octadecanoids with nitric oxide can yield nitro fatty acids (NO₂-FAs) in both plant and animal systems.^104,105 To our knowledge, fungal NO₂-FA formation has not been reported. The catabolism of octadecanoids is similar to other oxylipins and fatty acids in general, with a combination of initial β-oxidation in conjunction with further rounds of β- and/or ω-oxidation.^41,106 Mitochondrial β-oxidation has been proposed to be a major metabolic regulatory checkpoint for oxylipins during inflammation.¹⁰⁷ However, these pathways have yet to be explicitly explored for the octadecanoids.

3.2. Dietary Octadecanoids

3.2.1. Dietary Intake of C18-PUFAs

LA constitutes ∼7% of the total energy uptake in the United States,⁴⁵ and is now the most highly consumed PUFA in the Western Diet.¹⁰⁸ LA is readily available from vegetable oils (e.g., sunflower, safflower, soybean, corn, and canola) as well as nuts and seeds.¹⁰⁸ Worldwide, there has been a large and rapid increase in the amount of LA consumed in response to the work of Ancel Keys, who recommended a dietary shift from saturated to unsaturated fat for cardiovascular health.¹⁰⁹ In particular, global diets have become high in soybean and corn oils. The health effects associated with consumption of LA are unclear and context dependent,^110−113 and can be associated with genotype.¹¹⁴ The effects of LA supplementation upon octadecanoid levels are also ambiguous. In some studies, dietary levels of LA have been reported to associate with observed octadecanoid levels, for example, increased dietary LA resulted in higher observed LA-derived octadecanoid levels in the brain,¹¹⁵ while lowering dietary LA resulted in lower plasma levels of 9- and 13-HODE (as well as 9- and 13-oxo-ODE).¹¹⁶ However, this relationship is not always clear, for example feeding mice a diet high in LA-derived octadecanoids did not affect the levels observed in liver.^117,118 The literature suggests that while dietary levels of parent PUFAs can affect observed octadecanoid levels, supplementation with octadecanoids directly does not. There are a number of reviews on this relationship for further reading.^119,120

While numerous studies cite the conversion of LA into AA (the precursor of “pro-inflammatory” eicosanoids) to be a pro-inflammatory process, the biochemical reality is more nuanced.^108,121 LA exerts biochemical effects in its own right,^122,123 and the bioactivity of eicosanoids is species and context dependent, including both pro-inflammatory and pro-resolving (e.g., lipoxins) actions.¹²¹ ALA is readily available in vegetable oils (e.g., soybean, canola, flaxseed, and perilla) as well as seeds and nuts (particularly walnuts) and other plant food sources.¹²⁴ Dietary ALA is generally associated with beneficial health effects,¹²⁵ including that of the Mediterranean diet,¹²⁶ and is associated with a reduced risk in all causes mortality.^112,127 However, concerns have also been expressed about associations with cancer.¹²⁷ In terms of other C18-PUFAs, the dietary levels are generally low and they are primarily consumed as supplements including GLA (e.g., borage seed oil, evening primrose oil, black currant, ahiflower, spirulina) and SDA (e.g., black currant, ahiflower, spirulina). Recent work has shown that SDA supplementation with ahiflower oil increases the circulating levels of EPA and EPA-derived oxylipins, but does not affect DHA levels.¹²⁸ CLAs are the most abundant conjugated PUFAs in the food supply, primarily present in foods derived from ruminants.¹²⁹ The less common CLnAs are found in various plant seed oils.^130,131 CLnAs are also present in some foods, with 9(Z),11(E),13(Z)-octadecatrienoic acid (i.e., punicic acid) being found at up to 70% in pomegranate seed oil.¹³⁰ Moreover, the edible seeds of the Chinese cucumber, Trichosanthes kirilowii, can also provide a dietary source of CLnAs (Scheme 8).¹³² This rare oil is currently under investigation as a nutraceutical.¹³³ The health effects associated with dietary PUFA consumption have been extensively studied and consist of an ongoing topic of investigation and will not be further discussed here. Interested readers are directed to several in-depth reviews.^134−138

3.2.2. Dietary Sources of Octadecanoids

Multiple octadecanoids are present in the diet from both plant and animal sources,^32,139−144 which constitutes a significant exogenous source of octadecanoid exposure. For example, in 2017 Mubiru et al. documented the C18-epoxy-FA levels in triglycerides present in 390 foods available in the Belgian market.¹⁴⁵ They reported levels between 12 and 687 mg/kg of total C18-epoxy-FAs with animal source foods, being substantially lower than those from plants. More recently, Koch et al. measured the oxylipin content of food oils and processed foods and reported high concentrations (up to 80 mg per serving) in fried falafel and processed foods such as vegetarian sausage/fish fingers.¹⁴⁶ They proposed that the epoxy-to-diol ratio could be a potential marker for refined oils (e.g., the ratio of 15(16)-EpODE to 15,16-DiHODE), with diols increased in refined oils. In the same year, a study quantified the levels of parent PUFAs and of the main metabolites of LA and ALA in different plant and algae edible oils.¹⁴² The results highlighted a relationship between the levels of parent PUFA and the related oxylipins for all oils except olive and flaxseed oil, which had higher levels of oxylipins in relation to the fatty acids. The levels of oxylipins detected in the different oils were very heterogeneous and spanned several orders of magnitude. Flaxseed oil had the highest absolute levels of ketones and hydroxys from both LA and ALA, whereas canola oil had the lowest levels for HODEs and oxo-ODEs and olive oil for HOTrEs and oxo-OTrEs. Soybean oil was found to be rich in DiHOMEs and DiHODEs, while corn and flaxseed oil had the highest levels of EpOMEs and EpODEs, respectively. The median concentrations determined in this work are summarized in Table 1, expressed in μM as the median value obtained in the various oil samples.¹⁴²

Table 1. Median Concentration Values of Octadecanoids Quantified in Commercial Edible Plant Oils (μM)¹⁴².

octadecanoid^a	parent PUFA	soybean oil	corn oil	canola oil	olive oil	flaxseed oil
13-HODE	LA	1.57	8.40	0.15	14.3	24.1
9-HODE	LA	0.33	5.45	0.22	14.0	68.1
13-oxo-ODE	LA	0.28	0.45	0.12	4.42	25.8
9-oxo-ODE	LA	0.89	2.32	0.68	10.3	2.88
12(13)-EpOME	LA	5.10	13.0	2.88	1.21	13.0
9(10)-EpOME	LA	4.43	17.3	2.55	0.25	9.20
12,13-DiHOME	LA	18.8	9.03	0.98	0.57	0.35
9,10-DiHOME	LA	23.9	15.4	0.36	0.76	0.44
9-HOTrE	ALA	0.99	0.23	0.42	2.09	50.8
13-HOTrE	ALA	0.16	0.42	0.95	0.91	19.3
15(16)-EpODE	ALA	3.59	2.43	3.84	0.22	249
12(13)-EpODE	ALA	0.57	0.21	0.85	0.85	15.1
9(10)-EpODE	ALA	0.74	1.01	1.03	0.18	20.7
15,16-DiHODE	ALA	28.0	1.53	11.2	0.15	7.99
12,13-DiHODE	ALA	0.72	0.19	0.10	0.14	0.381
9,10-DiHODE	ALA	9.16	1.72	0.15	0.70	0.31

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Compound nomenclature is as described in Scheme 3.

Our own preliminary investigations of octadecanoid levels in walnuts have revealed an array of octadecanoids with TriHOMEs > HODEs > oxo-OMEs > HOTEs > EpOMEs > EpODAs > EpODEs > DiHOMEs > DiHODAs > DiHODEs.¹⁴⁷ In general, the most well studied aspect of oxygenated lipids in food is the impact of oxidation on edible oils^148,149 and meat.¹⁵⁰ While the incubation of cooked meat in gastric fluids can promote lipid peroxidation, coincubation with plant-based antioxidants suppressing this activity.¹⁵¹ Moreover using oxygenated stearates in simulated gastric conditions, Márques-Ruiz et al. demonstrated that ∼20–60% of 9(10)-EpODA was converted to 9,10-DiHODA, and ∼50% of 12-HpODA was degraded, while 12-HODA and 12-oxo-ODA were unaltered.¹⁵² Using deuterium labeled compounds, 13-HODE was shown to be absorbed after oral gavage, with an incorporation half-life of 71 min in the rat.¹⁵³ The tracer was incorporated into multiple tissues, but not the brain. In rats, consumption of trilinoleoylglycerol hydroperoxide led to the formation of 13-HODE, oxo-EpOMEs, and aldehydes in the gut, with enterocyte absorption.¹⁵⁴ Animal studies considering the impact of oxidized oil consumption reported decreases in liver and plasma triglycerides and cholesterol.¹⁵⁵ Moreover, gastric cells exposed to 13-HODE increased the production of branched chain amino acids, suggesting a possible beneficial impact of oxidized oil consumption.¹⁵⁶

In contrast to the distribution of oxygenated lipids in foods, for nitro lipids, only (E,Z)-NODEs have been reported as endogenous components of extra virgin olive oil.¹⁵⁷ More typically, the acidic pH of gastric juices can promote conjugated linoleic and conjugated linolenic acid reactions with dietary nitrite (NO₂^–) to form bioactive nitro lipids in vivo.¹⁵⁸ Nitrite is present in many vegetables, including spinach, lettuce and beets, and foods cured with nitrate salts, including many cured meats.¹⁵⁹ Concurrent ingestion of conjugated fatty acids and nitrate increases conjugated NODE levels in plasma, urine and tissues,¹⁶⁰ with demonstrated anti-inflammatory and anti-hypertensive effects.^161,162 The putative health effects associated with dietary nitrate should also be considered within the context of the negative effects on nitrosative stress.¹⁵⁹

Therefore, while some ingested octadecanoids in foods can be substantially altered in the gut, others are unaltered, and some can be formed during the digestion process, with substantial proportions available for host absorption. The potential impact of these exogenous sources of octadecanoids on host metabolism deserves further attention.

4. Enzymatic Biosynthesis of Octadecanoids

Octadecanoids can be produced from MUFAs and PUFAs by the action of different enzymes present in plants and mammals. Analogous to the formation of eicosanoids, the main enzymes known to oxidize C18-FAs are the cyclooxygenases, the lipoxygenases and the cytochrome P450s. These enzymes produce a large panel of alcohols, diols, triols, ketones and epoxides, as well as metabolites possessing a combination of these moieties. C18-FAs are generally poorer substrates for these enzymes compared to their C20 analogues; however, due to their high abundance, the amount of produced octadecanoids is significant. In addition, bacteria and fungi possess their own set of enzymes that can create unique octadecanoids, influencing the octadecanoid profile of the host.

4.1. Cyclooxygenases

Cyclooxygenase (COX) enzymes, also called prostaglandin–endoperoxide synthases and prostaglandin G/H synthases, are heme-containing dioxygenase enzymes that possess oxygenase and peroxidase activity on PUFAs. COX enzymes are homodimers and possess two isoforms (COX-1 and COX-2), each consisting of three distinct domains. On both isoforms, the catalytic domain is located on the C-terminal domain and contains separate oxygenase and peroxidase active sites on opposite sides of the heme cofactor. The N-terminal domain facilitates dimerization and membrane binding, while the third domain is important for membrane binding.³¹ Both oxygenase and peroxidase evidence slight differences in their activities and expression between the two isoforms. The two COX isoforms possess similar V_max and K_m(95) but exhibit differences in expression, tissue distribution, allosteric regulation and substrate specificity. COX-1 is ubiquitously expressed, and its expression sites include blood vessels, prostate, immune cells (monocytes, T-cells), platelets, stomach, resident inflammatory cells, smooth muscles, and mesothelium of many organs.¹⁶³ On the other hand, COX-2 is generally synthesized in response to inflammatory stimuli in many tissues including prostate, immune cells (T-cells, B-cells, monocytes), and stomach, but is also constitutively expressed in the brain, lungs, gut, thymus, kidneys, and blood vessels.^163−165 AA is the main substrate of both isoforms, but LA is a known competitive substrate and inhibitor of prostaglandin formation by COX-1 and COX-2.¹⁶⁶ To a lesser extent, ALA is oxidized by COX-2 and while GLA is a poor substrate of both isoforms, it is a better substrate for COX-2.¹⁶⁷

4.1.1. Enzymatic Mechanism

The COX catalytic domain possesses a heme-contained iron. Fe(III) is converted into iron(IV)-oxoporphyrin-radical via the reduction of a fatty acid hydroperoxide or an organic hydroperoxide. The action of this oxoferryl porphyrin radical on tyrosine³⁸⁵ creates a tyrosyl radical that abstracts a bis-allylic hydrogen and forms a conjugated radical. A dioxygen molecule is trapped by this radical to form a peroxyl radical.¹⁶⁸ In contrast to the formation of prostaglandins from C20-PUFAs, this step is not followed by cyclization and addition of a second dioxygen molecule on C18-PUFAs. Instead, the peroxyl is protonated to yield a hydroperoxide (lipoxygenase-type reaction) that is reduced into a hydroxy group by oxidation of Fe(III) included in the peroxidase active site or by other enzymes such as glutathione peroxidase (Scheme 9). Hydroxylated C18-PUFAs are generated via the same pathway as side products of prostaglandin biosynthesis from C20-PUFAs.³¹

4.1.2. Cyclooxygenase-Derived Octadecanoids

The stereospecific removal of the LA 11-pro(S) hydrogen by COX enzymes forms two hydroperoxides, 13(S)-HpODE and 9(R)-HpODE, that are reduced into two main monohydroxylated products 13(S)-HODE and 9(R)-HODE (Scheme 10).¹⁶⁹ The ratio of 9-HODE and 13-HODE vary depending on the enzyme source (tissue vs. recombinant) and the species. In ovine vesicular gland, COX-1 catalyzes preferentially the oxygenation of the 9-position of LA and the formation of 9- and 13-HODE in an 8:2 ratio. In contrast, recombinant human COX-1 expressed in COS-1 cells yielded 13-HODE and 9-HODE in a 5/1 ratio consuming ∼10% of the provided LA, while under the same conditions human COX-2 transformed ∼35% of LA with a 13-HODE/9-HODE ratio of 8/1.⁹⁵ Both 9- and 13-HpODE can also undergo dehydration to produce the respective oxo-products 9-oxo-ODE and 13-oxo-ODE,¹⁷⁰ or be converted nonenzymatically into hydroxy-epoxides or in the skin by a subsequent LOX reaction.⁶¹ These hydroxy-epoxides can subsequently be hydrolyzed to trihydroxy metabolites (TriHOMEs) via a nonenzymatic pathway or by epoxide hydrolases present in both particulate and cytosolic fractions.¹⁷¹ A TriHOME is a stable end product of the oxidation sequence. It has been shown that the formation of 9-HpODE, 13-HpODE, and TriHOMEs is suppressed by inhibition of COX with indomethacin or acetylsalicylic acid in calf aorta.¹⁷² Contrary to the other substrates that undergo a hydrogen abstraction on the ω8 position, the hydrogen abstraction occurs on the ω5 position of ALA, creating 12-HOTrE as the main product (Scheme 10).

Scheme 10 — Nomenclature is as described in Scheme 3. The synthetic route is referenced if known.

4.2. Lipoxygenases

Lipoxygenases (LOX) are nonheme iron-containing dioxygenases that catalyze the conversion of PUFAs containing a 1,4-pentadiene system into conjugated hydroperoxy-fatty acids. The human genome contains 6 functional LOX genes (ALOX5, ALOX12, ALOX12B, ALOX15, ALOX15B, ALOXE3) encoding for 6 different LOX-isoforms: 5-LOX, 12(S)-LOX, 12(R)-LOX, 15-LOX-1, 15-LOX-2, and eLOX3. These isoforms differ by their expression in different mammalian tissues and by their activities (Table 2).

Table 2. Human Lipoxygenase (LOX) Tissue Expression and Substrate Preference^31,173.

human gene	name	major expression	substrates
ALOX15	15-LOX-1	eosinophils, bronchial epithelium, monocytes, macrophages, dendritic cells, reticulocytes	LA, GLA, ALA, AA, EPA, DHA
ALOX15B	15-LOX-2	hair roots, skin, prostate, macrophages, lung, cornea	GLA, ALA, AA, EPA, DHA (LA is a poor substrate)
ALOX12	12(S)-LOX	skin, platelets, umbilical vein endothelial cells, vascular smooth muscle cells	AA, DGLA, EPA, DHA
ALOX12B	12(R)-LOX	skin, hair roots, tonsil epithelial cells, bronchial epithelial cells	GLA, AA, DGLA (LA, EPA, and DHA are poor substrates)
ALOX5	5-LOX	leukocytes, macrophages, dendritic cell, mast cells, lung, placenta	ALA, AA, 5(S)-HpETE
ALOXE3	eLOX3	skin	9(R)-HpODE, 12(R)-HpETE

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12(S)-LOX is primarily expressed in platelets, while 12(R)-LOX and eLOX3 are most frequently found in skin. 15-LOX-1 is highly expressed in leukocytes and airways endothelial cells whereas 15-LOX-2 is expressed in multiple tissues including skin, prostate, lung, cornea, liver, colon, kidney, brain, and monocytes-macrophages.^31,173,174 Different LOX isoforms present varying affinities with octadecanoid PUFAs. For example, LA is the preferred octadecanoid substrate of 15-LOX-1, and an acceptable substrate for 15-LOX-2.⁹⁶ The selectivity of LOX isoforms evidence species-specific variability. For example, the human 15-LOX-2 isoform produces 13(S)-HpODE while the murine gene Alox15b encodes a 8(S)-LOX enzyme that forms 9(S)-HpODE from LA.^173,175

4.2.1. Enzymatic Mechanism

Nonheme ferric iron Fe(III) performs stereospecific hydrogen atom removal in a bis-allylic position via a mechanism involving proton-coupled electron transfer (PCET). First, an electron is transferred to Fe(III) to produce ferrous ion Fe(II), and the proton is simultaneously trapped by the iron’s hydroxide ligand. A radical rearrangement step creates a more stable conjugated diene. Then, the conjugated radical traps a dioxygen molecule to create a hydroperoxide after protonation via a second PCET mechanism that reforms Fe(III).¹⁷⁶ Under low oxygen pressure conditions, the enzyme’s iron is in the ferrous state (Fe(II)) instead of staying in the ferric state (Fe(III)). Fe(II) can promote degradation of hydroperoxides and cause a Fenton reaction.^177−179 In these conditions, the peroxyl can be cleaved by Fe(II) ions to create an alkoxyl radical. The formation of this alkoxyl radical is responsible for the production of various oxygenated metabolites (ketone, hydroxy-epoxide, keto-epoxide, triol, etc.) (Scheme 11).³¹

In the skin, eLOX3 exhibits a hydroperoxide isomerase activity that converts hydroperoxides to hydroxy-epoxides and ketones, but it does not have an oxygenase activity. In contrast to the classical Fe(II)/Fe(III) mechanism of LOX oxidation, where Fe(III) is the active form, the active species of eLOX3 oxidation is the ferrous ion. Fe(II) initiates a homolytic cleavage of the peroxide bond, creating an alkoxyl radical and a ferric ion possessing a hydroxide ligand. The alkoxyl radical cyclizes to form an epoxyallylic radical. Finally, a radical addition of the hydroxide included in the Fe(III)–OH complex forms a hydroxy-epoxide product and restores the active Fe(II) form. A ketone is formed as a side-product of the catalytic cycle (Scheme 12).¹⁰⁰

4.2.2. Lipoxygenase-Derived Octadecanoids

In humans, 15-LOX-1 and 12(R)-LOX produce, respectively, 13(S)-HpODE⁹⁹ and 9(R)-HpODE^180,181 from LA. In mouse skin, 8(S)-LOX can convert LA into 9(S)-HpODE.¹⁷⁵ These hydroperoxides can in turn be converted to hydroxys, ketones, hydroxy-epoxides, keto-epoxides, and triols (Scheme 13). Hydroperoxides can be reduced to hydroxy groups to produce 13(S)-HODE in human,^65,182 and 9(S)-HODE in mice.¹⁷⁵ The oxo-products 13-oxo-ODE and 9-oxo-ODE are produced from an alkoxyl radical derived from LOX-assisted hydroperoxide cleavage.¹⁸³ 13-oxo-ODE can also be formed from 13-HODE by a NAD⁺ dependent dehydrogenase first identified in rat colon mucosa.¹⁸⁴ The relative formation of the HODEs in mammals via LOX or COX activity is unclear and a topic that requires further investigation.

Scheme 13 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known or indicated with a question mark if unknown. The dotted arrows refer to the formation of compounds in mice for which a primary citation could not be identified in the literature.

Hydroperoxide isomerization generates hydroxy-epoxides and keto-epoxides. In human skin, eight hydroxy-trans-epoxides and keto-trans-epoxides formed from 9(R)-HpODE and 13(S)-HpODE have been identified.⁶¹ The authors hypothesize that the enzymes eLOX3 and CYP2S1 are involved in the formation of these epoxy-metabolites. Isomerization of 9(R)-HpODE creates two hydroxy-epoxides possessing an epoxide at the 9(10)-position and a hydroxy at the 11- or 13-position, as well as the two corresponding ketones. From 13(S)-HpODE, two hydroxy-epoxides possessing an epoxide at the 12(13)-position and a hydroxy group at the 9- or 11-position are created, along with their corresponding ketones.⁶¹ This isomerization is performed nonenzymatically via the action of free radicals. Exposure of 13(S)-HpODE with FeCl₃ and cysteine creates 10(E)-9-oxo-12(13)-trans-EpOME, which has been detected in human plasma.¹⁸⁵ The hydroxy-epoxides can then be opened to generate triol metabolites, either by a nonenzymatic pathway as shown in eosinophils by Fuchs et al.¹⁷² or via the action of an epoxide hydrolase as suggested by Funk et al.¹⁷¹ It has been shown recently that alkaline treatment of the keto-epoxide 11(E)-13-oxo-9(R)(10(R))-trans-EpOME esterified to ceramides in skin form two cyclic metabolites, 10-OH-13-oxo-cyclohexenone and 10-OH-9-oxo-cyclohexenone.¹⁸⁶

Trihydroxy derivatives are end products of C18-PUFA oxidation in mammals.⁵⁹ 12(R)-LOX and eLOX3 are abundantly expressed in human skin. 9(R)-HpODE is isomerized by eLOX3 into 9(R),10,13(R)-TriHOME.⁵⁹ The major product in pig and human skin possesses a 10(S)-configuration coming from an SN₂ hydrolysis step that reverses the configuration at C-10 of the 9(R),10(R),13(R)-hydroxy-epoxide precursor.⁵⁹ This high specificity supports the hypothesis that an epoxide hydrolase enzyme is involved in hydrolysis of hydroxy-epoxides in these tissues.¹⁸⁷ In a cell line of human mast cells (LAD2 cells), TriHOME formation is LOX-independent and yields a 13(R) configuration, whereas in eosinophils this synthesis is initiated by 15-LOX, resulting in a 13(S)-configuration.¹⁷² Epidermal triols are almost exclusively esterified to ceramides and, together with epidermal keto-epoxides, possess an important role in maintaining the integrity of the water–skin barrier.⁵⁹ Notably in plants, TriHOMEs contained in cuticle waxes have antifungal properties,¹⁸⁸ and a parallel role in fungal defenses of the epithelial barrier can be postulated in animals.

LOX metabolites of ALA are of major importance in plants, although in animal systems ALA metabolism is less extensively studied, and only a small number of metabolites have been reported to date. The major product of ALA incubation with 15-LOX-1 is 13-HpOTrE, which is further reduced to 13-HOTrE.^189,190 Finally, the diol 10(E),12(Z),14(E)-9(S),16(S)-DiHOTrE has been identified after further oxidation of 9(S)-HpOTrE (an ALA-metabolite of plant 5-LOX) by human 15-LOX-2.¹⁰¹ The biosynthesis of TriHODEs has not yet been reported (Scheme 14).

Scheme 14 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known or indicated with a question mark if unknown. The dotted arrows refer to the formation of compounds for which a primary citation could not be identified in the literature.

The metabolism of sebaleic acid by 5-LOX in neutrophils results in the production of 5-HODE, which can be further metabolized into 5-oxo-ODE by 5-hydroxyeicosanoid dehydrogenase in neutrophils and keratinocytes. The formation of 6(E),8(Z)-5(S)-18-DiHODE and 6(E),8(Z)-18-OH-5-oxo-ODE from sebaleic acid have also been reported, but the responsible enzymes remain unknown (Scheme 16).¹⁹¹

Scheme 16 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

Very few studies have been performed to identify mammalian SDA-derived octadecanoids. However, Arterburn et al. did report that the incubation of SDA with porcine 12-LOX followed by reduction with NaBH₄ of the product led to the identification of two hydroxy-metabolites: 10-HOTE and 16-HOTE (Scheme 15).¹⁹²

Scheme 15 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

4.3. Cytochrome P450s

In mammals, CYPs are important enzymes that participate in oxidative, peroxidative, and reductive metabolism of a broad range of lipid substrates. CYPs are a superfamily of heme-containing monooxygenases that are ubiquitous across all domains of life.¹⁹³ Humans possess 57 functional CYP-encoding genes, and their products can be divided into 18 families and 41 subfamilies. The CYP2, CYP3, and CYP4 families contain far more genes than the other 15 families. CYPs are widely distributed in mammalian tissues and are highly expressed in the lung, liver, brain, and kidney. Most CYPs are bound to either mitochondrial membranes or the endoplasmic reticulum, but they are also present in the plasma membrane and the nucleus. Their major functions include drug and lipid metabolism.¹⁹⁴ CYPs catalyze four different reactions: ω-side chain hydroxylation, epoxidation, bis-allylic hydroxylation, and hydroxylation with double-bond migration (“allylic hydroxylation”).⁹⁸

4.3.1. Enzymatic Mechanism

CYP-catalyzed oxygenations are NADPH-dependent and involve the scission of dioxygen with one oxygen atom reduced to water and the other incorporated into the substrate. In the prototypical reaction cycle, substrate recruitment displaces H₂O from the axial position of the heme Fe(III). A NADPH-cytochrome P450 reductase undergoes a 1-electron reduction to form Fe(II), then O₂ binding creates an oxygen–iron complex, and another reductase-assisted 1-electron reduction generates a negatively charged iron(III)-peroxo complex. Double protonation of this iron(III)-peroxo complex followed by the scission of a dioxygen molecule creates an iron(IV)-oxoporphyrin-radical cation intermediate, a direct oxidant in many CYP oxidation reactions. This oxoferryl porphyrin is responsible for hydroxylation reactions and undergoes a hydrogen abstraction followed by a radical insertion of an hydroxy group that recreates the starting Fe(III) ion (Scheme 17).^195,196

The mechanism of epoxidation by CYPs includes formation of a π-complex between the iron(IV)-oxoporphyrin-radical cation intermediate and the alkene. The reaction proceeds with the insertion of the oxyl entity that initiates the formation of a radical intermediate. The radical scission of the oxygen–iron bond creates an epoxide and reforms Fe(III) ion (Scheme 18).

4.3.2. Cytochrome P450-Derived Octadecanoids

Epoxides/Diols. CYPs oxidize LA (Scheme 19), ALA (Scheme 20), GLA, and OA (Scheme 21) into a wide range of alcohols and epoxides. LA possesses two double bonds, each of which can undergo a CYP-catalyzed epoxidation to create 9(10)-EpOME and 12(13)-EpOME. Epoxide hydrolase-catalyzed epoxide hydrolysis forms the corresponding vicinal diols 9,10-DiHOME and 12,13-DiHOME.¹⁹⁷ In humans, CYP2J2 does not possess any hydroxylation function, resulting in the sole production of EpOMEs from LA.¹⁹⁸ CYP2E1¹⁹⁸ and CYP2C9¹⁹⁹ are the major human LA epoxygenases but are also responsible for the formation of monohydroxylated species. The other CYPs responsible for epoxide and diol formation in humans are CYP1A2, CYP2E1, CYP2C8, CYP2C9, CYP2C19, CYP2J2, CYP2J3, CYP2J5, CYP2J9, and CYP3A4.^97,198,200 These enzymes produce EpOMEs and DiHOMEs in mixture with the HODEs. In the same fashion as LA, ALA can be epoxidized by CYP enzymes on its three unsaturations to generate 9(10)-EpODE, 12(13)-EpODE, and 15(16)-EpODE.²⁰¹ The monoepoxides can be hydrolyzed into 9,10-DiHODE, 12,13-DiHODE, and 15,16-DiHODE.⁹⁹

Scheme 21 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known. The dotted arrows refer to the formation of compounds for which a primary citation could not be identified in the literature.

It is known that the cis- or trans-configuration of an epoxide has a strong impact on the opening reaction catalyzed by sEH to form vicinal diols. For example, in the case of alkyl-phenyloxirane, the cis-epoxides have been found to be poor substrates for sEH. Among the two trans-epoxides (R,R) and (S,S), both plant and mammalian sEH have been shown to be enantioselective. Mouse and human sEH are both enantioselective for (S,S)-alkyl-phenyloxiranes, while cress sEH is enantioselective for (R,R)-alkyl-phenyloxiranes.²⁰²

Relatively recent studies demonstrated the lipoxygenase-catalyzed formation of hydroxy-endoperoxides from EpETrE-type fatty acid epoxides.²⁰³ One example is the transformation of 15(R),16(S)-EpODE by soybean LOX-1. In association with the expected formation of a 13(S)-hydroperoxide, intramolecular nucleophilic substitution (S_Ni) between the hydroperoxy (nucleophile) and 15(16)-epoxy group (electrophile) forms a 13(S),(15(S))-epidioxy-16(S)-HODE (Scheme 20). The configuration of the endoperoxide (cis or trans side chains) was shown to depend on the steric relationship of the new hydroperoxy moiety to the enantiomeric configuration of the fatty acid epoxide.²⁰³ The results were proposed as a potential interaction of lipoxygenase and cytochrome P450 product formation.

GLA is also a CYP substrate and its three double bonds can be epoxidized by CYP2B1, giving 6(7)-EpODE-γ, 9(10)-EpODE-γ, and 12(13)-EpODE-γ.²⁰¹ 12(13)-EpODE-γ as well as a smaller amount of 9(10)-EpODE-γ can also be generated by CYP2CAA and CYP2C2.²⁰¹ CYPs (2C8, 2C9, 3A7, and 3A4 mainly) can also oxidize monounsaturated fatty acids to form the epoxide and diol metabolites. For example, 9(10)-epoxy-octadecanoic acid (i.e., 9(10)-EpODA) and 9,10-dihydroxy-octadecanoic acid (9,10-DiHODA) are produced from OA.^204,205

Alcohols. Certain CYPs are also able to catalyze bis-allylic hydroxylation of LA to produce 11-HODE.^200,206 This bis-allylic alcohol is unstable under acidic conditions and rearranges into racemic 9-HODE and 13-HODE. In human liver microsomes, the main CYPs responsible for the formation of 9(Z),12(Z)-11-HODE are CYP1A2 and CYP3A4.²⁰⁰ Allylic hydroxylation of LA also produces 9-HODE and 13-HODE possessing an (R)-configuration. The CYP responsible for the allylic hydroxylation of LA are CYP1A2, CYP2C9, and CYP3A4.⁹⁷

Finally, LA can be hydroxylated in the 5 positions ω1 to ω5, creating 9(Z),12(Z)-14-HODE, 9(Z),12(Z)-15-HODE, 9(Z),12(Z)-16-HODE, 9(Z),12(Z)-17-HODE, and 9(Z),12(Z)-18-HODE.^{97,200,207,208} CYP2C9 can generate all of these metabolites, but other CYPs are specific for hydroxylation at single positions.⁹⁷ For example, CYP2C19⁹⁷ and CYP2E1²⁰⁷ form 9(Z),12(Z)-17-HODE as the major product, and CYP4A1 and CYP4A11 form primarily 9(Z),12(Z)-18-HODE.^207,208 CYP1A2, CYP2C8, CYP2C9, CYP2C19, and CYP3A4 can catalyze hydroxylation on both the ω1 and ω2 positions.²⁰⁰ The ω1-hydroxylation of ALA to produce 9(Z),12(Z),15(Z)-18-HOTrE, has been reported by CYP2U1.²⁰⁹ Mammalian CYP2E1 can form the ω1 and ω-hydroxylated products 9(Z)-17-HOME and 9(Z)-18-HOME from OA,²⁰⁷ and 17-HODA and 18-HODA are produced from SA in rat microsomes in a NADPH-dependent reaction.²¹⁰ The formation of 8-HODE from LA by CYP2C9 has been reported.⁹⁷

Hydroxy-epoxides/Nonvicinal Diols. A few LA metabolites derived from a combination of these activities have also been observed. CYP4F3 enzymes are the main enzymes involved in the oxidation of epoxy-FAs, and are responsible for the ω1-hydroxylation, and to a lesser extent (ω2)-hydroxylation, of EpODA and EpOMEs. The ratios of ω/(ω1) hydroxylation were 8 and 7 for 9(10)-EpODA and 9(10)-EpOME, respectively, but decreased to 1.6 for 12(13)-EpOME.^211,212 CYP4F11 produces 3,18-DiHODA by ω1-oxidation of 3-HODA in human liver.¹⁷¹

5. Microbial and Fungal Sources

5.1. Gut Microbiota Octadecanoids

Microbes and bacteria that constitute the gut microbiome are human symbionts that exert a vital function in the health status of their hosts.²¹³ Among the various interactions, the gut microbiota exert profound effects on the host metabolism by transforming dietary fatty acids into bioactive molecules.^214,215 Lipid metabolites produced by the gut microbiota can impact human health, interacting with different organ systems including gut–artery,^216,217 gut–brain,²¹⁸ and gut–lung^219,220 axes. Gut dysbiosis can therefore induce multiple physiological consequences. For example, neonates at risk of childhood atopy, eczema, and asthma exhibit perturbation of the gut microbiome, including an increased number of bacterial epoxide hydrolase genes.^70,71 This enzyme forms the 12,13-DiHOME, which has physiological functions associated with fatty acid catabolism and energetics^63,221 but also exerts deleterious effects on lung function in patients with severe burns or COVID-19 infection.^68,71,222 Moreover, specific lipid metabolites produced by the gut microbiota have been shown to confer resistance to inflammation caused by high fat diet (HFD)-induced obesity in mice.⁷⁹ Accordingly, there are multiple health and physiological effects associated with microbiome-derived octadecanoids that are deserving of investigation.

The metabolism of C18-FAs by gastrointestinal anaerobic microbiota (e.g., lactic acid bacteria), generates multiple conjugated fatty acids and trans-fatty acids that can affect host lipid metabolism.²²³ Lactic acid bacteria transform growth-inhibiting PUFAs containing cis–cis pentadiene moieties into less toxic saturated fatty acids.²²⁴ In the process, CLAs, and trans-FAs are released into the host tissues. In ruminants, a multitude of these intermediate products are generated and may end up in consumed meat as well as milk.^225,226 CLA defines a broad family of isomers possessing at least two conjugated double bonds, that exert a variety of effects on human health, the nature of which is still debated. However, abundant species such as rumenic acid and its isomer 10(E),12(Z)-octadecadienoic acid, were found to possess anticarcinogenic activity.²²⁷ The role of CLA^227,228 and CLnA^229,230 in cancer prevention have been reviewed. The group of 28 CLA isomers represents a significant source of substrate for downstream octadecanoids that have been little explored. The effects of trans-fatty acids on human health are clearer. For example, 10(E)-octadecenoic acid, the major isomer typically found in meat and milk, has been shown to have adverse effects on cardiovascular health.^231,232 The bioactivity and health effects of ruminant meat lipids has been extensively reviewed by Vahmani et al.²³³

Specific octadecanoids are also produced by most lactic acid bacteria and possess an alcohol or a ketone at the 10- or 13-position. The bacterial pathways leading to both series of metabolites were first described by the pioneering work of Jun Ogawa and colleagues.²²³ It is important to highlight that these enzymatic reactions occur in an anaerobic environment, and the introduction of the hydroxy group occurs via hydration and not oxidation. Hira et al. characterized the production of a large number of C18-FA-derived metabolites by gut lactic acid bacteria.²³⁴ In a representative gut bacterium Lactobacillus plantarum from a mouse model, four enzymes involved in the bacterial metabolism of PUFAs were identified: CLA-hydrolase (CLA-HY), CLA-dehydrogenase (CLA-DH), CLA decarboxylase (CLA-DC), and CLA-enoyl reductase (CLA-ER).²²³ The first reaction of LA metabolism is the hydration of the 9,10-alkene catalyzed by CLA-HY, which exhibits both hydratase and dehydratase activities and converts LA into 12(Z)-10(S)-HOME. In Lactobacillus acidophilus, this step can also be catalyzed by the enzyme fatty acid hydratase 2 (FA-HY2).²³⁵ The resulting monohydroxylated metabolite is subsequently dehydrated by CLA-HY to generate the conjugated-LA derivative 10(E),12(Z)-ODE. The next step is the oxidation of 12(Z)-10-HOME catalyzed by CLA-DH in the presence of NAD⁺. CLA-DH possesses both dehydrogenase and oxidoreductase activities and generates the oxo-product 12(Z)-10-oxo-OME. It can also catalyze the reverse reaction and reduce the ketone into the corresponding hydroxy in the presence of NADH. The Δ¹²-cis double bond is isomerized by acetoacetate decarboxylase CLA-DC, to form the more stable conjugated enone product 11(E)-10-oxo-OME. This enone is saturated by the enone reductase CLA-ER to generate 10-oxo-ODA in the presence of FAD/FMN and NADH. The 10-oxo-ODA is then reduced by CLA-DH into the corresponding alcohol 10-HODA. The final elimination of the hydroxy by CLA-HY creates OA and 10(E)-OME. CLA-DY can also catalyze the reduction of 11(E)-10-oxo-OME into 11(E)-10-HOME, which is dehydrated by CLA-HY into two conjugated-LA compounds 9(Z),11(Z)-ODE and 9(E),11(Z)-ODE (Scheme 22).²²³ Additional C18-PUFAs, ALA (Scheme 23) and GLA (Scheme 24), are also metabolized by these enzymes following the same pathway.²²³ For example, FA-HY2 efficiently converts ALA, GLA, and OA into the corresponding 10-hydroxy fatty acids.²³⁵

Scheme 22 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

Scheme 23 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

Scheme 24 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

Most gut lactic acid bacteria generate the same metabolites as Lactobacillus plantarum. However, some lactic acid bacteria such as Lactobacillus acidophilus can also produce metabolites that are hydrated at the 13-position. The enzyme responsible for the hydration of C18-PUFAs on their 13-position has been identified in Lactobacillus acidophilus NTV001 as a hydratase termed FA-HY1.²³⁵ This enzyme can react with all C18-PUFAs possessing a Δ¹²cis-double bond including LA, ALA, GLA, and SDA. Incubation of PUFAs with FA-HY1 produces 9(Z)-13(R)-HOME from LA, 9(Z),15(Z)-13(S)-HODE from ALA and 6(Z),12(Z)-10-HODE from GLA. 9(Z)-13(R)-HOME is further metabolized by FA-HY1 into 9(Z),13(E)-ODE.²³⁵ The stereochemistry of 9(Z)-13(R)-HOME, produced from LA by FA-HY1 in the strain NTV001 of L. acidophilus, is distinct from that of the previously reported 9(Z)-13(S)-HOME produced by the strain 13951 of L. acidophilus.²³⁶ Through the screening of ∼300 strains of lactic acid bacteria, Pediococcus sp. AKU 1080 has also been identified as a strain with the ability to hydrate LA into three hydroxy-fatty acids, 12(Z)-10-HOME, 9(Z)-13-HOME, and 10,13-DiHODA. This last metabolite can be obtained from both 12(Z)-10-HOME and 9(Z)-13-HOME.²³⁷ The corresponding ketones have also been identified as gut bacteria metabolites; however, the enzymes responsible for their formation remain unknown.²³⁴

5.2. Other Bacterial and Microbial Metabolites

Multiple types of bacteria and microbes produce hydroxy fatty acids from different unsaturated fatty acids. Among them, Streptococcus,²³⁸Nocardia,²³⁹ and Flavobacterium(240) convert LA into 12(Z)-10-HOME. In addition to these well-known metabolites, some bacteria can produce unique metabolites of PUFAs, including diols, triols, tetrahydrofuranyl fatty acids (THFA), and bicyclic-fatty acids.

A bacterial source of C18-PUFA metabolites has been extensively studied by Hou and colleagues.²⁴¹ The bacterium responsible for the formation of a large range of unique PUFA metabolites is the strain Bacillus megaterium ALA2, which has been shown to effectively oxidize LA at the 7-, 12-, 13-, 16-, and/or 17-position, producing numerous unique triols and cyclic PUFA metabolites (Scheme 25). The production of an uncommon triol compound, 9(Z)-12,13,17-TriHOME, from LA by a microbial culture of B. megaterium ALA2 isolated from a dry soil sample, has been reported.²⁴² This triol is the major product formed from LA by this bacterial strain, and is generated from 12,13-DiHOME.²⁴³ It is a precursor of three diepoxy bicyclic fatty acids, 9(Z)-12(17);13(17)-DiEpOME, 9(Z)-7-OH-12(17);13(17)-DiEpOME, and 9(Z)-16-OH-12(17);13(17)-DiEpOME that have been identified from LA.^244,245 The generation of a second original triol (9(Z)-12,13,16-TriHOME), has also been reported by the same group. This second TriHOME can be epoxidized to a THFA (9(Z)-12-OH-13(16)-EpOME), which can be further metabolized into another THFA (9(Z)-7,12-DiOH-13(16)-EpOME).²⁴⁶

Scheme 25 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

B. megaterium ALA2 also produces THFA from ALA and GLA (Scheme 26). In 2003, Hosokawa et al. identified two new ALA metabolites, 9(Z)-13,16-DiOH-12(15)-EpOME and 9(Z)-7,13,16-TriOH-12(15)-EpOME,²⁴⁷ and three new GLA metabolites, 6(Z),9(Z)-12(17);13(17)-DiEpODE, 6(Z),9(Z)-12,13,17-TriHODE, and 6(Z),9(Z)-12-OH-13(16)-EpODE.²⁴⁸

Cyanobacteria possess mini-LOX enzymes, which show only the LOX’s C-terminal catalytic domain that contains iron. The mini-LOX of Cyanothece sp. (CspLOX₂) creates bis-allylic hydroperoxides, 11(R)-HODE from LA, as well as 9(R)-HODE and 13(S)-HODE.²⁴⁹ The lipoxygenase domain of a catalase-LOX fusion protein from the cyanobacterium Acaryochloris marina converts LA to 9(R)-HpODE and oxygenates omega-3 fatty acids at the ω7 carbon, forming 12(R)-HpOTrE from ALA.²⁵⁰

Microbial and bacterial transformations of OA have been extensively studied. OA can be metabolized by a large range of microbes and bacteria, most of which oxidize the 10-position, but oxidation of the 7-position has also been reported.^241,251,252Nocardia cholesterolicum,^239,241Rhodococcus sp.,²³⁹ and Flavobacterium DS5(253) can hydroxylate OA into 10-HODA. The corresponding ketone, 10-oxo-ODA, can be produced by Flavobacterium DS5,²⁵² as well as by fungal strains such as Saccharomyces sp. and Candida sp..²⁵¹ In their effort to convert agricultural oils to value-added industrial chemicals, Hou et al. extensively studied the bioconversion of OA by the bacterial stain Pseudomonas PR3. They reported the formation of a new dihydroxylated compound, 8(E)-7,10-DiHOME.²⁵⁴ In addition, they identified a monohydroxylated metabolite produced by the same strain, 8(Z)-10-HODE, and hypothesized it to be a potential intermediate in the bacterial biosynthesis of 8(E)-7,10-DiHOME.²⁵⁵ Guerrero et al. continued this study and showed that 8(E)-10(S)-HOME is generated by Pseudomonas 42A2 from OA and can be transformed into 8(E)-7,10-DiHOME.²⁵⁶ The enzyme responsible for this transformation was reported to be a LOX-like enzyme.²⁵⁷ Finally, OA can be epoxidized into 9(10)-EpODA by the enzyme CYP107N3 of Streptomyces peucetius (Scheme 27).²⁵⁸

Scheme 27 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

5.3. Fungal Metabolites

Fungi can penetrate an organism via alimentation (mushrooms) or by the airways due to their presence in inhaled air.²⁵⁹ Fungi can generate biologically active PUFA metabolites that are also produced in humans (e.g., 13-HODE, EpOMEs), which may interfere with the host metabolism as shown for eicosanoids.²⁶⁰ It is known that fungal octadecanoids can interfere in plant signaling pathways by mimicking endogenous oxylipins;^261−263 however, the influence of fungal-generated endogenous octadecanoids in animals is mostly unexplored. Fungi also possess diverse enzymes that exhibit unusual catalytic activities and can generate a high diversity of specific metabolites, whose effects on animals remain to be investigated.

The transformation of LA in mushrooms has a long history aimed at unravelling the mechanistic basis for formation of the characteristic mushroom aroma of the 8-carbon alcohol, 3(R)-1-octen-3-ol. In the 1980s, Wurzenberger and Grosch uncovered the LA metabolism leading to this 1-octen-3-ol, identifying the two steps as the oxygenation of LA to 10(S)-HpODE followed by its cleavage to the 10-carbon acid-aldehyde 8(E)-10-oxo-decanoate and the 8-carbon alcohol.^264−266 They also showed that ALA is converted via the equivalent transformations, generating 1,5(Z)-octadien-3-ol and 2(Z),5(Z)-octadien-1-ol.²⁶⁷ It is apparent that a LOX enzyme would be incapable of oxygenating LA to the 10-hydroperoxide and therefore the participation of a heme oxygenase-peroxidase has been long suspected. In 2012, a cyanobacterial catalase-heme dioxygenase fusion protein was show to transform LA to the two products.²⁶⁸ The proof of the conjecture with mushroom proteins came quite recently with the characterization of a heme dioxygenase in Coprinopsis cinerea;²⁶⁹ the expressed cDNAs of two heme dioxygenase-P450 fusion proteins converted LA to the 10(S)-HpODE. At least in this mushroom species, the second step of hydroperoxide lyase cleavage was not catalyzed by the expressed cDNAs and the hydroperoxide lyase enzyme remains uncharacterized.

The understanding of fungal enzymatic systems involved in the production of octadecanoids has its roots in the seminal work of Ernst Oliw who investigated these mechanisms in various fungal species. Dioxygenase–cytochrome P450 fusion enzymes (DOX-CYP) are common fungal enzymes that are divided into seven subfamilies (5,8-LDS, 7,8-LDS, 8,11-LDS, 10(R)-DOX, 10(R)-DOX-EAS, 9(R)-DOX-AOS, and 9(S)-DOX-AOS).²⁷⁰ Linoleate diol synthases (LDS) contain a heme and exhibit two related enzyme activities. They catalyze the dioxygenation of the 8-position of LA and the isomerization of a hydroperoxide group into dihydroxy-LA.²⁷¹ LDS oxidize LA to 9(Z),12(Z)-8(R)-HpODE (8(R)-DOX activity), which is further metabolized into 9(Z),12(Z)-5,8-DiHODE by 5,8-LDS (in Aspergillus sp.),^272,273 9(Z),12(Z)-7,8-DiHODE by 7,8-LDS (in Gaeumannomyces graminis(274,275) and Magnaporthe grisea(276)), and 9(Z),12(Z)-8(R),11(S)-DiHODE by 8,11-LDS (in Aspergillus clavatus,²⁷⁷Aspergillus fumigatus,²⁷³ or Penicillium chrysogenum(278)). 10(R)-DOX-EAS oxidizes LA into 10(R)-HpODE, which is further metabolized into 8(E)-10-OH-12(13)-EpOME. This enzyme also produces smaller amounts of monohydroxylated metabolites 9(Z),12(Z)-8-HODE and 10-HODE, and it has also been shown to oxidize the 8- and 10-position of ALA and OA, but not of GLA, which is a poor substrate.²⁷⁹ 9(R)-DOX-AOS and 9(S)-DOX-AOS oxidize LA to 10(E),12(Z)-9(S)(10)-EpODE and 10(E),12(Z)-9(R)(10)-EpODE.²⁷⁹ The production of 9(Z),12(Z)-17-HODE, 9(Z),12(Z)-8,17-DiHODE, and 9(Z),12(Z)-8,16-DiHODE in vitro from LA by an of as yet undetermined CYP has also been reported in the rice blast fungus M. grisea.²⁷⁶

In the cytosolic fraction of G. graminis, the products of LA hydroxylation to ω2 or ω3 alcohols and of ALA’s ω3 unsaturation epoxidation and hydrolysis have been observed, but the responsible CYP(s) remain unknown (Scheme 28).²⁷⁵

Scheme 28 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known. The scheme is split into two due to size limitations (see Scheme 29).

Fungi also possess LOX enzymes containing a manganese ion instead of the more common ferric ion in their active site.²⁸⁰ The G. graminis 13(R)-MnLOX, also called Mn-LO, forms 13(R)-HpODE and 13(R)-HpOTrE from LA and ALA, respectively. 9(Z),12(Z)-11(S)-HpODE is first produced and then rearranges into 13(R)-HpODE via a linoleoyl radical.²⁸¹ GLA is a poor substrate for Mn-LO, but minor amounts of 11-HOTrE-γ and 13-HOTrE-γ have been observed.²⁸⁰

A fungal 9(S)-Mn-LOX and an epoxy-alcohol synthase have been found in the mycelium of the rice stem pathogen, Magnaporthe salvinii. This new Mn-LOX generates 9(S)-HpODE from LA, which is further metabolized by EAS into 12(Z)-9(S)-OH-10(R)(11(R))-trans-EpOME. The hydrolysis of the epoxide generates two triols, 9(S),10,11-TriHOME and 9(S),12,13-TriHOME. ALA is metabolized with little positional specificity at the 9-, 11- or 13-position into 9(S)-, 11(R)- or 13(R)-HpOTrE. 9(S)-HpOTrE is further metabolized into 10(E),12(Z),14(E)-9,16-DiHOTrE (Scheme 29). Finally, 9(S)-Mn-LOX can oxidize GLA into 9-HOTrE-γ.²⁸²

Scheme 29 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known. The scheme is split into two due to size limitations (see Scheme 28).

Another original enzyme found in Fusarium oxysporum is an iron-containing 13(S)-LOX named FoxLOX. This enzyme was determined to act as a regular 13-LOX when LA or GLA were used as substrates; however, further metabolism of ALA-derived hydroperoxy products has been observed. Among the products formed from ALA, an hydroxy-epoxide (9(Z),15(Z)-11-OH-12(S)(13(S))-trans-EpODE), a ketone (13-oxo-OTrE), and two diols (10(E),12(Z),14(E)-9,16-DiHOTrE and 15,16-DiHOTrE) have been identified.²⁸³

A fungal catalase (Fg-cat) from the fungus Fusarium graminearum, possessing a 13(S)-peroxidase activity, has been identified by Teder et al. This new enzyme is responsible for the production of a keto-epoxide from LA (10(E)-9-oxo-12(13)-cis-EpOME), which is generated as the main product from 13(S)-HpODE, with the trans-epoxide detected as a minor product. Fg-cat can also oxidize ALA into 13(S)-HOTrE, which is further metabolized into a keto-epoxide (10(E),15(Z)-9-oxo-12(13)-EpODE) as well as a diepoxide (10(E)-9-oxo-12(13);15(16)-DiEpOME).²⁸⁴

Finally, fungi possess unspecific peroxygenases that metabolize PUFAs into epoxides. OA is an excellent substrate for these enzymes and is oxidized into 9(10)-EpODA (main product). ALA is metabolized into monoepoxides and 14-OH-EpODEs, by the UPO enzyme of Collariella virescens.²⁸⁵ LA is a poorer substrate for this enzyme, but can also be oxidized into 9(Z)-11-HOME and 12(13)-EpOME (main products) as well as 9(10)-EpOME and 9(Z)-11-OH-12(13)-EpOME in C. virescens.²⁸⁵ Production of 9-oxo-OTrE and 13-oxo-OTrE from ALA by the fungus Aspergillus niger has been described by Petta et al. in 2014, as well as the production of a unique metabolite (11(E),16(E)-9,10,13-TriOH-15-oxo-ODE). However, the enzymes involved in the production of these three metabolites have not been investigated (Scheme 29 and Scheme 30).⁸²

Scheme 30 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

In the commonly eaten champignon (Agaricus bisporus also called Psalliota bispora), several unique metabolites were discovered, but the enzymes responsible for their formation remain unknown. Identified LA metabolites include 9(Z),12(Z)-8(R)-HODE and 9(Z),12(Z)-8(R),11(S)-DiHODE as the main products, and 8(E),12(E)-10-HODE, 10(E),12(Z)-8,9-DiHODE and 9(Z),11(E)-8,13-DiHODE, 12(Z)-8(R)-HOME, 9-HODE and 13-HODE as secondary products. OA, ALA and GLA are also converted into their 8-hydroxy derivatives and, in the case of ALA, into the nonvicinal diol 9(Z),12(Z),15(Z)-8,11-DiHOTrE (Scheme 31).²⁷¹

Scheme 31 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

6. Nonenzymatic Biosynthesis of Octadecanoids

In addition to enzymatic biosynthetic pathways described in the previous chapter, a high diversity of octadecanoids can be formed nonenzymatically by the action of free radicals or small reactive molecules such as singlet oxygen.²⁸⁶ Endogenous sources of reactive oxygen include the CYPs,²⁸⁷ xanthine oxidoreductase,²⁸⁸ NADPH oxidases,²⁸⁹ flavoproteins, myeloperoxidase, as well as normal mitochondrial function as a consequence of cellular respiration.²⁹⁰ The release of ROS by NADPH oxidase is stimulated by saturated²⁹¹ and unsaturated C18-FAs.^292,293

If dysregulated by external (e.g., exposure to pollution or smoking) or internal (e.g., neurodegenerative or inflammatory) insults, activation of these systems can lead to oxidative stress with diverse ramifications. For instance, within the mitochondria, increased ROS production leads to the oxidation of the LA rich cardiolipin, and subsequent phospholipase activity can release an array of species including oxo-OMEs, HODEs, EpOMEs, HpODEs, DiHODEs, and Hp-oxo-ODEs.²⁹⁴ Moreover, radical leakage into tissues can oxidize proteins, lipids, and DNA, leading to their degradation.²⁹⁵ Most of these radicals are ROS such as hydroxyl radical (HO^•), the most reactive radical for peroxidation of PUFAs. ROS comprise the nonradicals singlet oxygen and hydrogen peroxide, and encompass the highly reactive hydroxyl radical (HO^•), superoxide radical (O^2•–), and fatty acid peroxyl and alkoxyl radicals that strongly enhance the propagation of lipid peroxidation. Superoxide radical, by itself a weak oxidant, participates in the Fenton reaction,²⁹⁶ and in the formation of ¹O₂ and of H₂O₂ by spontaneous dismutation.^297,298

The chemical nature of LA oxidation products was studied principally by Gardner’s group in the 1970s and early 1980s. Radical reaction of oxygen with PUFAs has been extensively reviewed by Gardner in 1989,²⁹⁹ and the nature and formation mechanisms of nonenzymatic LA metabolites has been reviewed by Spiteller in 1998.³⁰⁰ Linear products as well as endoperoxides are formed from LA, whereas cyclic metabolites such as phytoprostanes or phytofurans can be formed from ALA, GLA, and SDA, which possess 2 or more bis-allylic carbons.¹⁰² The first step of lipid peroxidation is the abstraction of a bis-allylic hydrogen, generating a stabilized pentadienyl radical. After radical mesomeric rearrangement, a dioxygen molecule is trapped by the conjugated radical to generate a peroxyl radical. Hydrogen abstraction by the peroxyl radical forms a hydroperoxide that can be reduced into a hydroxy group by glutathione peroxidase or a metal complex (Scheme 49, orange path).^301−304 As opposed to enzymatic oxidation, which generates enantiopure hydroperoxides, the action of free radicals on PUFAs is not stereoselective and generates racemic mixtures.²⁸⁶ Nonenzymatic abstraction of a bis-allylic hydrogen on LA generates 9-HpODE and 13-HpODE,³⁰⁵ and 9-HpOTrE, 12-HpOTrE, 13-HpOTrE, and 16-HpOTrE are formed from ALA.³⁰⁶ Even if not yet reported in the literature, the same reactions can occur with other C18-PUFAs including GLA, forming 6-, 9-,10-, and 13-HpOTrE-γ, and SDA, forming 6-,9-,10-,12-, 13-, and 16-HpOTE. Similarly, the abstraction of a hydrogen on carbons 8 and 11 of OA generate 8-HpODA, 9-HpODA, 10-HpODA, and 11-HpODA.^177,307

Scheme 49 — The orange color indicates the formation of linear hydroperoxide as a side reaction.

Nonenzymatic oxidation can also be initiated by nonradical ROS species such as singlet oxygen. Singlet oxygen (¹O₂) is an excited state of dioxygen that can be generated by photoactivation. Singlet oxygen is a highly electrophilic species and can easily react with alkene moieties to produce hydroperoxides via an ene-reaction (Scheme 32).^308,309 PUFAs can be oxidized by singlet oxygen, resulting in the formation of hydroperoxides in a process called photosensitized oxidation. Reaction of singlet oxygen with LA in mice skin creates four hydroperoxides, (9-HpODE, 10-HpODE, 12-HpODE, and 13-HpODE), among which 10-HpODE and 12-HpODE are specific products of singlet oxygen oxidation.³¹⁰ Photosensitized oxidation of ALA generates 9-HpOTrE, 10-HpOTrE, 12-HpOTrE, 13-HpOTrE, 15-HpOTrE, and 16-HpOTrE, among which terminal hydroperoxides (9-HpOTrE and 16-HpOTrE) are favored. 10-HOTrE and 15-HOTrE are not generated by autoxidation of ALA.³¹¹ OA, which does not possess any bis-allylic hydrogen, is less sensitive to autoxidation but can undergo photosensitized oxidation by singlet oxygen to produce 9-HpOME and 10-HpOME.³¹⁰

Hydroperoxides can be transformed by the same enzymatic systems responsible for the production of octadecanoid oxylipins, as previously described, but can also undergo specific nonenzymatic oxidation. This results in a high diversity of octadecanoids, possessing both linear and cyclic structures. Among the former, mono-, di-, and trihydroxy groups, ketones, hydroxy-ketones, mono-, and diepoxides, keto-epoxides and hydroxy-epoxides have been described.^{301−304,306}

Reactive nitrogen species (RNS), such as nitrogen dioxide radical (^•NO₂) created from nitric oxide radical (^•NO) in the presence of oxygen, or peroxynitrite (ONOO^–) created from ^•NO in the presence of superoxide radical, are also able to react with mono-FAs and PUFAs to produce nitro-FAs.³¹² While peroxynitrite can initiate lipid peroxidation reactions,³¹²^•NO can terminate such reactions,³¹³ and ^•NO₂ can react directly with conjugated fatty acids to produce various nitro fatty acids including NODEs and NOTrEs.³¹⁴ Notably, multiple molybdopterin-based nitrate reductases have been identified including xanthine oxidoreductase (XOR), aldehyde oxidase (AO) and sulfite oxidase (SO), which are capable of reducing NO₂^– to ^•NO in low pH anoxic microenvironments as found in hypoxic inflammatory sites.³¹⁵ In addition, hemoglobin has also been reported to execute the NO₂^– to ^•NO conversion under hypoxic conditions.³¹⁶ Therefore, it is important to realize that while the classic arginine-dependent production of ^•NO is dominant when oxygen tensions are high, when oxygen tension is low alternate NO₂^–-dependent mechanisms are available and may contribute to nitro-FA production.

6.1. Linear Metabolites

6.1.1. Alcohols

Hydroperoxides can be reduced into hydroxy groups in two steps in the presence of a metal. First, an alkoxyl radical is formed by reduction of hydroperoxides by a variety of free metals or metal complexes (e.g., ferrous chloride) and metalloproteins (e.g., hemoglobin and myoglobin).^298,300 The alkoxyl radical can then abstract a hydrogen to generate a hydroxylated FA.³¹⁷ Alternatively, an electron transfer from Fe(II) can directly form an alkoxyl anion which generates a hydroxy group after protonation (Scheme 33).³⁰⁰

The primary autoxidation (and enzymatic) products of LA (9- and 13-HpODE) were originally characterized by Nobelist Sune Bergstrom and Ralph Holman in the 1940s.^318,319 In detailed extension of this work, the formation of 10(E),12(Z/E)-9-HODE and 9(Z/E),11(E)-13-HODE by autoxidation of LA and formation of 8-HOME, 9-HOME, 10-HOME, and 11-HOME from OA was reported by Gardner et al. in 1974.¹⁷⁷ Then, in 1977, Frankel described the generation of 9-HOTrE, 12-HOTrE, 13-HOTrE, and 16-HOTrE by autoxidation of ALA followed by NaBH₄ reduction of the hydroperoxides.³⁰⁶

6.1.2. Ketones/Hydroxy-ketones

Ketones are readily available from PUFA hydroperoxides via transformations of peroxyl or alkoxyl radicals. The classic “Russell mechanism” involves the combination of two peroxyl radicals followed by the elimination of singlet oxygen and formation of one alcohol and one ketone.³²⁰Scheme 34 illustrates the reactions of peroxyl radicals from 13-HpODE, combining to form a linear tetroxide that decomposes to give 13-HODE, 13-oxo-ODE, and singlet molecular oxygen. The predictions of singlet oxygen formation have been tested over the years and although early experiments fell short in the quantitative yield of ¹O₂,³²¹ later experiments have garnered more support for the Russell mechanism.^322,323

An interesting twist on the diradical combination is a proposed fusion of peroxyl and alkoxyl radicals to yield a ketone and regenerate a hydroperoxide (Scheme 35).³²⁴ The reactions occurred in the micro environment of lipid (linoleate) micelles, potentially bringing together these fleeting radical species. Radical–radical dismutation between a (E/Z)-linoleate alkoxyl radical and (E/E)-linoleate peroxyl radical explained the production of 9- and 13-(E/Z)-oxo-ODEs and (E/E)-HpODEs.³²⁴

Alkoxyl radicals are generated by metal-dependent reduction of hydroperoxides and their further transformations lead to complex mixtures prominently featuring ketones and alcohols (Scheme 36). In 1974, Gardner et al. demonstrated the formation of 9(Z/E),11(E)-13-oxo-ODE, 10(E),12(E/Z)-9-oxo-ODE, 10(E)-9-OH-13-oxo-OME, and 11(E)-13-OH-9-oxo-OME from LA in the presence of cysteine and a catalytic amount of Fe (III), or equimolar amount of Fe(II), in ethanol, at room temperature.¹⁷⁷ Zhu et al. reported that autoxidation of LA enhanced by Fe(II)/ascorbate under physiological condition (37 °C, pH 7.4) generates three main metabolites: a hydroxy-ketone possessing a trans-unsaturation (11(E)-10-OH-13-oxo-OME), a dihydroxy-ketone (11(E)-9,10-DiOH-13-oxo-OME), and an all-trans ketone (9(E),11(E)-13-oxo-ODE).³²⁵

Marnett and co-workers studied these mechanisms and the ensuing products by hematin-catalyzed degradation of oleic, linoleic and linolenic hydroperoxides.^298,326,327 They noted that “When a double bond is β to the (alkoxyl) radical, the principal fate is loss of the α-H to form a ketone.”^298,326,327 The transformations of 13-HpODE gave multiple epoxy derivatives plus 13-HODE and 13-oxo-ODE.³²⁷ Nonenzymatic oxidation of 13-HpOTrE into a small amount (7%) of 13-oxo-OTrE at room temperature in dichloromethane and in the presence of 5,10,15,20-tetraphenyl-21H,23H-porphyrin iron(III) chloride (TPP/Fe(III)) and 2,4,6-tri-tert-butylphenol, which generate an alkoxyl anion from the hydroperoxide, has been reported by Wilcox et al.,²⁹⁸ whereas the formation of a small amount (1.2%) of all-trans ketones (10(E),12(E)-9-oxo-ODE and 9(E),11(E)-13-oxo-ODE) by photosensitized oxidation of LA-derived 9- and 13-HpODEs has been described by Neff in the presence of methylene blue at 0 °C in dichloromethane (Scheme 37).³²⁸

Scheme 37 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

6.1.3. Diols

In 1983, Hamberg reported on the hemoglobin-catalyzed transformation of 13(S)-HpODE to a leukotriene A-type epoxide, 8(E),10(E)-12(13(S))-EpODE, with its rapid nonenzymic hydrolysis on the 8-carbon to two 8,13-dihydroxy diastereomers (Scheme 38).³²⁹ In analyzing the autoxidation of LA, Hamberg followed up on these findings by identifying four dihydroxy derivatives with conjugated dienes (two 8,13-diols and two 9,14-diols), postulated as the products from 9-HpODE and 13-HpODE.³³⁰ Subsequently, an equivalent enzymic transformation of 9(R)-HpODE to an allylic 9(10)-epoxide and its hydrolysis to 9,14-diols was shown to be catalyzed by an Anabaena catalase-LOX fusion protein.³³¹

Scheme 38 — Nomenclature is as described in Scheme 3.

In 1982, Frankel et al. reported the formation of small amount of 9,10-DiHODE, 9,12-DiHODE, 9,13-DiHODE, 10,12-DiHODE, 10,13-DiHODE, and 12,13-DiHODE (0.3–5.3 wt % depending on peroxide value) as secondary products of photosensitized oxidation of LA’s 9- and 13-HpODEs at 0 °C in dichloromethane and in the presence of methylene blue.³³² The formation of 9,14-DiHpODE from 9-HpODE and 8,13-DiHpODE from 13-HpODE in the presence of singlet oxygen and methylene blue at 0 °C in dichloromethane has also been observed by the same group in 1983, and represent 17.0% of total products.³²⁸

In 1982, Neff et al. described the photosensitized oxidation of ALA into a mixture of different dihydroperoxides: 10(E),13(E),15(Z)-9,12-DiHpOTrE, 9(Z),11(E),14(E)-13,16-DiHpOTrE, 8(E),13(E),15(Z)-10,12-DiHpOTrE, 9(Z),11(E),16(E)-13,15-DiHpOTrE, 8(E),12(Z),14(E)-10,16-DiHpOTrE, 10(E),12(Z),16(E)-9,15-DiHpOTrE, 10(E),12(Z),14(E)-9,16-DiHpOTrE, and 10(E),12(E),14(E)-9,16-DiHpOTrE. These dihydroperoxides have been identified as major secondary products of ALA oxidation in the presence of singlet oxygen.³³³ The 9,12-, 13,16-, and 9,16-dihydroperoxides may be formed from the 9- and 16-hydroperoxides by the same mechanism suggested for autoxidized methyl linoleate that proceeds through pentadienyl radicals. The other dihydroperoxide products (1,3-disubstituted and 1,7-disubstituted) are created by concerted addition of singlet oxygen on monohydroperoxides. An overview of the described diols formed from LA and ALA is provided in Scheme 39.

Scheme 39 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

6.1.4. Epoxides/Hydroxy-epoxides/Triols

In 1973, Hamberg et al. first reported the autoxidation of 13-HpODE into 9(Z)-11-OH-12(13)-trans-EpOME by homolysis of the hydroperoxide, formation of an epoxide, and attack by a water molecule.³³⁴ More recently, these hydroxy-epoxide compounds have been shown to be generated from an alkoxyl radical, which tends to rearrange into more stable mesomeric epoxy-allylic radicals, even in the presence of compounds with a readily abstractable hydrogen.^335,336 These radicals can perform hydrogen abstraction to form epoxides or, alternatively, they can also be scavenged by oxygen to form hydroperoxy-epoxides after hydrogen abstraction.³³⁷ In 1981, Gardner et al. described formation of 9(10)-EpOME and 12(13)-EpOME following this mechanism, in the presence of FeCl₃/Cys to generate the alkoxyl radical. The formation of 10(E)-9-Hp-12(13)-trans-EpOME from 13(S)-HpODE in the presence of Fe(III)/cysteine has also been reported.³³⁸ Cleavage of the hydroperoxide followed by hydrogen abstraction generates hydroxy-trans-epoxide compounds that can be hydrolyzed into triols (Scheme 40).³³⁵

Scheme 40 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

In the presence of Fe(III) and cysteine, Gardner et al. showed that oxidation of LA results in the formation of 9(Z)-11-OH-12(13)-trans-EpOME, 12(Z)-11-OH-9(10)-trans-EpOME, 9(E)-11-OH-12(13)-trans-EpOME, and 12(E)-11-OH-9(10)-trans-EpOME as well as the triols 10(E)-9,12,13-TriHOME and 11(E)-9,10,13-TriHOME.¹⁷⁷ In 1978, the formation of 10(E)-9-Hp-12(13)-trans-EpOME from 13-HODE was reported by the same group.³³⁷ Then, in 1981, they showed that these products are generated from monohydroperoxides. Indeed, 11(E),15(Z)-9-OH-12(13)-trans-EpODE can be generated by decomposition of 13-HpOTrE. Decomposition of 13-HpODE or 9-HpODE in the presence of FeCl₃/Cys resulted in formation of 9(Z)-erythro/threo-11-OH-12(13)-trans-EpOME, and 12(Z)-erythro/threo-11-OH-9(10)-trans-EpOME as well as lesser amount (about 15–25%) of the (E)-isomer (Scheme 41).³³⁵

Scheme 41 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known. The dotted arrows refer to the formation of compounds for which a primary citation could not be identified in the literature.

6.1.5. Keto-epoxides

Keto-epoxides are mainly formed from hydroperoxy-epoxides via alkoxyl radical formation. The first step of the proposed mechanism is the formation of an alkoxyl radical by a metal-dependent reduction of the hydroperoxide, followed by a rearrangement which gives a hydroxy group possessing a radical in the α-position. This radical electron is trapped by a ferric ion to form a cation which, after deprotonation of the alcohol, forms a ketone. (Scheme 42).^{177,300,335−337}

Another reported mechanism describes the formation of the ketone via cleavage of the hydroperoxide into an alkoxyl radical that can degrade into a ketone.³³⁵ In the presence of FeCl₃/Cys, Gardner et al. showed the formation of 11(E)-13-oxo-9(10)-trans-EpOME, 11(E)-13-oxo-9(10)-cis-EpOME, 10(E)-9-oxo-12(13)-trans-EpOME, and 10(E)-9-oxo-12(13)-cis-EpOME by autoxidation of LA.¹⁷⁷ In 1981, the same group showed that, in the presence of FeCl₃/Cys, 10(E)-9-oxo-12(13)-trans-EpOME, or 11(E)-13-oxo-9(10)-trans-EpOME are formed from 13-HpODE and 9-HpODE, respectively, in ∼18% overall yield. The cis-epoxides are produced as well but in lower yield (5% overall).³³⁵ The formation of these four keto-epoxyoctadecenoic acids possessing a trans-double bond was shown by Lin et al., simultaneously with two new keto-epoxy-metabolites by autoxidation of LA promoted by Fe(II)/ascorbic acid (Scheme 43).³³⁹

Scheme 43 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known.

6.1.6. Nitro-FAs

When the enzymatic production of superoxide (e.g., NADPH oxidases) and ^•NO occur in close proximity, these species can combine to form peroxynitrite, which can further react with carbon dioxide to yield ^•NO₂.³⁴⁰ Under normoxic conditions, nitric oxide synthase is the primary source of ^•NO; however, under hypoxic conditions, nitrite is reduced to ^•NO by deoxygenated myoglobin, xanthine oxidoreductase, and other systems.³⁴¹ Nitrated fatty acids (NO₂-FA) are the best-known products of RNS and are endogenous signaling molecules. They are formed by the attack of nitrogen dioxide radical (^•NO₂) or nitronium cation (NO₂⁺) on unsaturated FAs, leading to the introduction of a nitro group on any position of the double bonds.^342,343 The mechanism of formation of the NO₂-FAs is still not known with certainty, because several biosynthetic routes can form these metabolites.

The most commonly reported NO₂-FAs identified in vivo are the nitroalkenes, such as nitro-oleic acids (NOMEs, 2 regioisomers, Scheme 44), nitro-linoleic acids ((Z,Z)-NODEs, 4 regioisomers, Scheme 44), and nitro-conjugated linoleic acids ((E,Z)-NODEs, 2 regioisomers, Scheme 44).^344,345 Notably, conjugated linoleic acid appears to be the preferred substrate for these reactions¹⁶⁰ and mechanisms associated with ^•NO₂ production are increased during inflammation and metabolic stress with both 9(E),11(E)-9-NODE and 9(E),11(E)-12-NODE being produced.³⁴⁶

Nitroalkenes can also be precursors of other nitrated species because they can undergo additional reactions with ROS and RNS to be further nitrated, and form nitroso, dinitroso, nitronitroso, di- and trinitro species. They can also be oxidized to generate nitrohydroxy, nitrohydroperoxy, nitro-epoxy, or nitro-keto.^160,347

Two mechanisms coexist for the synthesis of the nitroalkene metabolites. One of them is a radical mechanism that begins by a radical-induced nitration that generates a β-nitroalkyl radical. The abstraction of a hydrogen radical gives a nitro-allyl derivative. If the β-nitroalkyl radical undergoes a second addition of a ^•NO₂, a nitronitrite compound is obtained, and can be hydrolyzed into a nitrohydroxy fatty acid. It can also lose nitrous acid to give a nitroalkene.³⁴⁶ Formation of nitro allyl derivatives can also proceed through bis-allylic hydrogen abstraction, and reaction of the resulting stabilized pentadienyl radical with nitrogen dioxide radical (^•NO₂) (Scheme 45, orange path).³⁴⁶

Scheme 45 — The orange path indicates formation of nitro allyl derivatives through bis-allylic hydrogen abstraction.

Similarly, the formation of NO₂-CLA starts by radical-induced nitration that generates an allylic radical. This radical can isomerize via resonance and reacts with a molecule of ^•NO₂ or ^•O₂ in the gamma position of the nitro group to form a nitronitrite intermediate or a nitro-peroxide intermediate. The elimination of the nitrite gives the nitroalkene, while the reduction of the peroxide into an alkoxyl radical gives a nitro-hydroxy metabolite by reduction of a nitro-oxo metabolite by oxidation (Scheme 46).

The second mechanism for the formation of nitroalkene consists of an electrophilic substitution of a nitronium cation on a double bond. The nitronium cation can be formed by the reaction of a transition metal with peroxynitrite (Scheme 47).

6.2. Cyclic Metabolites

In addition to linear metabolites, three families of cyclic metabolites can be generated nonenzymatically from C18-PUFAs: endoperoxides and diendoperoxides, phytoprostanes (PhytoPs), and phytofurans (PhytoFs). These compounds are named in accordance with the eicosanoid classes isoprostanes and isofurans. Cyclic epoxides are discussed in the linear metabolites section. Endoperoxides can be generated from all PUFAs, while PhytoPs and PhytoFs are only generated from C18-PUFAs possessing at least three unsaturations (ALA, GLA, SDA). Differently from C20-PUFAs, for which cyclic metabolites can be also produced enzymatically by COX action, C18-PUFAs only produce cyclic metabolites via nonenzymatic oxidation. The resulting compounds contain 5-membered rings and can be created both via radical and singlet oxygen-dependent processes including a host of biologically mediated processes.³⁴⁸ Production of cyclic nonenzymatic metabolites of PUFAs, including octadecanoids, has been extensively studied and reviewed by Durand and colleagues.^{102,103,349,350}

6.2.1. Endoperoxides

The ¹⁸O₂-labeling evidence by Samuelsson for an endoperoxide intermediate in prostaglandin biosynthesis³⁵¹ and the subsequent isolation of PGG₂ and PGH₂³⁵² sparked great interest in the occurrence of fatty acid endoperoxides and the mechanisms of their formation. Subsequent mechanistic studies were conducted by the Porter and Pryor laboratories using C18:3 substrates, undoubtedly due to the ready availability of these plant-derived fatty acids in the 1970s. Porter and Funk formed prostaglandin-like bicyclic endoperoxides by free radical chemistry, thus supporting the involvement of this mechanism in prostaglandin biosynthesis.^353,354 The focus of the Pryor group was on implicating bicyclic endoperoxides as reactants in the widely used thiobarbituric acid method to detect and assay fatty acid oxidants in autoxidation.³⁵⁵ Interesting enzymological production of monocyclic endoperoxides was reported by Roza and Francke in their studies on the soybean lipoxygenase-catalyzed transformations of α-linolenate methyl ester.³⁵⁶ Using soybean flour and neutral pH conditions, the main products were the hydroxy-endoperoxides 16-hydroxyperoxy-13,15-endoperoxy-linolenate and 9-hydroxyperoxy-10,12-endoperoxy-linolenate. A chemical point of interest is the resistance of these monocyclic endoperoxide moieties to reduction by sodium borohydride; instead, for analytical purposes, the endoperoxides were opened to diols by hydrogenation using palladium on a carbon catalyst.³⁵⁶

Octadecanoid endoperoxides can be created via both radical and singlet oxygen-dependent processes. In 1980, the Chan and Mihelich groups reported for the first time the formation of 5-membered ring endoperoxide hydroperoxides by both photosensitized oxidation and autoxidation of LA.^357,358 In 1981, Neff et al. separated and identified 13(E),15(Z)-10(12)-epidioxy-9-HpODE, 9(E),11(E)-13(15)-epidioxy-16-HpODE, and 9(Z),11(E)-13(15)-epidioxy-16-HpODE as major secondary products of autoxidized ALA.³⁵⁹ The same year, the group published a study concerning the photosensitized oxidation of LA and separated and identified the structures of the endoperoxides previously discovered by Mihelich. These products are diastereoisomeric pairs of 8(E)-10(12)-epidioxy-13-HpOME and 13(E)-10(12)-epidioxy-9-HpOME and are generated from specific singlet oxygen hydroperoxides (10-HpODE and 12-HpODE).³³²

The following year, the Neff and Chan groups showed that 6 endoperoxides already identified among the autoxidation products of ALA^359,360 could be also produced by photosensitized oxidation and described the formation mechanisms.³³³ These products (13(E),15(Z)-10(R)(12(S))-epidioxy-9(R/S)-HpODE and 13(E),15(Z)-13(S)(15(R))-epidioxy-16(R/S)-HpODE and their enantiomers) are derived from 12-HpOTrE and 13-HpOTrE and represent 97% of the endoperoxides generated by photosensitized oxidation of ALA. The remaining 3% consisted of 8(E),15(Z)-10(12)-epidioxy-13-HpODE and 9(Z),16(E)-13(15)-epidioxy-12-HpODE, generated by cyclization of 10- and 15-HpOTrE. These endoperoxides tend to cyclize again to form the diendoperoxides 16(E)-10(S)(12(R));13(S)(15(R))-diepidioxy-9(R/S)-HpOME and 8(E)-10(R)(12(S));13(R)(15(S))-diepidioxy-16(R/S)-HpOME as well as their enantiomers, which were reported for the first time by Neff et al. in 1982.³³³ Therefore, 8(E),15(Z)-10(12)-epidioxy-13-HpODE and 9(Z),16(E)-13(15)-epidioxy-12-HpODE, as well as diendoperoxide metabolites, are specific products of singlet oxygen-dependent oxidation of ALA and are not formed by radical mediated autoxidation.

In 1983, Neff et al. identified the 6-membered ring endoperoxides 10(Z)-9(12)-epidioxy-13-HpOME and 11(Z)-10(13)-epidioxy-9-HpOME as major products (57.9%) of the reaction of 9- and 13-HpODE with singlet oxygen.³²⁸ Six-membered hydroperoxy-epidioxides are formed by 1,4-addition of singlet oxygen to the 1,3-diene system of linoleate hydroperoxides (Scheme 48).³⁶¹ The formation of 6-membered ring endoperoxides has been hypothesized by Frankel et al. to explain the formation of 9,10,12-TriHODE and 13,15,16-TriHODE from 12-HpOTrE and 13-HpOTrE, respectively.³⁰⁶

Scheme 48 — Nomenclature is as described in Scheme 3. Synthetic route is referenced if known. ^aThe enantiomer is also generated.

6.2.2. Phytoprostanes

PhytoPs can be produced from all C18-PUFAs possessing at least three unsaturations (e.g., ALA, GLA, and SDA). PhytoPs are abundant in plant-based food¹⁴⁴ but can also be synthesized in humans by direct nonenzymatic oxidation of these PUFAs. In 2009, a study published by Barden et al. showed that supplementation with flaxseed oil, an oil rich in ALA, leads to significant increases in F₁-PhytoP in human plasma and urine.³⁶² The biosynthesis mechanism of PhytoPs has not been determined directly but it has been inferred by analogy with isoprostanes (IsoPs).¹⁰² It initiates by the formation of a peroxyl by abstraction of a hydrogen radical on a bis-allylic position, followed by a radical rearrangement and trapping of a dioxygen molecule. The next step is a 5-exo-trig cyclization of the peroxyl that generates an endoperoxide.^363,364 A second 5-exo-trig cyclization forms a bicyclic endoperoxide and creates mostly a cis-configuration between the two lateral chains.^365,366 A second molecule of dioxygen is trapped to form a hydroperoxide that can be subsequently reduced together with the endoperoxide into hydroxy groups via the action of antioxidant species (e.g., glutathione or α-tocopherol), generating mostly type F-PhytoP.^367,368 The hydroxy moieties on the ring can be in cis-configuration with the two lateral chains, creating the cis-PhytoPs (e.g., F_1c-PhytoPs) or in a trans-configuration with the lipidic chains, creating trans-PhytoP (e.g., F_1t-PhytoPs). In total, 8 isomers of PhytoPs can be generated from ALA. When the concentration of reducing agent is lower and H-PhytoP is present in an aqueous environment, type D- and E-PhytoPs are mostly formed via the rearrangement of H₁-PhytoP.^352,369,370 Dehydration of type D- and E-PhytoPs occurs easily under physiological conditions and creates type J- and A-PhytoPs analogous to the isoprostanes.³⁷¹ Finally, type B- and L-PhytoP are obtained by isomerization of type A- and J- PhytoPs’ intracyclic double bond (Scheme 49).³⁷²

As ALA possesses two bis-allylic positions, two different families of PhytoPs can be formed, namely 9-PhytoPs (if the abstraction takes place on the 11-position) and 16-PhytoPs (if the abstraction takes place on the 14-position) (Scheme 50). GLA also possesses two bis-allylic positions, and the abstraction of a hydrogen radical on the 8-position generates the 6-GLA-PhytoPs, whereas the abstraction of a hydrogen radical on the 11-position forms 13-GLA-PhytoPs (Scheme 51). SDA possesses three bis-allylic positions, and four families of SDA-PhytoPs can be created. A bis-allylic hydrogen radical can be abstracted on the 8-, 11-, and 14-positions, generating 6-SDA-PhytoPs, 9-SDA-PhytoPs, 13-SDA-PhytoPs, and 16-SDA-PhytoPs respectively (Scheme 51). One study on GLA-derived phytoprostanes was reported by Porter et al. in 1975. In this study, it was shown that 13-F₁-GLA-PhytoP is generated from LOX-generated hydroperoxide 9-HpODE in the presence of a free radical source in O₂-saturated benzene.³⁵³ To the best of our knowledge, the formation of cyclic metabolites from SDA has not yet been reported.

Scheme 51 — The α-chain is displayed in blue color and the ω-chain in green color.

6.2.3. Phytofurans

Under increased oxygen tension, such as in highly oxygenated tissues like lung, brain, and kidneys, cyclic nonenzymatic metabolites possessing a substituted tetrahydrofuran ring (PhytoFs) are preferentially formed compared to phytoprostanes.^373,374 PhytoFs exist in two distinct families, termed alkenyl PhytoFs and enediol PhytoFs. In 2002, Fessel et al. were the first to propose two distinct mechanisms for the formation of the two families of PhytoFs.³⁷³ A unified route for the formation of both families was then proposed by Jahn et al. in 2008,¹⁰² based on the known formation of a bis-epoxide.¹³ The first step of phytofuran biosynthesis is identical to isoprostane biosynthesis and a cyclic endoperoxide is created by addition of a dioxygen molecule followed by a 5-exo-trig cyclization. After generation of the cyclic endoperoxide, a 1,3-S_Hi (intramolecular homolytic substitution) reaction, a 3-exo cyclization generates a radical diepoxide that traps a dioxygen molecule to form a diepoxyhydroperoxide after protonation. After hydrolysis, two regioisomeric epoxy diols are created. An intramolecular nucleophilic ring opening of the epoxide by a hydroxy group generates the phytofurans (Scheme 52). Analogously to PhytoPs, the biosynthesis mechanism of PhytoFs has not been determined directly but it has been inferred by comparison with isofurans (IsoFs).³⁵⁰

Scheme 52 — Color indicates the site of attack for a given chemical moiety and the resulting metabolite.

7. Physiological Distribution and Compartmentalization

As with their parent fatty acids, oxygenated octadecanoids are incorporated into various biomolecular complexes through esterification with the free alcohols of glycerol and cholesterol, and their cellular compartmentalization, physiological distribution, and trafficking behavior can be best understood in this context.^375−377 Similarly, NO₂-FAs are stable in both esterified and nonesterified forms, are transported in lipoprotein particles³⁷⁸ and accumulate in peripheral tissues.³⁷⁹ While it has long been recognized that complex lipids can contain functionalized lipid species, the tools for their efficient investigation are still in development, and the importance of the esterified fraction is just beginning to be broadly considered and addressed.^378,380,381 The majority of oxylipin investigations still focus on the free acid; however, there is an increasing appreciation that the esterified fraction is often the larger oxylipin pool.³⁸⁰ While the biosynthesis of esterified oxylipins has been reviewed in plants,³⁸² less is known about these pathways in mammals. Free oxylipins can be produced from free fatty acids and then subsequently be esterified into complex lipids. Alternatively, fatty acids can be esterified into complex lipids and then directly transformed into oxylipins.^383,384 For example, it has been shown that both human 15-LOX-2 and mouse 8-LOX can transform phospholipid esterified AA to 15-HETE.³⁸⁵ It has long been known that cholesteryl linoleate esters of LDL are a substrate for 15-LOX, leading to formation of 13(S)-HODE by recombinant 15-LOX⁴⁶ or rabbit 15-LOX.³⁸⁶ Accordingly, future investigations in the oxylipin field should further characterize the formation and function of esterified octadecanoids.

While the nonesterified octadecanoids are thought to be bioactive fractions, esterified forms can act as lipid mediator reservoirs and may play a role in the activity and function of lipid microdomains. For instance, at the cellular level, the Lands cycle describes the fundamental biochemical process of phospholipid membrane remodeling.³⁸⁷ Both epoxides and alcohols of AA participate in this process.^377,388,389 Notably long-chain acyl-CoA synthetase (ACSL)4 and ASCL1 prefer AA at similar rates, but epoxy and hydroxy arachidonates differentially.³⁸⁹ By analogy, the octadecanoids would also be expected to be active participants in this biochemical network, however little direct evidence exists. Interestingly, ASCL1 is broadly distributed and has a distinct preference for oleate and linoleate,³⁹⁰ suggesting that it may be a candidate to regulate the intercellular phospholipid distribution of some octadecanoids. We have previously reported that adipose tissue harbors a significant pool of esterified eicosanoids and octadecanoids, whose composition and relative distribution between free and esterified pool are influenced by dietary fat content.³⁹¹ The postprandial decline and late rebound of nonesterified HODEs, EpOMEs, DiHOMEs also suggest that this pool is substantially influenced by release from adipose triglycerides.^392,393 Similar results were reported with insulin infusions in early and late fasting elephant seals, a unique animal model of insulin resistance.³⁹⁴ In unpublished works in the Syrian hamster (J.W. Newman, personal communication), octadecanoids were detected in cholesteryl esters, phospholipids and triglycerides of adipose, liver and muscle tissues. Notably, adipose and muscle tissue triglycerides were particularly enriched in HODEs, HOTrEs, EpOMEs, and EpODEs, and the relative balance of these LA and ALA metabolites were responsive to the dietary lipid balance. In supplemented adipocytes, the NODA, NOME, (E,Z)-NOMEs as well as their chain shortened dihydro- and tetranor- metabolites were also observed in mono-, di- and triacylglycerol pools as well as various phospholipid classes.³⁷⁹

Considering whole body octadecanoid trafficking, recognition of these bioactive agents as integral parts of lipoprotein particles raises a number of intriguing possibilities. For instance, receptor mediated delivery with lipase-mediated release of bioactive agents may represent an endocrine signaling avenue for such compounds.³⁷⁶ Within lipoprotein fractions, octadecanoid contributions have been reported to range from 75 to 90% of the totals, with HODEs, oxo-ODEs > EpOMEs ≫ DiHOMEs and vary by particle density class.^376,395 Since upward of 90% of oxidized fatty acids in circulation are found in the esterified pools,^375,396 these lipoprotein bioactive payloads will provide a steady pulsatile signal to the periphery, that will covary with that of other nutrients. Therefore, it will likely be necessary to fully speciate lipoprotein particle classes and subclasses to fully understand the passive and/or active dynamics associated with octadecanoid trafficking in the intact organism.

Another issue for consideration in investigating octadecanoids is the use of serum vs plasma. It has been extensively shown that the levels of oxylipins can vary with the blood preparation examined and that this must be taken into account during experimental preparation. This is primarily due to platelet activation during the blood collection process and is a known issue of concern with some eicosanoids (e.g., TxB₂, 12-HETE).³⁹⁷ For many of the octadecanoids, there are relatively few differences in the choice of blood preparation protocol and there is not a major need to consider this issue (Table 3).

Table 3. Comparison of Octadecanoid Levels in Different Human Blood Preparations^a.

parent fatty acid	octadecanoid	serum^b	serum^c	serum^d	plasma EDTA^b	plasma EDTA^c	plasma EDTA^d	plasma heparin^b	plasma citrate^b
linoleic acid	9(10)-EpOME	2.7	0.4	0.8	1.6	0.3	0.8	1.8	1.7
	12(13)-EpOME	10	4	2.0	7.0	3	2.0	6.4	6.5
	9,10-DiHOME	14	5	2.9	11.9	4	3.0	10.4	10.7
	12,13-DiHOME	3.7	6	3.1	2.9	5	3.0	2.7	2.6
	9-HODE	15	14	7.0	9.3	12	7.1	9.9	8.4
	13-HODE	37	21	13.6	29.3	17	14.4	27.2	25.2
	9-KODE	2.6	2	2.3	1.9	<1	3.3	1.9	1.9
	13-KODE	0.8	<1	NR	0.6	2	NR	0.6	0.6
	10(Z)-9-oxo-12(13)-EpOME	4.0	NR^e	1.1	2.3	NR	1.7	1.7	5.3
	9,10,13-TriHOME	2.3	NR	NR	1.3	NR	NR	1.2	3.0
	9,12,13-TriHOME	9.2	4	NR	8.3	5	NR	6.7	8.3

alpha linolenic acid	9(10)-EpODE	0.1	0.2	0.1	0.1	0.2	0.1	0.1	0.1
	12(13)-EpODE	0.2	0.2	NR	0.1	<4	NR	0.1	0.1
	15(16)-EpODE	15	3	2.0	10.1	2	2.0	10.8	8.9
	9,10-DiHODE	0.4	0.3	0.3	0.4	0.2	0.3	0.3	0.3
	12,13-DiHODE	0.5	0.3	NR	0.3	0.2	NR	0.4	0.4
	15,16-DiHODE	9	15	10.5	8.5	13	10.3	7.1	7.5
	9-HOTrE	2.3	0.5	0.6	1.6	0.5	0.5	1.6	1.3
	13-HOTrE	1.9	1	1.0	1.5	0.8	1.1	1.4	1.3

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Concentration in nM.

Data are from a single male in which blood was collected and simultaneously prepared with the different anticoagulants. Oxylipins were measured with the analytical method published by Kolmert et al.³⁹⁸

Data from Pedersen et al.³⁹⁹

Data from Rajan et al.⁴⁰⁰

NR = not reported. Compounds were not reported in the original publications.^399,400

8. Preparation of Octadecanoids

In most cases, a given octadecanoid can be prepared in more than one way. The reactions and methods listed below have all been tested in our laboratory and can serve as useful starting points for investigators new to the field of octadecanoids and oxylipins. The compounds mentioned are in most cases derived from LA; however, as a rule, the reactions are also applicable to the other C18-PUFAs.

8.1. Hydroperoxides

The 9(S)-hydroperoxide of 9(S)-HpODE 1 (Scheme 53) is conveniently prepared using the LOX enzyme present in tomato fruit.⁴⁰¹ The sodium salt of the fatty acid is stirred at 23 °C under oxygen atmosphere with a crude homogenate of tomato fruit in 0.1 M potassium phosphate buffer pH 6.0.⁴⁰¹ Purification is performed by silica gel chromatography followed by normal-phase HPLC. The amount of hydroperoxide can be determined gravimetrically or by UV spectroscopy using an ε value of 27 000 at the λ_max (around 235 nm). For discussion of the ε value of H(p)ODEs, see the review published by Gardner.⁴⁰² For preparation of 13(S)-HpODE 2 (Scheme 53), commercially available soybean LOX-1 is the preferred enzyme. The fatty acid (e.g., LA, ALA) is stirred at 0 °C under oxygen atmosphere with the enzyme in 0.1 M sodium borate buffer pH 10.4; detailed conditions are provided in previous publications.^402,403 Purification and quantitation are performed as outlined above.

E. J. Corey in an early paper noted that irrespective of the position oxygenated in the fatty acid chain, lipoxygenases produce hydroperoxides of the (S) absolute configuration.⁴⁰⁴ Since then, lipoxygenases producing (R)-hydroperoxides have been discovered, notably the 12(R)-lipoxygenase in human skin,⁴⁰⁵ and for preparation of (R)-hydroperoxides such enzymes can be utilized. Alternatively, enzymatic resolution of a racemic hydroperoxide solution can be used. As an example, 13(R)-HpODE was prepared by incubation of 13(R,S)-HpODE with corn allene oxide synthase, an enzyme that specifically converts the (S)-enantiomer into product, but does not react with the (R)-enantiomer.¹¹

Racemic fatty acid hydroperoxides can be obtained by autoxidation or by nonradical oxygenation using singlet oxygen. For the preparation of 9(R,S)- and 13(R,S)-hydroperoxides derived from LA by the former method, LA is evaporated in a round-bottom flask to produce a film covering the wall. The flask is kept at 37 °C for 15 h under an atmosphere of oxygen gas.⁴⁰⁶ This will produce a mixture of (E,Z)-9(R,S)-, (E,E)-9(R,S)-, (E,Z)-13(R,S)-, and (E,E)-13(R,S)-HpODEs. If desired, formation of the (E,E)-isomers can be suppressed by adding α-tocopherol.⁴⁰⁷ Isolation of products can be performed by normal-phase HPLC, in which case the order of elution is (E,Z)-13(R,S)-HpODE (first), (E,E)-13(R,S)-HpODE, (E,Z)-9(R,S)-HpODE, and (E,E)-9(R,S)-HpODE. Racemic hydroperoxides of ALA, GLA and SDA can be prepared in the same general way.

Photosensitized oxidation with generation of singlet oxygen provides a general method for producing racemic hydroperoxides. Illumination with visible light of oxygen-bubbled solutions containing methylene blue generates singlet oxygen, which produces two regioisomeric racemic hydroperoxides from each double bond in a PUFA. For example, irradiation (250 W lamp) of an oxygen-purged solution of LA in methanol containing 0.05% methylene blue at 5 °C produces a mixture of 9(R,S)- and 13(R,S)-HpODEs as well as 10(R,S)-HpODE 3 and 12(R,S)-HpODE 4 (Scheme 54).⁴⁰⁸

In the same way, photosensitized oxidation of ALA produces 6 regioisomeric hydroperoxides that can be separated by normal-phase HPLC.⁴⁰⁹

8.2. Hydroxides

NaBH₄ in methanol (0 °C, 30 min) is commonly used to prepare a fatty acid hydroxide from its corresponding hydroperoxide.⁴¹⁰ Milder reduction methods include SnCl₂ in ethanol and triphenylphosphine in diethyl ether or methanol.^203,402 The two first-mentioned methods require extractive isolation of the product, whereas the last method requires elimination of the triphenylphosphine oxide formed using e.g., a silica gel column. The fatty acid hydroxides can be separated into regioisomers and (E,Z)- and (E,E)-isomers using normal-phase HPLC. Various bacteria, notably Lactobacilli, produce hydroxy acids by a nonlipoxygenase route involving hydratase, dehydrogenase and isomerase enzymes.²²³ Examples of such compounds are 12(Z)-10-HOME 5 and 11(E)-10-HOME 6 formed from LA (Scheme 55). The corresponding compounds are produced from ALA. Such octadecanoids can be prepared using Lactobacillus plantarum enzymes²²³ or can be synthesized by chemical methods.⁴¹¹

The complete series of saturated hydroxystearates has been synthesized as described in the classical paper by Bergström et al.⁴¹² Most of these compounds were obtained by reduction of the corresponding oxo-stearate (see below).

8.3. Ketones

Fatty acid ketones (such as 9-oxo-ODE 7 and 13-oxo-ODE 8, Scheme 56) can be prepared from the corresponding hydroxides using the Dess–Martin periodinane in dichloromethane at 0 °C.⁴¹³ Alternatively, fatty acid hydroperoxides can be directly dehydrated into the corresponding ketones by treatment with acetic anhydride–pyridine (v/v 1:1) at 0 °C.⁴¹⁴ Amounts of products can be determined gravimetrically or by UV spectroscopy using ε = 24 000 at the λ_max (around 278 nm).

Microbially generated fatty acid ketones include 12(Z)-10-oxo-OME 9 and 11(E)-10-oxo-OME 10 (Scheme 57) derived from LA as well as the corresponding compounds formed from ALA. Lactobacillus plantarum enzymes can be used to prepare these compounds²²³ or they can be synthesized using chemical methods.⁴¹¹ In the latter case, the conjugated enone 10 is prepared by treatment of 9 with perchloric acid in tetrahydrofuran, and its reduction with NaBH₄ affords the hydroxy derivative 6.

The complete series of 2- to 17-oxo-stearates has been synthesized.⁴¹² Coupling of dialkyl-cadmium derivatives to acid chlorides was employed in many of these syntheses and in other cases β-ketoesters were reacted with ω-haloesters.

8.4. Epoxides

The monoepoxides of LA and other fatty acids can be obtained either by nonselective epoxidation of double bonds^415,416 or be isolated as natural products, in the latter case as defined stereoisomers. As an example of the first-mentioned approach, methyl linoleate in chloroform is treated with 1.1 equiv of peracetic acid and the solution stirred at 23 °C for 2–3 h.⁴¹⁵ In order to optimize the yield of the two monoepoxides (Scheme 58), the reaction progress has to be followed by TLC or GC-MS. Alternatively, m-CPBA is also a broadly used reagent to perform the epoxidation of LA and ALA in a nonselective way.^417−419 Unreacted methyl linoleate and diepoxide can be removed by silica gel chromatography and the monoepoxide fraction subjected to normal-phase HPLC for separation of the (±)-cis-9(10)-epoxy-12(Z)- and (±)-cis-12(13)-epoxy-9(Z)-octadecenoates (9(10)-EpOME 11 and 12(13)-EpOME 12, respectively, of which the latter is the first eluting). Saponification to the free epoxy acids can be performed by treatment with 0.2 M NaOH in 80% methanol at 23 °C for 15 h. The enantiomers of 9(10)-EpOME 11 are resolved on a Chiralpak AD column in the order 9(S),10(R)-isomer eluting before 9(R),10(S)-isomer.⁴²⁰

By applying the methodology outlined above on the linoleate trans-isomers methyl 9(E),12(Z)- and 9(Z),12(E)-octadecadienoates, it is possible to prepare the corresponding trans-epoxides, i.e., (±)-trans-9(10)-epoxy-12(Z)- and (±)-trans-12(13)-epoxy-9(Z)-octadecenoates, (9(10)-trans-EpOME 13, and 12(13)-trans-EpOME 14, respectively (Scheme 59)).

(+)-Coronaric acid (9(R)(10(S))-EpOME 15(421)), (+)-vernolic acid (12(S)(13(R))-EpOME 16(422)), and (−)-vernolic acid (12(R)(13(S))-EpOME⁴²³) are examples of epoxy acids readily available by extraction of certain seeds, e.g., those from Chrysanthemum coronarium, Euphorbia lagascae, and Malope trifida, respectively (Scheme 60).

8.5. Diols

Ring opening of the above-mentioned epoxides by refluxing with acetic acid followed by saponification or methanolysis affords the corresponding diols.⁴²² Perchloric acid in tetrahydrofuran is often used to effect opening of epoxides into diols; however, in the case of the fatty acid epoxides 11–16 such treatment results in the formation of byproducts due to the presence of the homoallylic double bond. Racemic diols having the threo relative configuration are formed from the cis-epoxides, e.g., 17 from 11, whereas trans-epoxides produce erythro diols, e.g., 18 from 14 (Scheme 61).

Interestingly, attack by acetic acid takes place with preference at the epoxide carbon, which is most distal to the double bond (C-9 in the 9(10)-epoxides and C-13 in the 12(13)-epoxides). This means that diols enriched (about 65:35) with one enantiomer can be prepared if enantiopure epoxides are used.⁴²⁴ Another way of producing erythro diols is dihydroxylation of double bond(s) in an unsaturated fatty acid using, e.g., the so-called Upjohn method.⁴²⁵

8.6. Hydroxy-epoxides

Fatty acid hydroperoxides can produce a variety of hydroxy-epoxide and keto-epoxides, chemically as well as enzymatically. For example, the 2,3-hydroxy-epoxides in which the epoxide and hydroxy groups are vicinally located and additionally possess a double bond vicinal to either the hydroxy group or the epoxide (cf. e.g., structures 19 and 21). As an example of the preparation of a hydroxy-epoxide of the former type, the methyl ester of 13(S)-HpODE is treated in dichloromethane at −78 °C with trifluoroacetic anhydride and lutidine for 30 min followed by K₂CO₃ in methanol.⁴²⁶ This produces the methyl esters of 9(Z)-11(S)-OH-12(S)(13(S))-EpOME and 9(Z)-11(R)-OH-12(S)(13(S))-EpOME (19 and 20, respectively, Scheme 62) of which the first-mentioned (erythro) isomer is the predominant one. These isomers can be easily separated by normal-phase HPLC, the erythro form being the first eluting, and then saponified by mild alkali treatment.

2,3-Hydroxy-epoxides of the latter type can also be prepared from fatty acid hydroperoxide methyl esters, in this case by treatment with a hexane solution saturated with vanadium oxyacetylacetonate.⁴²⁷ As an example, from the methyl ester of 13(S)-HpODE is produced a 1:1 mixture of the methyl ester of 9(Z)-13(S)-OH-11(R)(12(R))-EpOME and 9(Z)-13(S)-OH-11(S)(12(S))-EpOME (21 and 22; threo and erythro, respectively, Scheme 63). Separation of these isomers can easily be performed by normal-phase HPLC, and again the erythro form elutes first. The allylic epoxide present in these hydroxy-epoxides renders them quite sensitive to acid (reported half-life times at pH 3, 1–2 min⁴²⁸), and special care has to be taken during the workup following their saponification.

In addition to the chemical methods mentioned above, enzymes such as peroxygenase and other epoxy alcohol synthases/hydroperoxide isomerases can be used to generate hydroxy-epoxides from hydroperoxides. In this case hydroxy-epoxides having the double bond located between the epoxide and hydroxy functionalities are the exclusive or predominant products. For example, upon incubation of 9(S)-HpODE with preparations from beetroot, the hydroxy-epoxide 10(E)-9(S)-OH-12(R)(13(S))-EpOME 23 is formed,⁴²⁹ whereas 9(R)-HpODE generates 10(E)-9(R)-OH-12(R)(13(S))-EpOME 24 (Scheme 64).⁴³⁰

8.7. Trihydroxy Acids

Acid-catalyzed solvolysis of allylic epoxy alcohols of types 21 and 22 mainly takes place by solvent attack at the double bond carbon most remote from the epoxide followed by double bond isomerization and epoxide opening.^427,431 When carried out in aqueous medium, a given hydroxy-epoxide produces a trihydroxy acid formed as a 1:1 mixture of two diastereoisomers. For example, acid treatment of hydroxy-epoxide 21 in water produces the pair 9(S),12(S),13(S)-TriHOME (pinellic acid, 25) and 9(R),12(S),13(S)-TriHOME 26 (Scheme 65).

Normal-phase HPLC can be used for preparative separation of the methyl esters of trihydroxy acid regio- and diastereoisomers, and the 16 possible 9,10,13- and 9,12,13-trihydroxyoctadecenoic acids derived from LA have been resolved by chiral-phase HPLC.⁴³² Stereospecific opening of allylic hydroxy-epoxides catalyzed by epoxide hydrolases is an alternative to the acid-catalyzed reaction shown above. Thus, potato leaves contain an epoxide hydrolase activity which catalyzes stereospecific hydrolysis of the 2,3-hydroxy-epoxide 12(Z)-9(S)-OH-10(S)(11(S))-EpOME 27 into trihydroxy acid 9(S),10(S),11(R)-TriHOME 28(188) (Scheme 66), and a partially purified epoxide hydrolase from oat seeds⁴³³ catalyzes stereospecific hydrolysis of 23 into trihydroxy acid 25 (pinellic acid).

Additional trihydroxy acids of this and related types have been prepared using chemo-enzymatic methods,^430,434 and by total chemical synthesis.⁴³⁵

8.8. Nitro

In 2013, Woodcock et al. reviewed nitro fatty acid synthesis procedures, which were classified into three approaches: (1) nitrogen dioxide radical/nitronium ion reactions, (2) nitroselenation/nitromercuriation reactions, and (3) total syntheses based on Henry nitroaldol reactions for isomeric synthesis. The review also included a general procedure for nitroselenation of OA and total synthesis of 9(E)-9-NOME as well as purification and analysis of nitrated fatty acids.⁴³⁶

(1) Nitrogen Dioxide Radical/Nitronium Ion Reactions. This method, which mimics the biosynthesis of nitro-FAs, is better adapted for the production of endogenous metabolites from a biological matrix in order to study the formation of nitro-derivatives under pathophysiological conditions. It is based on the exposure of rats to nitrogen dioxide (NO₂) in order to generate in situ nitrogen dioxide radical (^•NO₂).^437−439 This approach gives a mixture of products due to the high reactivity and lack of selectivity of the nitrogen dioxide (Scheme 67). Nitronium ions (i.e., NO₂BF₄) can also be used to generate a mixture of nitro-alkene products.⁴⁴⁰

(2) Nitroselenation/Nitromercuriation Reactions. Nitro-alkenes can be obtained by nitroselenation of unsaturated FAs via a two-step method. For example, the nitro-selenyl intermediates 29 and 30 are created by treatment of OA by mercury chloride, phenylselenyl bromide and sodium nitrite at 0 °C. Then, a treatment of the crude reaction by H₂O₂ at 0 °C gives a mixture of 9(E)-9-NOME 32 and 9(E)-10-NOME 31, that can be separated by HPLC (Scheme 68).^345,436

(3) Total Syntheses Based upon Henry Nitroaldol Reactions for Isomeric Synthesis. The utility of the total synthesis of nitro-alkenes is the ability to obtain a specific isomer. The synthesis of nitrated metabolites of OA^441,442 and LA^443,444 has been published and all methods use a Henry nitroaldol reaction. Typically, the free acid is protected as a methyl^441,443,444 or allyl⁴⁴² ester, and a Henry nitroaldol reaction is performed using either DBU⁴⁴² or t-BuOK^441,443 as a base. The generated nitro-hydroxy intermediate is acetylated and the acetoxy group is eliminated using DMAP^441,443 or Na₂CO₃⁴⁴² to yield the nitro-alkene (Scheme 69).

Recently, an elegant one-pot synthesis has been published by Hassan et al.⁴⁴⁵ In this work, they first describe a synthesis that enables obtaining either (E)- or (Z)-nitroalkene with good stereoselectivity (>95:5 for both stereoisomers). This is the first multistep synthesis of a (Z)-nitro-FA reported. First, the free acid of bromoheptanoic acid 40 is protected by a prenyl group, then prenyl nitroheptanoate 42 is generated and the Henry nitroaldol step was performed with 1,1,3,3 tetramethylguanidine (TMG) and nonanal. The generated hydroxy group of 43 was eliminated with Burgess reagent, a mild dehydrating reagent, and both 9(E)-9-NOME 32 and 9(Z)-9-NOME 45 were obtained with good selectivity (>95:5) following separation of the isomers and deprotection (Scheme 70).

The (E)-isomer can be obtained selectively by a one-pot synthesis. First, the Henry nitroaldol reaction between nonanal and prenyl heptanoate 42 was performed with TMG, then trifluoroacetic anhydride (TFAA) and triethylamine were added in situ to eliminate the hydroxy group, and finally boron trifluoride diethyl etherate was added to the mixture to deprotect the acid and obtain 9(E)-9-NOME 32 with an efficient one-pot procedure (Scheme 71).

8.9. Furan Fatty Acids

Octadecanoids incorporating a furan ring exist as ring-methylated as well as nonmethylated compounds and several methods have been described for their preparation by total organic synthesis.^446−448 For easy preparation of the nonmethylated furans 9(12)- and 10(13)-epoxyoctadecadienoic acids, it is possible to use naturally occurring starting materials as exemplified with preparation of the first-mentioned compound 49 from the methyl ester of ricinoleic acid 46. Oxidation with the Dess–Martin periodinane (i) and dihydroxylation (ii) is followed by warming (iii; 80 °C for 30 min) and saponification (iv). Reaction (iii) involved spontaneous attack by the C-9 hydroxy group at the C-12 carbonyl forming an unstable cyclic hemiketal that loses 2 molecules of H₂O (Scheme 72).⁴⁴⁹

The related 10(13)-epoxy-10,12-octadecadienoic acid 52 can be prepared starting from (±)-vernolic acid 50. Acetolysis followed by saponification and methylation (i, ii, iii) affords the threo-12,13-diol 51, which undergoes aerobic cyclodehydrogenation in the presence of PdCl₂ and CuCl₂ (iv; 95 °C for 20 min) to produce the desired product 52 in good yield following saponification (v).³⁶¹ Purifications are performed by RP-HPLC (Scheme 73).

8.10. Isoprostanes

8.10.1. F Series

F-PhytoPs have been synthesized by Durand and colleagues, and all isomers possessing a syn–anti–syn configuration have been obtained, for the two coexisting series (9th and 16th series).^450,451 The syn-anti-syn ester 54 was first prepared in 8 steps from diacetone l-glucose 53.^452,453 The chains of the phytoprostanes were subsequently introduced by Wittig and Horner–Wadsworth–Emmons (HWE) reactions to obtain 9-F_1t-PhytoP 55 and 16-F_1t-PhytoP 56 in 9 steps from the ester 54(Scheme 74).⁴⁵⁰ The enantiomers ent-9-F_1t-PhytoP and ent-16-F_1t-PhytoP were also obtained following the same strategy starting from diacetone d-glucose.⁴⁵¹

Recently, the first synthesis of a prostaglandin version of a phytoprostane has also been performed by Jahn and colleagues, who developed a strategy based on a radical anion oxidative cyclization and copper(I)-mediated alkyl–alkyl coupling.⁴⁵⁴

8.10.2. D/E Series

The 16-epi-16-E_1t-PhytoP 59 and ent-9-D_1t-PhytoP 62 have been synthesized in the laboratories of Durand¹⁰² and Spur.⁴⁵⁵ The synthesis of 16-epi-16-E_1t-PhytoP 59 was initiated with preparation of the furan precursor 57 by Friedel–Crafts acylation of furan using the anhydride of the azelaic acid monoester.⁴⁵⁵ A ZnCl₂-induced rearrangement followed by treatment with chloral, enzymatic resolution and TBS protection yielded the key intermediate 58, on which the second chain of 16-epi-16-E_1t-PhytoP 59 was introduced by a conjugated addition (Scheme 75). The ent-9-D_1t-PhytoP was prepared in 2009 by Durand and colleagues following a procedure developed for the synthesis of 15-D_2t-IsoP and 15-epi-15-E_2t-IsoP.⁴⁵⁶ The ent-9-D_1t-PhyotP 62 was prepared in 25 steps from 1,3-cyclooctadiene⁴⁵⁷ via the functionalized intermediate 61, which was nicely obtained through a regioselective enzymatic acetylation of the corresponding diol^102,458 (T. Durand, personal communication) (Scheme 76).

8.10.3. J/A Series

9-J₁-PhytoP 65 and 9-A₁-PhytoP 66 were prepared from the common sulfone intermediate 64 by the group of Vidari and Zanoni.⁴⁵⁹ This common intermediate was obtained in 4 steps from the enantiopure hydroxymethyl γ-lactone 63. The preparation of the enantiopure starting material was previously described by the same group.^460,461 From this sulfone, both 9-J₁-PhytoP 65 and 9-A₁-PhytoP 66 were obtained in 4 and 5 steps, respectively (Scheme 77). It is interesting to note that 9-A₁-PhytoP 66 was obtained as a methyl ester instead of a free acid, which is rather unstable due to the lability of the hydrogen on the 12th position. This is the only description of the synthesis of A- and J-PhytoP to date.

8.10.4. B/L Series

The four existing B- and L-PhytoPs have been synthesized by several groups. 9-L₁-PhytoPs 69 and 16-B₁-PhytoP 72 and their enantiomers ent-69 and ent-72 were first prepared by Boland and colleagues.⁴⁶² A few years later, these phytoprostanes were also made by several research groups including Mikołajczyk,^463,464 Riera,^465,466 Durand,⁴⁶⁷ and Vidari.⁴⁶⁸ Finally, the synthesis of 16-L₁-PhytoP 79 was described by Vidari and colleagues in 2015.⁴⁶⁸

9-L₁-PhytoP 69, 16-B₁-PhytoP 72 and their enantiomers ent-69 and ent-72 were easily obtained from the conjugated dienones 68 and 71,⁴⁶⁷ or from the vinyl iodides 74 and 75.⁴⁶² These required intermediates were prepared from the commercially available 1,3-cyclopentandione 67(467,462) or from the cyclooctene 70.⁴⁶⁷ The second chains of the phytoprostanes were then introduced either by a metathesis reaction on the conjugated dienones 68 and 71(467) or by a Heck-type alkylation of the vinylic iodides 74 and 75(462) (Scheme 78).

16-L₁-PhytoP 79 was prepared in 10 steps from O-TBS-protected 2-iodo-3-bromocyclopentenol 76 by the group of Zanoni and Vidari. A I/Li exchange on intermediate 76 followed by formylation with dimethylformamide formed the aldehyde 77. Then, the first chain of the phytoprostane was introduced by a HWE reaction using diethyl (2-oxobutyl)phosphonate to yield the intermediate 78. Finally, 7 extra steps were necessary to obtain the 16-L₁-PhytoP 79 from intermediate 78(Scheme 79).⁴⁶⁸

8.11. Phytofurans

Only one strategy for the synthesis of phytofuran has been reported in the literature. Durand and colleagues described the synthesis of ent-16(R,S)-13-epi-ST-Δ¹⁴-9-PhytoF 83 in 2015.⁴⁶⁹ Following the same strategy, ent-16(R,S)-9-epi-ST-Δ¹⁴-10-PhytoF 84 and ent-9(R,S)-12-epi-ST-Δ¹⁰-13-PhytoF 85 were also prepared by the same group in 2017.⁴⁷⁰ The ent-16(R,S)-13-epi-ST-Δ¹⁴-9-PhytoF 83, ent-16(R,S)-9-epi-ST-Δ¹⁴-10-PhytoF 84 and ent-9(R,S)-12-epi-ST-Δ¹⁰-13-PhytoF 85 were made in 20 steps from the commercially available but-2-yne-1,4-diol 80, via a Payne rearrangement of the bis-epoxide intermediate 81, which yielded the tetraol THF intermediate 82(Scheme 80).

9. Measurement of Octadecanoids

A vital component in the study of octadecanoids is the ability to reliably measure these metabolites, either singly for the confirmation of synthesis and purification, or as metabolite suites for biological investigations. There are several decades of literature on the quantification of oxylipins, and an in-depth discussion is beyond the scope of this review. The vast majority of published methods focus on the eicosanoids given their long history of study. For applications in biomedicine, there are few methods designed to specifically target the octadecanoids. Instead, selected octadecanoids have been included in eicosanoid and oxylipin metabolic profiling methods. Accordingly, these general oxylipin methods will be briefly summarized in this section. A consistent challenge experienced in the development of methods for the measurement of oxylipins has been the pM to nM range of endogenous levels. Early efforts employed a range of methods to measure AA metabolites, including radioimmunoassay (RIA), enzyme linked immunoassay (ELISA) and spectrophotometry.⁴⁷¹ RIA and ELISA methods are sensitive (detection limits of low aM to pM, respectively) and relatively simple to implement; however, they lack specificity and are artifact-prone. Commercial ELISA kits are available for the more common octadecanoids from linoleic acid (e.g., 9-HODE, 13-HODE, EpOMEs, DiHOMEs); however, they are generally not available for other species and there is a paucity of radiolabeled octadecanoids. Accordingly, there are limitations in the application of RIA and ELISA methods for the study of octadecanoids.

UV chromatography has been used extensively for investigations of the eicosanoids and docosanoids.^472−474 However, it is less useful for many of the octadecanoids, which do not exhibit a selective absorption spectrum due to a lack of conjugated double bonds. The octadecenoids containing a cis–trans conjugated diene (i.e., HODEs and HOTrEs) possess a λ_max = 235–237 nm and a molar extinction coefficient (ε) of 27000 M^–1 cm^–1 and the corresponding keto metabolites absorb equally strongly around 275–285 nm (λ_max being solvent dependent for oxo-ODEs). Metabolites possessing one or more nonconjugated alkene moieties (e.g., HOMEs) exhibit only “end absorbance” with a weak spectrum near 200 nm and lacking in any diagnostic utility. For products with a characteristic chromophore, UV spectrometry is useful for quantitation of micromolar concentrations of authentic standards (5–100 μM). UV with diode array detection on HPLC is capable of producing high quality UV spectra on low nanomole levels of product and thus has a role in product identification. Beyond these applications, UV-based analyses are of limited utility for measurements or assay of the octadecanoids.

The coupling of a gas chromatograph (GC) to a mass spectrometer was first accomplished by Ragnar Ryhage at the Karolinska Institute in 1962, setting the stage for much of the structural elucidation work for the discoveries of eicosanoids that led to awarding of the Nobel Prize in 1982.⁴⁷⁵ Accordingly, GC-based investigations have been vital for the successful development of the oxylipin field and many of the steps in the metabolic pathway for AA were elucidated using the conventional techniques of thin layer chromatography (TLC) and GC. For GC-based work, initially flame ionization (FID) and electron capture (ECD) detectors were employed, but the development of mass spectrometer detectors increased the specificity of the measurements. The GC-MS methods have been a staple of oxylipin measurements for several decades.^476,477 This approach requires derivatization of the oxylipins, generally targeting the carboxylic acid to form the methyl ester with diazomethane or the trimethylsilyl (TMS) ester, while pentafluorobenzyl bromide has been used to form the pentafluorobenzyl esters.⁴⁷⁸ In addition, oximes of keto-metabolites can provide additional benefits, and have been used to profile an array of structurally labile plant octadecanoids.⁴⁷⁹ Unfortunately, poor sensitivity due to substantial fragmentation can be a limiting factor in GC-MS detection strategies. The formation of pentafluorobenzyl esters can limit fragmentation and greatly enhance sensitivity. However, the bulk of this derivative can overwhelm the chromatographic resolution of structurally simple regioisomers like epoxides. In this case, regioisomer purification prior to analysis may be required prior to detection if regioisomeric abundance is of interest. This limitation can be overcome by epoxide-directed derivatizations.⁴⁸⁰ For example, the use of 2,3,5,6-tetrafluorobenzenethiol to derivatize arachidonate and linoleate epoxides was demonstrated, with the hydroxysulfanyloctadecanoids having detection limits of 5 pg/μL.⁴⁸⁰ An issue with TMS esters is that they can be readily hydrolyzed and therefore require care in their handling and storage. While still a useful method for oxylipin measurement, the predominant application has shifted away from GC-MS with the advent of LC coupled to electrospray ionization (ESI).

LC-MS has become the preferred method for measurement of oxylipins. While GC-MS and LC-MS have comparable sensitivity in terms of lower limit of detection (LLOD) and lower limit of quantification (LLOQ), the initial primary advantage of LC-MS was its simplicity due to a lack of need for derivatization. For in-depth reading, multiple reviews have been written on the LC-MS quantification of oxylipins.^481−490 The majority of the LC-MS methods employ solid phase extraction with mixed-mode or reversed phase sorbents as a cleanup and concentration step prior to measurement, limiting matrix specific ion suppression. The extraction protocols are optimized on a matrix- and analyte-specific basis and can significantly affect both absolute and apparent oxylipin recoveries. Generally, the methods employed for octadecanoids are identical to those used for other oxylipins and these efforts are not reviewed here.⁴⁹¹ The key points include evaluation of recovery, matrix-effects, and ion-suppression. More recently, some simplified analysis schemes have emerged using protein precipitation by organic solvent (e.g., acetonitrile, methanol). While this approach can be associated with considerable matrix and ion-suppression effects, if sufficient dilutions are employed, robust and sensitive methods can result.^399,492 These methods routinely target over 100 oxylipins along with other metabolites, with quantification in good agreement with methods using more extensive cleanup. For many oxylipins, there is a potential for artifactual formation (or destruction) during the sample collection and cleanup steps. In particular, the activation of platelets during blood collection can result in commensurate increases in LOX-derived compounds (e.g., 12-HETE, 15-HETE, leukotrienes)⁴⁹³ as well as thromboxanes. This issue is less relevant for the octadecanoids (Table 3), which are less dependent upon LOX and COX activity (in mammals). In studies of oxylipin levels in tissue samples, it is important to instantaneously stop enzymatic generation of products during handling (i.e., by freezing in liquid nitrogen or addition of an organic solvent such as 2-propanol). Artificial formation by autoxidation can be minimized by adding butylated hydroxytoluene (BHT) or another antioxidant; however, many analytical workflows for oxylipin quantification no longer use antioxidants. Despite these challenges, an interlaboratory analysis using a harmonized LC-MS-based protocol for 133 oxylipins was able to achieve low technical variability.⁴⁶¹ Some of these factors affecting variability in observed free oxylipin levels in mammalian tissues have been recently reviewed.⁴⁹⁴

The dominant trend in the literature for the past 20 years has been the development of increasingly broad coverage oxylipin profiling methods. Newman and colleagues developed one of the first LC-MS profiling methods for linoleate and arachidonate derived epoxides and vicinal diols in 2002,⁴¹⁷ measuring 13 oxylipins in 31 min with LOQ ⩽1.5 nM. Further developments in the field due to the availability of commercial compounds, advances in column technology, and the attention of dedicated analytical chemists dramatically increased the number of oxylipins that could be analyzed in a single run. Edward Dennis and co-workers developed some of the first large-scale oxylipin metabolic profiling methods in 2007, reporting the analysis of 60 eicosanoids in 16 min.⁴⁹⁵ In 2009, Yang et al. measured 39 oxylipins in 21 min with LOQ ranging from 20 pM-10 nM,⁴⁹⁶ which included for the first time focus on the HODEs, oxo-ODEs, EpOMEs, DiHOMEs and TriHOMEs. A number of general oxylipin profiling methods were subsequently published; however, the octadecanoid coverage was limited to the primary commercially available standards from LA (EpOMEs, DiHOMEs, 9- and 13-HODE, 9- and 13-oxo-ODE as well as the TriHOMEs). The analogous octadecanoids from ALA became commercially available and were commensurately included in the profiling methods (EpODEs, DiHODEs, 9- and 13-HOTrE, 9- and 13-oxo-OTrE). Ramsden and colleagues expanded these efforts to focus on the octadecanoids, measuring 57 oxylipins of which 28 were octadecanoids including novel hydroxy-epoxy- and keto-epoxy-octadecenoic acids.⁴⁹⁷ Multiple oxylipin profiling methods have since been published,^{398,487,498,499} with Schebb and colleagues publishing a number of comprehensive oxylipin panels that include multiple octadecanoid species.^500,501 A more recent effort focused on the microbial products of C18-FAs and developed a method for 45 different octadecanoids derived from LA, ALA and GLA.⁵⁰² Advances in octadecanoid-specific methods are proceeding in alignment with the availability of the analytical standards.

There have been a number of specialized LC-MS methods developed to improve oxylipin analysis. The use of micro-UHPLC was shown to be useful for low volume analysis of oxylipins.⁵⁰³ Polarity switching is regularly used in profiling methods.⁵⁰⁴ Kampschulte et al. developed a multiple heart-cutting achiral–chiral 2D-LC-MS method that enables simultaneous oxylipin quantification and determination of stereochemistry.⁵⁰⁵ In an attempt to decrease the time required for sample preparation and analysis, online SPE-LC-MS/MS methods for oxylipins have been developed.⁴⁹¹ Nontargeted methods have also been developed using high resolution mass spectrometry (HRMS); however, these methods generally report oxylipin identification at the MS1 level, which results in significant uncertainty for reporting the different oxylipin isomers.⁵⁰⁶ The development of oxylipin-specific libraries will enhance the utility of these approaches as demonstrated by Galano and colleagues who employed molecular networking with high resolution MS/MS data to expand the strategies for oxylipin annotation.⁵⁰⁷ Attempts have also been made to improve the sensitivity via the use of nitrogen-containing derivatization agents including primary amines, secondary amines, aromatic amines, hydrazines and hydrazides, and hydroxylamines.⁵⁰⁸ For example, Bollinger et al. reported a derivatization reagent N-(4-aminomethylphenyl)pyridinium (AMPP) that can be coupled to eicosanoids via an amide linkage to improve sensitivity by 10–20-fold.⁵⁰⁹ This method has not yet been applied to octadecanoids, but was successfully used for the analysis of fatty acids.⁵¹⁰

Determining the chirality of stereocenters in oxylipins is important for understanding the route of formation as well as biological function. The different approaches for chiral-based analyses have been previously reviewed.⁵¹¹ Initial efforts relied on normal phase chromatography;⁴⁹⁵ however, developments in column technologies have resulted in the availability of reversed phase compatible columns.³⁹⁸ While initial particle sizes were large (<3 μm),⁵¹² newer phases have smaller particle size (<2 μm) with the associated improved separation.⁵¹³ For example, Fuchs et al. employed reversed phase chiral chromatography to separate 16 different isomers of the LA-derived TriHOMEs using an >100 min isocratic gradient.⁴³² Supercritical fluid chromatography (SFC) has been used to perform chiral analysis, with a recent octadecanoid-specific method capable of simultaneously screening the HODEs, oxo-ODEs, EpOMEs, DiHOMEs and TriHOMES from LA as well as the corresponding analogues from ALA and GLA in addition to multiple microbial metabolites.^514,515 This method is the first dedicated method for octadecanoid metabolic profiling, requiring the custom synthesis of many of the analytical standards.

In terms of other methods for oxylipin analysis, nuclear magnetic resonance (NMR) has also been used extensively and is ideal for confirmation of synthesized compound structure and purity. NMR is particularly useful for assigning double bond configurations and stereochemistry of oxylipins, which is challenging to perform via mass spectrometry-based methods. Capillary electrophoresis (CE) has been used for analyzing fatty acids, but there is little information on applications in oxylipins.⁵¹⁶ Limited efforts have demonstrated the ability to measure eicosanoids,^517−519 LA-derived epoxy octadecenoic acid isomers⁵²⁰ and LA-derived hydroperoxides.⁵²¹ Specialized methods such as immunoaffinity chromatography (IAC) are only commercially available for a few eicosanoids.⁵²² Ion mobility spectrometry (IMS) has strong potential for increasing the specificity in oxylipin analysis with a combination of IMS- and LC-based separation. In early efforts, Kyle et al.⁵²³ successfully employed IMS to separate 9-HODE and 13-HODE as well as 9-oxo-ODE and 13-oxo-ODE. Jónasdóttir et al. then demonstrated that differential mobility spectrometry (DMS) was able to separate leukotrienes that partially coeluted by LC-MS/MS.⁵²⁴ Recently, Fedorova and co-workers evaluated the implementation of IMS in LC-HRMS workflows for eicosanoids and concluded that while deprotonated ions of eicosanoids were poorly resolved, adducts evidenced good separation.⁵²⁵ While the application of IMS to improve the separation of complex oxylipin mixtures is promising, there is currently a lack of collision cross section (CCS) libraries, which is a significant limitation. Recently, mass spectrometry imaging (MSI) approaches have been developed to detect oxylipins in intact tissue. Lanekoff and colleagues successfully visualized prostaglandins in uterine tissue using desorption electrospray ionization (DESI)-based imaging, reporting that PGD₂ tissue localization and abundance could not be inferred by COX distribution and that it was necessary to perform in situ imaging of the prostanoids.⁵²⁶ In addition, they developed targeted methods to discriminate prostanoid isomers (PGD₂ vs. PGE₂) using cationization with silver ions.^527,528 Nano-DESI interfaced with a triple quadrupole has been used to image monohydroxylated isomers of AA (e.g., 11-HETE vs 12-HETE);⁵²⁹ however, the selectivity of the annotated isomers depended upon unique transitions, which are challenging to establish for oxylipins. Accordingly, it is recommended to confirm the structural identify with an orthogonal approach (i.e., LC-MS of the same tissue). Octadecanoids have also been successfully imaged using a novel DESI-MRM-based approach, visualizing metabolites from both LA and ALA in lung tissue.⁵³⁰ Accordingly, the use of DESI-MRM-based MSI to map the spatial heterogeneity of octadecanoids (and other oxylipins) is promising. This approach should be able to provide useful insight into the in situ signaling pathways associated with octadecanoid production.

In order to advance octadecanoid research, there is a need to further develop targeted methods for octadecanoid measurement. The use of chiral chromatography to determine the route of formation (i.e., enzymatic or autoxidative) is an important component of octadecanoid method development. However, the biggest obstacle is the paucity of available analytical standards. Commercial suppliers have begun to recognize the increasing interest in octadecanoids and continue to improve their offerings of standards. The microbial-derived compounds are of particular interest and further standards will be required to develop this research area.

10. Bioactivity of Octadecanoids

10.1. Cellular Targets of Action

Octadecanoids display an array of biological effects with actions dependent on their structure, their site of action, and their context of formation. For the oxygenated octadecanoids, these effects are propagated from interactions with various nuclear receptors (e.g., peroxisome proliferator activated receptors (PPARs), G-coupled protein receptors (e.g., GPCR132; i.e., G2A), transient receptor potential receptors (TRPs) including the TRP vanilloid type 1 (TRPV1), and processes modulating cell surface receptor translocation (e.g., vitronectin receptor)). Importantly, both regio- and enantioselectivity in receptor activation have been reported in some of these processes as discussed below.

For the nitrated octadecanoids, effects are also initiated from multiple interactions including those at nuclear receptors (e.g., Nrf2, PPARs, NFκB, heat shock response, stimulator of interferon genes (STING)), and cellular receptors (e.g., angiotensin II type 1 receptors), as well as by direct modulation of enzyme activity (e.g., soluble epoxide hydrolase (sEH), 5-LOX, xanthine oxidase).⁵³¹ These actions appear to largely stem from their ability to participate in reversible Michael additions with reactive thiols.⁵³² Recent efforts identified 184 high confidence intracellular NO₂-FA targets in macrophages, expanding the reported list to include such important targets as the retinoid X receptor and Toll-like receptors.⁵³³ The aggregate activity of the effects of oxygenated octadecanoid along with points of known NO₂-FA interactions are shown in Figure 1.

Aggregated octadecanoid mechanisms of action with points of nitro-octadecanoid interactions indicated. Linoleic acid (LA)-derived octadecanoids are shown as an example. Structural analogues are expected to exhibit similar interactions, with differential potency; however, in most cases those data are not yet available. Lines indicate direct (solid) and indirect (dashed) octadecanoid interactions, while line terminations indicate positive (arrow heads) and negative (bars) actions. In mammals, enzymatic and abiotic linoleate oxygenations yield a variety of octadecanoids.^{58,169,534−538} These products reportedly bind PPARs,^170,539,540 the G-coupled protein receptor GPR132,^534,541 transient receptor potential (TRP) channels,^542,543 the vitronectin receptor⁵⁴⁴ and β-catenin.⁵⁴⁵ In addition, the microbial enzyme CLA-HY can produce metabolites in the gut^541,546 which activate GPR40 (i.e., FFAR1). These ligand binding events influence cell proliferation,^545,547 inflammation,^{71,541,546−550} lipid metabolism,⁵⁴⁹ cell adhesivity,⁵⁴¹ and insulin secretion,⁵⁵¹ along with the central and peripheral perception of pain. Actions within cell types will reflect the availability of biosynthetic machinery, receptors profiles, and downstream response elements, and can be regulated at multiple points by reversible nitroalkylation of redox sensitive thiols, indicated by a yellow S in the displayed system where known. The line colors indicate the directionality of impact (green increase, red increase). Octadecanoid nomenclature is as described in Scheme 3.

PPARs are important octadecanoid sites of action. Eighteen carbon alcohols, ketones, epoxides, and nitro-lipids are all ligands, activators and suppressors of PPARs.^{170,552−556} For instance, 13-HODE can activate PPARα and PPARγ, yet suppress PPARδ, while 9(10)-EpOME is a PPARγ activator.^540,557 In addition, substantial regio- and enantioselective interactions with PPARs are reported. 13-HODE is the most potent PPARγ activator within the linoleate derived alcohols, while the autoxidation product 10(E),12(E)-9-HODE is the weakest.⁵⁵⁸ Moreover, while 10-HODE, 12-HODE and 9(E),11(E)-13-HODE increase PPARγ-mediated transcription, the 10(E),12(E)-9-HODE regioisomer reduces this action.⁵⁵⁸

The stress inducible GPR132 is another important site of action for the HODEs. While the downstream effects of GPR132 vary by cell type and the coactivation of other processes, it was first identified as a factor causing the accumulation of cells in G2/M of the cell cycle with tumor suppressor-like properties.⁵⁵⁹ GPR132 is activated by oxidized nonesterified fatty acids, but not their esterified forms, with 9-HODE being a particularly potent activator.^{542,560−562} Moreover GPR132, is transcriptionally suppressed by PPARγ activation, indicating a point of potential regulatory cross talk by octadecanoids.⁵⁴⁸ For example, in the context of atherosclerosis, the early activation of 15-LOX-1 leads to 13-HODE production and PPARγ-dependent enhanced lipid clearance. Moreover, HODEs may also be involved in atherosclerosis risk reduction. For instance, 13-HODE may increase reverse cholesterol transport early in atherosclerosis through PPARα activation.^563−565 It has been suggested, however, that with the oxidative stress-dependent generation of racemic 9- and 13-HODEs, 9-HODE-driven GPR132-dependent pro-inflammatory processes take over.⁵³⁴

HODEs are also reported activators of the TRPV1, but their importance as endogenous regulators of TRPV1-dependent physiology (e.g., nociception) is still debated.⁵⁶⁶ Regardless, in the context of chemically induced oxidative stress and nociception in the dorsal root ganglion, 9-HODE-dependent activation of GPR132-PKC-TRPV1 coupled signaling appears credible.⁵⁴⁶ In addition, EpOMEs and DiHOMEs are also activators of both TRPV1 and TRPA1.⁵⁶⁷ A comprehensive evaluation of octadecanoids as effectors of the TRP-superfamily would appear to be extremely valuable and may unravel potential mechanisms of action in various cell systems.

The electrophilic NO₂-FAs impact physiological processes by modulating the function of a variety of proteins through the reversible modification of regulatory thiol and histidine residues. Such processes have been extensively reviewed elsewhere, but are summarized and integrated here.^76,532 These NO₂-FA targets include redox sensing mechanisms (e.g., Keap/Nrf2), metabolic and growth regulators (e.g., glyceraldehyde-3-phosphate dehydrogenase, Rad51, PPARs) and inflammatory modulators (e.g., NFκB), along with other targets with broad physiological impacts (e.g., sEH, TRPV1). It is particularly noteworthy that a variety of protein targets of electrophilic lipids have been identified, and while only a subset of these have been investigated for interactions with the NO₂-FAs, these lipids do share some specific amino acid targets with other electrophilic lipids like 15-deoxy-Δ^12,14-prostaglandin J₂ (15-deoxy-Δ^12,14-PGJ₂) and hydroxynonenal.⁷⁶ The demonstration of cellular NODE catabolism, with prostaglandin reductase-1 identified as a functional nitroalkene reductase, has also provided a satisfying endogenous mechanism to halt such nitro fatty acid-dependent signaling cascades.⁵⁶⁸ Therefore, this mechanism alone allows the NO₂-FAs to have broad actions in both plants and animals, many of which likely remain to be described.^105,314,569

10.2. Inflammation and Immunomodulation

Octadecanoids, like other oxygenated fatty acids including eicosanoids and docosanoids, are involved in the regulation of inflammatory processes, with both pro- and anti-inflammatory activity. For instance, μM concentrations of 13-HpODE, but not 13-HODE, stimulate RAS-dependent inflammatory signaling cascades and induce the formation of the transcription factor NFκB in vascular smooth muscle cells.⁵⁷⁰ However, both 9- and 13-HODEs can induce the maturation of monocytes into macrophages through PPARγ-dependent processes.⁵⁷¹ The 15-LOX-1 dependent formation of 13-HODE has numerous demonstrated anti-inflammatory effects. For instance, Iversen et al. showed that 13-HODE inhibits the human neutrophil production of LTB₄in vitro.⁵⁷² Another LOX product, the nonvicinal diol 9,16-DiHOTrE, has been shown to inhibit recombinant COX and 5-LOX, and decrease the production of pro-inflammatory mediators including LTB₄ and prostaglandins. The effect was the same for both the 9(R),16(S)- and 9(S),16(S)-stereoisomers.¹⁰¹ 13-HODE can also inhibit platelet-activating factor (PAF)-induction, but amplifies formyl-methionyl-leucylphenylalanine (fMLP)-induced polymorphonuclear leukocyte degranulation.⁵⁷³ Alternatively, GRP132 activation by 9-HODE, but not 13-HODE, stimulates pro-inflammatory cytokine production, and cell cycle arrest in normal human epidermal keratinocytes,⁵⁶² but not GRP132-dependent activities in macrophages.^574,575 Anti-inflammatory effects were also demonstrated for 13-HOTrE and the corresponding hydroperoxide 13-HpOTrE, with both compounds decreasing pro-inflammatory cytokine/enzyme levels while simultaneously increasing anti-inflammatory cytokines in LPS-challenged RAW 264.7 and mouse peritoneal macrophages. This anti-inflammatory activity is induced by inactivation of the NLRP3 inflammasome complex via activation of PPARγ. Additionally, both metabolites also deactivated autophagy and induced apoptosis in LPS challenged macrophages.⁶⁷ The role of ALA metabolites in the resolution of inflammation and immunomodulation have been extensively reviewed by Cambiaggi et al.⁵⁷⁶

Ketones produced from both LA and ALA have been found to exert anti-inflammatory effects through PPAR-mediated processes. Altmann et al. showed that 13-oxo-ODE can activate PPARγ, inducing transcriptional repression of pro-inflammatory factors and ameliorating colitis and mucosa inflammation in human epithelial colon cells,¹⁷⁰ whereas the two ALA-derived ketones, 9-oxo-OTrE and 13-oxo-OTrE inhibited inflammatory responses by significantly decreasing nitric oxide (NO) and TNF-α release in a LPS-stimulated RAW 264.7 macrophage cell line.⁸² The anti-inflammatory role of ALA was evaluated in M1-like macrophages from THP-1 cells, resulting in an increase in both ALA- and LA-derived octadecanoids and a reduction in LPS-induced IL-1β, IL-6, and TNF-α production.⁵⁷⁷ The authors concluded that ALA and its associated octadecanoids may act to dampen the inflammatory phenotype of M1-like macrophages.

A role for epoxy and dihydroxy octadecanoids in inflammatory processes is also slowly emerging. For instance, the generation of epoxides from LA during inflammation limited the accumulation of pro-inflammatory Ly6chi monocytes and pro-inflammatory phenotype macrophages.⁶⁹ The 9(10)-EpOME and 9,10-DiHOME can activate inflammation-associated transcription factors NFκB and AP-1,²⁸⁷ while 9,10-DiHOME promotes neutrophil chemotaxis⁵⁷⁸ and inhibits the neutrophil respiratory burst.⁵⁷⁹ Interestingly, acute inflammation causes a rapid regiospecific decline in the constitutively high levels of EpOMEs in the mouse peritoneal cavity, with the 9(10)-EpOME being more affected than the 12(13)-regioisomer.⁶⁹ These changes paralleled responses of 9-HODE to this acute inflammatory challenge. Notably, serum levels of 12,13-DiHOME were significantly elevated in severe burn-injured mice, causing immune cell dysfunction through hyperinflammatory neutrophilic and impaired monocytic actions.⁶⁸ A 12,13-DiHOME-induced neutrophil dysfunction has been implicated in intralipid-associated immunosuppression in men (no investigation was performed in women).⁵⁸⁰ Importantly, the 12,13-DiHOME has also been identified as a gut microbiota-derived octadecanoid that can impact the inflammatory state of the host. Two studies from Lynch and colleagues demonstrated increased levels of 12,13-DiHOME associated with overexpression of bacterial sEH in neonates with atopic asthma.^70,71 Intra-abdominal treatment with 12,13-DiHOME induced pulmonary inflammation and decreased the number of regulatory T (Treg) cells in the lungs of mice.⁷¹ Similarly, treatment of human dendritic cells with this diol reduced anti-inflammatory cytokine secretion as well as the number of Treg cells in vitro, via the alteration of PPARγ-regulated gene expression.⁷¹ The 12,13-DiHOME also facilitated the maturation and activation of stimulated neutrophils, while impeding monocyte and macrophage functionality and cytokine generation. A recent study demonstrated a pro-inflammatory role for 12,13-DiHOME by enhancing NLRP3 inflammasome activation in human macrophages.⁵⁸¹

Recent work has further highlighted the potential for microbial octadecanoids to influence mammalian inflammation. Two ALA-derived metabolites produced by lactic acid bacteria, 9(Z),15(Z)-13-HODE and 9(Z),15(Z)-13-oxo-ODE, were shown to favor the differentiation of macrophages into the anti-inflammatory M2-type via activation of GPR40 receptor.⁵⁸² The LA-derived 12(Z)-10-HOME, produced by gut bacteria, was recently identified as a potent mediator of inflammation. Both the 12(Z)- and 11(E)-isomers are PPARα agonists, while the 12(Z)-isomer is recognized by GPR40.⁵⁸³ Administration of 12(Z)-10-HOME resulted in reduced intestinal inflammation in DSS-induced colitis and protection from fat-induced obesity in mice.⁵⁸⁴ The same compound was found to down-regulate the pro-inflammatory response while activating the nuclear factor erythroid 2 (NF-E2) p45-related factor-2 (Nrf2)-induced cytoprotective defenses in LPS-matured dendritic cells and MODE-K murine intestine cell lines. Moreover, 12(Z)-10-HOME stimulated an anti-inflammatory cytokine pattern and dampened the production of pro-inflammatory mediators in the extracellular space of LPS-matured dendritic cells.⁵⁸⁵ Finally, the production of 12(Z)-10-HOME by gut bacteria reduced the production of pro-inflammatory mediators from arachidonic acid by diverting the excess of LA from the eicosanoid cascade.⁵⁸⁴

Nitro fatty acids also exhibit an array of anti-inflammatory properties.^586,587 For instance, 10-NOME and 9- and 12(E/Z)-NODEs can reduce monocyte chemoattractant protein-1 (MCP-1) and interleukin 6 production.^588,589 In addition, by forming reversible post-translational modifications of Kelch-like ECH-associated protein (Keap) 1, NODEs can lead to nuclear factor (erythroid-derived 2)-like 2 (Nrf2) release from the Keap1/Nrf2 complex, with subsequent Nrf2 translocation to the nucleus and the induction of an array of antioxidant genes.^590−592 The direct modification of NFκB by NODEs can also inhibit its inflammatory signaling capacity.⁵⁹¹ The cyclic guanosine monophosphate–adenosine monophosphate (i.e., GMP-AMP) synthase-stimulator of interferon genes (cGAS-STING) signaling pathway is a fundamental system involved in the innate and adaptive immune response to infection and inflammation. Activation of this pathway stimulates both interferon regulatory factor3 (IRF3) and NFκB activation and an array of downstream pro-inflammatory responses (Figure 2).^593,594 The modification of the transmembrane portion of STING at Cys88 and Cys91 10-NOME blocks the palmitoylation-dependent recruitment activation of this pathway, suppressing interferon release in both human and murine cells.⁵⁹⁵

Nitrated fatty acid (NO₂-FA)-mediated anti-inflammatory and cytoprotective effects. The nitration of fatty acids, particularly conjugated fatty acids, produces reactive electrophiles that can participate in Michael additions with exposed reactive thiols (S) in a multitude of proteins to elicit an array of anti-inflammatory and cytoprotective effects. For example, direct nitro alkylation of xanthine oxidoreductase (XOR) diminishes its activity, reducing the production of reactive oxygen species (ROS). In a more complex interaction, nitro alkylation of the chaperone protein disulfide isomerase (PDI) directs this protein to proteosomal degradation, preventing p47 oxidation, protein kinase C (PKC)-dependent phosphorylation and assembly into the NADPH oxidase (NOX) activating complex, again reducing ROS production. NO₂-FA modifications to Keap1 (Kelch-like ECH-associated protein) prevent de novo nuclear factor erythroid 2-related factor 2 (Nrf2) capture, ubiquitination, and degradation. The resulting Nrf2 buildup translocates to the nucleus followed by heterodimerization with small musculoaponeurotic fibrosarcoma protein (sMaf). The Nrf2–sMAf heterodimer binds to the antioxidant response element (ARE) transactivating a battery of antioxidant and detoxification genes (e.g., glutathione (GSH), NAD(P)H quinone dehydrogenase 1 (NQO1), catalase (CAT), superoxide dismutase (SOD), and heme oxidase (HO-1). Nitroalkylation of NF-κB p65 at Cys-38 also results in proteasomal degradation of this component reducing the availability of the NF-κB heterodimer of this redox-sensitive transcription factor. NO₂-FAs can also nitroalkylate IκB kinase (IκK), retarding IκB-activated NF-κB-dependent inflammatory signaling. The line colors indicate the directionality of NO₂-FA impact (green increase, red decrease).

In addition to disrupting inflammatory signaling cascades at the receptor and transcription factor level, the NOMEs can also modulate the function of a variety of enzymes involved in oxylipin biosynthesis and degradation. For instance, NODE interactions with catalytic cysteines are able to inactivate lipoxygenase as a class.^105,596 In contrast, NODE-additions to Cys-521 of the murine sEH inhibits fatty acid epoxide hydrolysis and would be expected to have an anti-inflammatory effect by extending the action of endogenous epoxy-fatty acid.^161,597,598

10.3. Octadecanoids in the Mammalian Skin Permeability Barrier

The integrity of the mammalian skin permeability barrier depends on octadecanoids and specifically on LA and its LOX products. The classic paper by Burr and Burr in 1929 first described the existence of dietary EFAs and noted that a feature of EFA deficiency is development of a scaly skin and its resolution by topical application of EFA.⁵⁹⁹ The ability of different fatty acids to correct the symptoms of EFA deficiency was studied in detail during the subsequent 50 years (most commonly by monitoring correction of the trans-epidermal water loss), and the results indicated a structural requirement for at least two double bonds positioned as in linoleate and suitable for transformation by a lipoxygenase.¹⁵ Other ω6 fatty acids such as arachidonate can cure the scaly skin symptoms of EFA deficiency, although evidence indicates their retro-conversion to linoleate for functionality in the epidermis.⁶⁰⁰

Ichthyosis, named for the fish-like scaly skin, is a human congenital disease that mimics the phenotype of EFA deficiency; it occurs in rare human families with an inactivating mutation in a gene critical for skin barrier formation.⁶⁰¹ There are over a dozen lipid-related genes whose primary action is epidermal barrier formation,⁶⁰¹ and these include three working in series on linoleate, starting with 12(R)-LOX, then eLOX3,⁶⁰² then the short-chain dehydrogenase-reductase SDR9C7.⁶⁰³ Loss of any one of these results in ichthyosis in humans while the gene knockout in mice has a neonatal lethal phenotype; rodent pups cannot survive the trans-epidermal water loss and die of dehydration within hours of birth.^603−606

Linoleate in the outer barrier layer of the epidermis is esterified in the skin-specific Ceramide-EOS (Esterified Omega-hydroxy Sphingosine), and is the substrate for 12(R)-LOX, eLOX3, and SDR9C7 (Scheme 81). As shown in Scheme 81, 12(R)-LOX forms the 9(R)-hydroperoxide on linoleate esters, which is isomerized by eLOX3, and the NAD-dependent dehydrogenase SDR9C7 oxidizes the 13-hydroxy group to the ketone.^180,603 These transformations are required for the covalent binding of ceramides to the epidermal barrier proteins,^{180,603,606,607} forming a substructure known as the corneocyte lipid envelope, detectable by electron microscopy.⁶⁰⁸ Each of the three mouse knockouts results in loss of covalent binding of ceramides and absence of the corneocyte lipid envelope.

The pathway to the linoleate triols becomes especially prominent after knockout of the Sdr9c7 gene in mice. The esterified linoleate triol levels increase 100-fold in mouse epidermis after genetic disruption of Sdr9c7.⁶⁰³

This result points to a rapid turnover of the pathway, with a large buildup of triols as a side product when the route to the epoxy-enone is disrupted. The epoxide hydrolase EH3 (EPHX3) was shown to be responsible for epoxide hydrolysis of the esterified Cer-EOS-epoxyenone in epidermis.¹⁸⁷ The chemical reactivity of the epoxy-enone moiety is the basis for a postulated mechanistic link between the requirement for the 12(R)-LOX pathway of Cer-EOS metabolism and covalent binding of the ceramides (Scheme 82). Simple chemical inspection of this substructure indicates its proclivity for formation of Michael adducts with cysteine and histidine and Schiff base formation with lysine (Scheme 83).

Scheme 82 — The linoleate alkyl chain is shown in red.

The reactions of linoleate epoxy-enone methyl esters with the free amino acid lysine were reported in 1995 and shown to include the formation of pyrrole adducts.^609,610 Pyrrole formation would represent a stable adduct formation although reaction with the epoxy-enone methyl esters in vitro was extremely sluggish under the conditions reported (CH₃CN/H₂O, 7 days at rt). The reactions of analogues of the linoleate epoxy-enone with imidazole nucleophiles were studied by Sayre and co-workers.³³⁹ Rates of reaction of the epoxy-enones with imidazole and NR-benzoyl-l-histidine were reported and the adducts identified by LC-MS. Nonetheless, in a direct comparison of the reactions of lysine, histidine, serine and cysteine with a synthetic keto-epoxide analogue, only Cys adducts were detectable after 1h reaction at 37 °C in physiological buffer, and Ohno and co-workers went on to identify the adducts as formed by Michael addition to the conjugated enone.⁶⁰⁷ Michael addition is a facile and potentially reversible reaction, and back in 2011, along with the original characterization of LOX involvement in the metabolism and covalent binding of ceramides to protein, reversibly bound Cer-EOS-keto-epoxide was isolated from covalent attachment to mouse epidermal proteins.¹⁸⁰ Ohno et al. made use of this reversibility to compare the proportions of reversibly and irreversibly bound ceramide covalently bound to mouse epidermal proteins; the results indicated 60% of the total as reversibly bound, with 46% as irreversibly bound⁶⁰⁷ (the latter potentially comprised of ester conjugates with glutamate as reported for human epidermis in 1998⁶¹¹). The raison d’être of the LOX metabolism of the epidermal barrier lipids is to promote the covalent binding of ceramides and formation of the CLE,^180,603 and as the evidence stands, a major component of the Cer-EOS oxidized through the 12(R)-LOX pathway is covalently bond via Michael addition to Cys residues in the skin barrier protein.⁶¹¹ While not reported to the best of our knowledge, it is interesting to speculate that ^•NO₂-dependent nitration of the epoxy-enone could also occur, as elevated levels of (E,Z)-NODEs can accumulation in inflamed skin.⁶¹²

Other LA regioisomers play important roles in the correct function of human epidermis. Sebaleic acid, the 5(Z),8(Z)-regioisomer of LA, is secreted by sebaceous glands and is the main component of sebum and of sebaceous cell membrane phospholipids. Increased sebum production results in localized imbalance of fatty acid abundance in skin, with locally decreased levels of LA, which is replaced by sebaleic acid, in follicular epithelium. This was identified as a factor to potentially favor the insurgence of acne in hyperseborrheic individuals.⁶¹³ A more recent study identified a bioactive metabolite of sebaleic acid, 6(E),8(Z)-5-oxo-ODE, which stimulated calcium mobilization in human neutrophils and induced desensitization to 5-oxo-ETE (5-KETE), but not LTB₄, indicating that this effect was mediated by the oxo-eicosanoid receptor (GPR170). 6(E),8(Z)-5-oxo-ODE and its 8-trans-isomer were equipotent with 5-oxo-ETE in stimulating actin polymerization and chemotaxis in human neutrophils. Because of these chemoattractant properties, 6(E),8(Z)-5-oxo-ODE could be involved in neutrophil infiltration and support acne and seborrheic dermatitis.¹⁹¹

10.4. Nociception

The oxylipins involved in persistent pain states (both eicosanoids and octadecanoids), their biosynthesis and role in inflammation and pain, the corresponding activated receptors, as well as the therapeutic implications of targeting lipid signaling in chronic and neuropathic pain were recently reviewed by Osthues and Sisignano.⁶¹⁴ Far less has been reported on the potential for NO₂-FAs in the regulation of pain.

In the past decade, key studies performed by the research groups of Hargreaves, Ramsden, and Sisignano have unveiled that octadecanoids are key mediators of nociceptive processes. In particular, LA-derived octadecanoids play a pivotal role in thermal and mechanical pain modulation, as well as in itch perception, acting both locally and systemically.^61,222,615 The molecular mechanisms behind nociception in rats were investigated and summarized by Domenichiello et al.⁶² High levels of LA-derived octadecanoids are present in skin due to enrichment of LA, as well as the elevated expression of genes coding for oxylipin production, and have been demonstrated to interact with the main receptors involved in the pain circuit.^62,616 For example, high levels of 9- and 13-HODEs are produced in mouse and rat skin after exposure to noxious heat,⁶¹⁷ and were significantly increased in inflamed paw tissue and in the corresponding dorsal root ganglia in the subchronic phase of inflammation.⁶¹⁸ These HODEs, as well as their oxo derivatives 9- and 13-oxo-ODEs, activate TRPV1 the main receptor for heat perception in the peripheral nervous system,⁶¹⁷ leading to allodynia and hyperalgesia in vivo in rats.⁶¹⁵ Hargreaves and co-workers determined that the biosynthesis of HODEs and oxo-ODEs in sensory neurons was due to the activity of CYP; treatment with CYP inhibitors completely abolished LA-evoked calcium influx in sensory neurons in inflammatory dental pain, whereas LOX inhibitors had no effect.⁶¹⁹ As a result, administration of the selective CYP3A inhibitor ketoconazole reduced postburn thermal allodynia in rat hindpaw skin by preventing TRPV1 activation by HODEs and oxo-ODEs,⁶²⁰ while local treatment with anti-HODE antibody reversed inflammatory heat hyperalgesia.⁶¹⁹ Oxidative enzymes (e.g., CYP) are rapidly upregulated in immune cells in human dental pulpitis and in trigeminal nociceptive afferent neurons after orofacial inflammation, resulting in a great capacity to generate lipid mediators able to activate the pain circuits.⁶²¹ Other CYP metabolites of LA, 9(10)-EpOME and 12(13)-EpOME, as well as their hydrolysis products 9,10-DiHOME and 12,13-DiHOME were also found to be increased in spinal cord tissue after burn injury. These four octadecanoids activated both TRPV1 and TRPA1, resulting in postburn mechanical and thermal allodynia (i.e., pain from a typically nonpainful stimulus), which could be reversed by treatment with CYP inhibitors.⁵⁶⁷ Activation of TRPV1 by 9(10)-EpOME⁶²² and 12,13-DiHOME²²² resulted in increased TRPV1-dependent calcitonin gene-related peptide (CGRP)-release from sensory neurons in mice. The subsequent effect on thermal pain sensitivity is opposite for the two compounds, with the diol increasing⁶²² and the epoxide decreasing pain hypersensitivity. Reduced thermal hyperalgesia could then be achieved by in vivo inhibition of sEH, which results in a decreased concentration of the diol in the nerve tissue.²²²

Activation of GPR132 (i.e., G2A) by 9-HODE in a chemically induced model of neuropathic pain was associated with a protein kinase C-dependent TRPV1-sensitization in peripheral sensory neurons.⁵⁴⁶ The concentration of 9-HODE was strongly increased at the site of nerve injury during neuropathic pain,⁶²³ whereas increased systemic circulatory levels of both 9- and 13-HODEs correlated with pain intensity in women suffering from chronic neck pain.⁶²⁴ Blocking the production of the HODEs led to a significant relief of mechanical and thermal hypersensitivity in vivo.^617,618 Increased levels of HODEs, EpOMEs, and DiHOMEs were detected after UVB irradiation in mouse and rat skin. Given that the biosynthesis of these octadecanoids is COX-independent, this finding may explain why COX-inhibitors (e.g., ibuprofen) only show weak antinociceptive effects in UVB-induced mechanical allodynia in rodents.⁶²⁵ The identification of a CYP-dependent enzymatic pathway in pain modulation led the authors to propose CYP inhibitors as an alternative to opioids in postburn pain management, given the lower rate of adverse effects and the extremely decreased potential for generating addiction.⁵⁶⁷ The biological functions of CYP enzymes, with a focus on lipid mediators, and the therapeutic implications of CYP inhibitors were discussed in a recent review that focused on CYP2J2.⁶²⁶

The eLOX3-derived hydroxy-epoxide octadecanoids are abundant LA metabolites in skin and play a crucial role in nociception. Ramsden and co-workers identified increased concentrations of hydroxy-epoxide and hydroxy-keto octadecanoids in psoriatic skin lesions and observed that injection of these compounds in rodents enhanced scratching behavior. In particular, 9(Z)-11-OH-12(13)-EpOME and 12(Z)-11-OH-9(10)-EpOME were involved in C-fiber–mediated pain-related hypersensitivity in rats via the release of CGRP from a sensitization of neurons in the dorsal root ganglion. These two metabolites share a 3-hydroxy-(Z)-pentenyl-cis-epoxide moiety that was identified as the pharmacophore involved in nociceptor sensitization mediation.⁶¹ The same group also observed that 9(Z)-11-OH-12(13)-EpOME elicited pain-related behavior in rats,⁶²⁷ and that both 9(Z)-11-OH-12(13)-EpOME and 12(Z)-11-OH-9(10)-EpOME (as well as the corresponding keto-epoxides 9(Z)-11-oxo-12(13)-EpOME and 12(Z)-11-oxo-9(10)-EpOME) stimulated trigeminal neurons by eliciting Ca²⁺ responses, suggesting these metabolites as mediators in chronic headaches and craniofacial pain syndromes.⁶²⁸ In addition, systemic levels of 9(Z)-11-OH-12(13)-trans-EpOME correlated with the frequency of headache events in patients suffering from severe chronic daily headache and diet-induced reduction of plasma levels reduced headache hours per day and headache days per month.⁶¹ Finally, high concentration levels of 9(Z)-11-OH-12(13)-EpOME and 11(E)-13-OH-9(10)-EpOME were observed in the brain of chronic pain model rats.⁶²⁷ These efforts have focused primarily on the LA derivatives and the affinity of eLOX3 for non-LA peroxides has not been determined and involvement of non-LA octadecanoids in nociception has not been explored.^180,629 However, it would be of particular interest to investigate if octadecanoids deriving from other PUFAs (e.g., ALA, GLA) interacted with the same pain circuit receptors.

More recently, the role of 9,10,13-TriHOME in nociception in mice was identified. Surprisingly, 9,10,13-TriHOME did not induce nociceptive behavior when tested on rats. As discussed above, this compound is present in its esterified form at high levels in the mammalian epidermis and is released through hydrolysis resulting in high local levels on the skin surface. Application of 9,10,13-TriHOME caused a rapid acute/hyper-acute pain response with short duration, indicating rapid inactivation via acylation and esterification back into lipid membranes and dehydrogenase-induced conversion of the alcohol to ketone moieties. 9,10,13-TriHOME caused hypersensitivity to noxious heat and noxious cold via a TRPA1-dependent mechanism, but also required simultaneous activation of TRPV1.⁶³⁰

Dietary intervention was shown to affect nociception in mice. A diet enriched in ω6 PUFAs such as AA and LA, mimicking the Western diet, led to the development of pain hypersensitivity, spontaneously active and hyper-responsive glabrous afferent fibers, and histologic markers of peripheral nerve damage reminiscent of a peripheral neuropathy. Both LA and AA were shown to accumulate in lumbar dorsal root ganglia, where the fatty acids are released by PLA₂ and subsequently oxidized to a range of octadecanoids (e.g., the nociceptive hydroxy-epoxides, HODEs, EpOMEs, DiHOMEs). The nociceptive hypersensitivity could be attenuated by inhibiting PLA2G7 and was completely reverted by switching to an ω3 enriched diet (rich in EPA and DHA but containing lower levels of ALA).⁶³¹ A similar result was also obtained by investigation of systemic levels of oxylipins in rats subjected to diets enriched in LA or OA. In the first case, the plasma levels of pro-nociceptive LA-derived octadecanoids and AA-derived eicosanoids were significantly increased and subsequent reduction with antinociceptive ω3 oxylipins derived from EPA and DHA. The authors, however, did not characterize the rats’ behavior in relation to the diet, but classified the pro- or antinociception property of the oxylipins based on previous literature reports.⁶³² EPA and DHA-derived oxylipins were found to be accumulated in the plasma of rats subjected to the OA-enriched diet, a rather unexpected result given the fact that ALA, the precursor of EPA and DHA, was present at the same level in both diets, and OA is not metabolized to either EPA or DHA. This was consistent with the competition between LA and ALA for elongation-desaturation (as reported by Taha et al.),⁶³³ further raising the possibility that high LA intake may reduce the benefits of EPA and DHA supplementation on the basis of substrate competition. In humans, tissue concentration from ankle punch biopsies in diabetic patients evidenced a correlation between LA concentration and the necessity for neuropathic pain pharmacotherapy; all patients requiring the therapy had LA > 100 nmol/mg tissue. This result, however, was limited by the low number of investigated subjects (n = 16) and requires further investigation.⁶³¹

With respect to nitrated species, NO₂-FA production in inflamed tissues may be involved in the initial activation, and later suppression of TRPV1 and TRPA1 receptors on afferent nerves.⁶³⁴ The presence of cysteine rich ankyrin-like repeats in the N-terminus of TRPV1, TRPV1 and TRPC make these proteins sensitive to modifications and activation by multiple electrophiles, including NO₂-FAs.⁶³⁵ For instance, Cys-414 and Cys-421 within the N-terminal intracellular domain are sensitive to electrophile additions,⁶³⁶ as are cysteines in the extracellular domain between transmembrane loops 4 and 5 of TRPV1 (i.e., Cys-616, Cys-621, and Cys-633) and TRPA1 (i.e., Cys-621, Cys-641, and Cys-665).^637,638 Due to the presence of such redox-sensitive regulatory sites in these pain modulating proteins suggests a role for nitrolipids in the regulation of pain.

10.5. Cell Proliferation

The 9- and 13-HODEs are able to influence cell proliferation and apoptosis through PPARδ and PPARγ interactions in multiple cell types.⁶⁶ In 1990, Miller and colleagues demonstrated that 13-HODE could reduce the proliferation of skin in guinea pigs suffering from epidermal hyperproliferation induced by a fat-free diet.⁶³⁹ Since then, the effect of HODEs in cell proliferation has been recognized and extensively studied. For example, 9-HODE inhibits proliferation and induces apoptosis in monocyte cell lines.⁶⁴⁰ Similarly, the age-related increases in skeletal lipoxygenase metabolism appears to influence bone loss, where 9- and 13-HODEs are shown to suppress β-catenin mediated Wnt signaling, reducing osteoblast proliferation and promoting apoptosis.⁶⁴¹ Moreover, 9(S)-HODE, but not 13(S)-HODE (the equivalent (R)-enantiomers were not evaluated), inhibited the proliferation of normal human epidermal keratinocytes cells by suppressing DNA synthesis, while inducing the secretion of inflammatory cytokines IL-6, IL-8, and granulocyte-macrophage colony stimulating factor (GM-CSF).⁵⁶² Conversely, 13(S)-HODE increased DNA synthesis in Syrian hamster embryo fibroblasts, while the 13(R)-enantiomer showed no activity.⁶⁴² The demonstrated enantioselectivity of these processes argues for an enzymatic role in the regulation of the antiproliferative response mediated by octadecanoids.

The role of the HODEs in the processes of cell adhesion, apoptosis, and mitogenesis in cancer has been extensively reviewed by Vangaveti et al. in 2016.⁶⁶ Honn et al. were the first to show that 13(S)-HODE is produced by tumor cells and is able to block the deleterious effect of 12(S)-HETE, which increases the metastatic potential of low-metastatic melanoma cells. Moreover, 13(S)-HODE reduced lung colonization by high-metastatic melanoma cells.⁶⁴³ Since then, several studies supported the finding that 13(S)-HODE is able to reduce cancer cell proliferation. 13(S)-HODE has been shown to restore apoptosis in a human colorectal cancer cell line⁵⁵³ and to inhibit cell growth in a dose-dependent manner in breast cancer cell lines via down-regulation of PPARδ.⁶⁴⁴ In addition, 13(S)- and 9(S)-HODEs decreased cell growth and DNA synthesis of nondifferentiated Caco-2 cells and showed an apoptotic effect via activation of PPARγ. Conversely, 9(R)- and 13(R)-HODEs are not ligands for this receptor and resulted in increased cell growth and DNA synthesis through a different mechanism, most likely via activation of the COX pathway. Thus, the two enantiomers used different receptors and exerted contrary effects.⁶⁵ Since vitronectin receptors play an important role in cell growth and differentiation,⁶⁴⁵ the 13-HODE-dependent inhibition of vitronectin receptor translocation to the cell surface may also be involved in this compound’s impact on cell growth.^544,646,647

LA epoxides and diols have also been implicated in the regulation of cell proliferation. The 12,13-DiHOME appears to promote hematopoietic progenitor cell proliferation and mobilization through modulation of canonical Wnt signaling, a process important for neovascularization.⁶⁴⁸ In addition, the EpOMEs, but not DiHOMEs, promote hair follicle stem cycling and enhance hair growth,⁶⁴⁹ and EpOMEs can induce colon tumorogenesis in vivo.⁶⁵⁰ A recent study in early stage breast cancer profiled oxylipins and reported that levels of the 9(10)- and 12(13)-EpOME were reduced in plasma from individuals with breast cancer (n = 169) relative to healthy controls (n = 152), while 9-HODE levels were increased. The authors concluded that the observed oxylipin changes likely reflect the general status of the individual rather than changes in a specific tumor tissue. Notably, intestinal Fusobacterium nucleatum infection has been shown to activate TLR4/AKT/Keap1/NRF2 signaling leading to increased 12(13)-EpOME production and oncogenesis.⁶⁵¹ The Keap1-Nrf2 pathway has been described as “the primary protective response to oxidative and electrophilic stress”⁶⁵² and is often disrupted in oncogenic process.⁶⁵³ Interestingly, in metastatic brain melanoma, EpOMEs prevented melanoma cell invasion and macrophage polarization into M1-like macrophages.⁶⁵⁴ The NO₂-FAs including the (E,Z)-NODEs are potent activators of the Nrf2-Keap1 pathway, thereby upregulating antioxidant gene expression and inhibiting cell proliferation.^{590−592,655} As the (E,Z)-NODEs also allosterically inhibit the sEH, this linkage between Keap1-Nrf2 signaling and changes in epoxy-fatty acid metabolism present a unifying linkage between oxylipin and NO₂-FA actions.

NOMEs also have antiproliferative effects linked to nitroalkylation of RAD51 recombinase, a critical component in the DNA repair machinery.⁶⁵⁶ Another important regulator of cell cycle progression, proliferation and senescence are the Ras-GTPase associated signaling pathway.^657,658 Like other Ras proteins, H-Ras activation is linked to the canonical Raf/MEK/ERK signaling leading to the induction of proliferation and survival genes. However, H-RAS is also linked to PI3K/AKT signaling, which lead to mTOR activation facilitating protein synthesis, migration, and proliferation.⁶⁵⁸ To the best of our knowledge, interactions between NODEs and the Ras proteins have not been reported. However, Ras modification by reactive nitrogen species has been reported.¹⁹ Moreover, H-Ras Cys184 binding to the electrophilic lipid mediator 15-deoxy-prostaglandin J₂ (15d-PGJ₂) prevents palmitoylation and activates Raf/MEK/ERK-mediated signaling, without AKT pathway activation.^659,660 Together, these results suggest that modulation of H-Ras regulated cell growth by nitro octadecanoids deserves attention.

10.6. Mitochondrial Respiration

A significant body of literature exists indicating that EpOMEs and DiHOMEs are negative effectors of mitochondrial function. The LA-derived 9(10)- and 12(13)-EpOMEs were first found in rice plants and suggested to be derived from rice blast fungus.⁶⁶¹ In 1986, Ozawa and colleagues reported the structures of two LA-derived epoxides that evidenced a potent uncoupling effect in rat liver mitochondria, simultaneously with relaxation of stomach smooth muscle in a dose-dependent manner.^662,663 The epoxides (9(10)-EpOME and 12(13)-EpOME) were demonstrated to be formed by incubating LA with leukocytes collected from lung lavages, and were therefore termed leukotoxin and iso-leukotoxin, respectively.⁶⁶² However, follow-up work stated that the leukotoxin nomenclature was due to the toxic activity toward mitochondrial respiration.^662,664 The 9(10)-EpOME was shown to be produced by neutrophils in a calcium ionophore enhanced process, while little 12(13)-EpOME was observed.^662,665 Importantly, in these early investigations, the other products of LA oxidation (e.g., 9-HODE) were reported to not affect mitochondrial function. The 9(10)-EpOME was detected in rat lung lavages after long exposure to hyperoxia (60 h) and in lung lavages obtained from patients with acute respiratory distress syndrome (ARDS).⁶⁶⁶ Additionally, injection of 9(10)-EpOME (100 μmol/kg) in rats resulted in acute edematous lung injury. However, this dose is approximatively 700 times higher than the physiological level of 9(10)-EpOME in human blood (from 1.5 to 5.5 nM according to the Human Metabolome Database). These collective findings suggested that 9(10)-EpOME played an important role in the development of lung injury observed in patients with ARDS.⁶⁶⁷ However, exposure to low concentrations of 9(10)-EpOME (10 μM) caused lung edema and cellular damage without evidence for mitochondrial dysfunction in rats, while, a higher dose of 30 μM induced a significant decrease of the mitochondrial respiration rate associated with decreased ATP content in the lung tissue.⁶⁶⁸ However, the direct cardiac administration of DiHOMEs, but not EpOMEs, leads to ARDS-like symptoms and rapid death in rodents, implicating an epoxide hydrolase-dependent pathology.^669,670 The effect of anesthetic agents and sedatives upon circulatory oxylipin profiles was examined in a large RCT of healthy males (n = 160).⁶⁷¹ The authors observed that injection of the anesthetic Propofol (which is used to sedate COVID-19 patients who require mechanical ventilation in the intensive care unit) resulted in elevated circulatory levels of the DiHOMEs in contrast to Dexmedetomidine, which resulted in decreased DiHOME levels. Accordingly, the choice of anesthetic used to sedate COVID-19 patients may have implications for the observed DiHOME levels, which are associated with ARDS-related COVID-19 mortality.

Hammock and colleagues have extensively studied the sEH responsible for converting the EpOMEs to the corresponding DiHOMEs as well as the associated biologies.^58,672 Their collective efforts have established that the EpOMEs are protoxicants that require activation by sEH to the DiHOMEs. For example, the cytotoxicity of 9(10)-EpOME in ARDS observed in severe burn patients was only observed in the presence of sEH. The study showed that the resulting 9,10-DiHOME was toxic to pulmonary alveolar epithelial cells.⁶⁶⁹ These pathological levels of DiHOMEs have been found to have direct adverse effects on mitochondrial function and engage the mitochondrial permeability transition.⁶⁷³ In addition, in the context of LPS-induced cardiac inflammation, 12,13-DiHOME induces severe mitochondrial dysfunction and structural abnormalities, while stimulating inflammatory cytokine production.⁶⁷⁴ The 9,10- and 12,13-DiHOMEs also induced renal proximal tubule cell death associated with mitochondrial dysfunction, while the corresponding epoxide did not induce cell death at concentrations up to 1 mM.⁶⁷⁵ Schuster et al. reported that addition of thermally stressed corn (of which the octadecanoid content was determined by LC-MS) to the diet of mice had an effect in the regulation of mitochondrial function and in the activation of the NLRP3 inflammasome. These effects could be ascribed to the high levels of LA-derived octadecanoids (259.6 ± 21.6 nM) in stressed oil, which mediated cell death in hepatocytes through a mechanism partly dependent on capsase-1 activation.¹¹⁸ These findings suggest that dietary octadecanoids are able to directly activate the innate immune response linking lipid metabolism with innate immunity. Interesting, in vitro studies in RAW264.7 cells suggested that 9(S)- and 13(S)-HODE might have different cellular functions, with 9(S)-HODE potentially possessing cytotoxic properties,⁶⁷⁶ further evidencing the need to focus on chirality.

While pathological levels of DiHOMEs cause severe mitochondrial dysfunction, EpOMEs can induce cell death in vitro by uncoupling oxidative phosphorylation.^198,677 Notable, 12(13)-EpOME was shown to increase mitochondrial non-ADP-stimulated and oligomycin-insensitive (i.e., state 4) respiration and reduce succinate-dependent and oligomycin-sensitive (i.e., state 3) respiration in isolated rabbit renal cortical mitochondria (at relatively high concentrations of 50 mM). Concomitantly, 12(13)-EpOME decreased the potential of the mitochondrial membrane, while 12,13-DiHOME did not exhibit similar effects at these concentrations.⁶⁷⁷ Nowak et al., then showed that pretreatment of rabbit renal tubular cells with 12(13)-EpOME (but not 12,13-DiHOME) before hypoxia protected the mitochondrial functions and accelerated the recovery of the intracellular ATP levels during reoxygenation.⁶⁷⁸ Moreover, sEH deletion or pharmacological inhibition also prevents the development of mitochondrial abnormalities in models of cardiac ischemia/reperfusion inflammation.^674,679,680 Likewise, NOME perfusion is cardioprotective in a heart ischemia/reperfusion injury model. These findings provide another plausible physiological linkage between the NO₂-FA-dependent metabolism and the sEH biology. However, NO₂-FAs are also able to directly inhibit mitochondrial respiration by nitroalkylation-dependent inhibition of complex-II linked respiration.⁶⁸¹ Mitochondrial adenine nucleotide translocase 1 (ANT1) Cys-57 has also been identified as a cardioprotective target of NO₂-FA adduction.⁶⁸² ANT1, which can account for up to 10% of total mitochondrial protein,⁶⁸³ is a component of the permeability transition pore⁶⁸⁴ that regulates ADP/ATP exchange across the inner mitochondrial membrane and regulates basal proton leakage.⁶⁸⁵ Knockdown of ANT1 is associated with increased mitochondrial ROS, concomitant with increased TNFα-induced NF-κB reporter gene activity and interleukin-6 and TNFα expression.⁶⁸⁶ Therefore, nitroalkylation of ANT1, sEH, and other cellular and mitochondrial targets likely work in concert to elicit an anti-inflammatory and cardioprotective phenotype.

Because the mitochondrial inner membrane is exposed to an oxidant abundant environment and consists of high levels of cardiolipin, a LA-rich membrane lipid when dietary LA is high,⁶⁸⁷ it is not surprising that linoleate oxidation products including HODEs, DiHOMEs, TriHOMEs, oxo-OMEs, oxo-EpOMEs, HpODEs and Hp-oxo-OMEs are constitutive members of the mitochondrial lipidome.^294,688 Notably, cytochrome c and iPLA₂γ are likely involved in cardiolipin oxidation and hydrolysis in vivo,²⁹⁴ and cardiotoxic chemotherapeutics and ω3 fatty acids can modulate their levels.⁶⁸⁸ Regardless, studies of the impact of these species on mitochondrial function are extremely limited. In the context of airway inflammation, 13(S)-HODE has been shown to negatively impact mitochondrial function in airway epithelium.⁶⁸⁹ Together, these findings suggest that future efforts to evaluate the potential for cardiolipin-derived octadecanoids as endogenous mediators of mitochondrial function have great potential.

10.7. Metabolism and Hormone Modulation

Octadecanoids exert an important role in fatty acid and glucose metabolism as well as in the modulation of hormone secretion because they can activate a number of receptors linked to metabolism (e.g., PPARα,⁶⁹⁰ TRPV1,⁶⁹¹ GPR40,⁵⁸⁴ prostaglandin receptor EP3⁷⁹). In 1998, Murthy et al. were the first to show that 13-HODE interferes with the assembly and composition of triacylglycerol-rich lipoproteins secreted by intestinal cells and diminishes the secretion of triacylglycerol in intestinal Caco-2 cells.⁶⁹² Interesting recent evidence in drosophila also suggests that 9-HODE stimulates JNK activation of Forkhead box (i.e., FOXO) proteins, which are important regulators of insulin signaling.⁶⁹³ In addition, both 9 and 13-HODEs increase fatty acid binding protein 4 (FABP4) expression in leukemia monocyte cell lines in a PPARγ-dependent manner.

While a similar impact of HODEs on adipose FABP4 expression has not been demonstrated, this protein is strongly associated with the development of insulin resistance and atherosclerosis⁶⁹⁴ and research in this area is warranted.⁵⁴⁹ More recently, the role of batokines during cold-exposure or exercise has been studied by Stanford and colleagues, revealing the importance of 12,13-DiHOME in fatty acid uptake and insulin resistance.^695−697 Lynes et al. demonstrated that 12,13-DiHOME decreases body-mass and insulin resistance in humans.⁶⁴ Interestingly, 12,13-DiHOME has been reported to reduce glucose uptake and inhibit insulin-dependent signaling in myotubes.⁶⁹⁸ 12,13-DiHOME is released from BAT after 1 h of cold exposure in both rodents (4 °C) and humans (14 °C). Moreover, the levels of epoxide hydrolase were significantly higher in BAT after cold exposure, leading to an increased production of the diol from the corresponding epoxide. The increased levels of the circulating diol enhance fatty acid uptake in BAT, which is involved in heat production in adults, and reduce levels of serum triglycerides.⁶⁴ Acute-intensity bouts of exercise caused the release of 12,13-DiHOME from BAT, increasing the circulating levels and enhancing fatty acid uptake and mitochondrial fatty acid oxidation in skeletal muscles.⁶³ Acute treatment with 12,13-DiHOME in mice increased skeletal muscle fatty acid uptake and oxidation, but had no effect on glucose uptake. BAT transplantation resulted in improved cardiac functions in mice through the release of 12,13-DiHOME, which negated the deleterious effects of a high-fat diet. Acute injection with 12,13-DiHOME affected the cardiomyocytes by increasing mitochondrial respiration resulting in increased cardiac hemodynamics. Moreover, the systemic concentration was decreased in individuals with heart disease.²²¹ More recent studies report that both the 9,10-DiHOME and 12,13-DiHOME are inversely associated with BMI and activate calcium influx in mouse brown and white adipocytes in vitro,⁶⁹⁹ further supporting that activating BAT is a promising target to treat metabolic syndrome.

Ketones derived from LA and ALA interact with PPARα, affecting the intake of triglycerides and fatty acids in cells and tissues. Cellular triglyceride accumulation in the hepatocytes was inhibited by 10(E),12(E)-9-oxo-ODE, a ketone produced from LA and isolated from tomato juice, via activation of PPARα in vitro.⁶⁹⁰ The effects were confirmed by in vivo treatment of obese KK-Ay mice fed with a high-fat diet, resulting in decreased levels of triglycerides in both plasma and liver.⁷⁰⁰ Activation of PPARα was also reported for 9-oxo-OTrE, resulting in enhanced fatty acid uptake in murine hepatocytes.⁷⁰¹

Exercise-induced changes in octadecanoids have also been reported, suggesting they may report on metabolic changes associated with metabolic exertion. Elevations in HODEs and DiHOMEs induced by a 75-km bout of cycling were partially resolved in 1.5h, and fully resolved within 1d, further suggesting these compounds as markers of physiologically relevant oxidative stress.⁷⁰² Notably, exercise-induced changes in HODEs were reportedly unrelated to isoprostane formation, muscle damage, or soreness, but negatively correlated with granulocyte colony stimulating factor and interleukin 6 (IL-6), suggesting associations with neutrophil chemotaxis and inflammation in this context.⁷⁰³ Given the impact of 12,13-DiHOME on glucose uptake and insulin-dependent signaling in myotubes,⁶⁹⁸ some of these changes may report on physiological changes in muscle energy metabolism.

Interesting metabolic effects were demonstrated for octadecanoids produced by gut microbiota. Alcohols and ketones oxidized at the 10-position can activate the PPARα receptor, while the two regioisomers of 10-oxo-OME (12(Z)- and 11(E)-10-oxo-OME) can also activate PPARγ, with the former possessing the highest potency. The activation of PPARα by 12(Z)-10-oxo-OME caused an increase of adiponectin production and insulin-stimulated glucose uptake.⁵⁸³ This ketone was also able to activate the TRPV1 receptor and improve noradrenalin turnover in adipose tissues. The intake of 12(Z)-10-oxo-OME by obese and diabetic KK-Ay mice improved obesity-associated metabolic disorders, such as glucose intolerance, insulin resistance, and increased adiposity.⁶⁹¹ Recently, Miyamoto et al. showed that the corresponding alcohol, 12(Z)-10-HOME attenuates HFD-induced obesity in mice. Acute 12(Z)-10-HOME administration promoted the secretion of GLP-1, a peptide hormone associated with appetite suppression and improvement of glucose homeostasis via activation of GPR40 and GPR120.⁷⁹ Finally, 12(Z)-10-HOME activated the EP3 receptor in the gut, promoting intestinal peristalsis.⁷⁹ Dietary-derived octadecanoids therefore appear to participate in a feedback system in which the gut microbiome converts dietary fatty acids into octadecanoid lipid mediators that attenuate obesity.

Hormone modulating effects were reported for three octadecanoids: a keto-epoxide metabolite of LA, 10(E)-9-oxo-12(13)-EpOME, and two gut microbial metabolites, 11(E)-10-oxo-OME, 9(Z),15(Z)-13-oxo-ODE, derived from LA and ALA, respectively. In particular, 10(E)-9-oxo-12(13)-EpOME promoted the production of aldosterone, a hormone regulating blood pressure, at low doses (from 0.5 to 5 μmol/L in vitro), and inhibited its production at higher doses,¹⁸⁵ while both 11(E)-10-oxo-OME and 9(Z),15(Z)-13-oxo-ODE stimulated production of a cholecystokinin, a gut hormone that helps digestion and reduces appetite, via activation of GPR40 (further demonstrating the role of octadecanoids in attenuating diet).²³⁴

There are limited studies investigating the utility of octadecanoids as indicators of endocrine disruption. Markaverich and colleagues showed that THF-diols, but not the DiHOMEs, have been reported to block male sexual behavior.^704,705 While the mechanism of action is unknown, Okamura’s and Hull’s groups hypothesized that it occurs via modulation of nitric oxide-dependent pathways that control gonadotrophin-releasing hormone release.^706−708 THF-diols and the DiHOMEs are thought to act additively to disrupt endocrine function in both male and female rats at low concentrations (0.5–1 ppm), which is ∼200-fold lower than those of classical phytoestrogen endocrine disruptors (e.g., isoflavones).⁷⁰⁹ Both DiHOMEs stimulated MCF-7 breast cancer cell proliferation equivalently, but did not compete for [3H]estradiol binding to the estrogen receptor or nuclear type II sites. Oral administration of the DiHOMEs at low doses (>0.8 mg/kg body weight/day) disrupted estrous cyclicity in female rats, but did not disrupt male sexual behavior, suggesting sex-specific differences in endocrine response.⁷⁰⁵

10.8. Cardiovascular Effects

Octadecanoids possess diverse effects in the cardiovascular system, influencing cardiac muscle contraction, arterial relaxation, ischemia, and platelet adherence. Injection of very high doses of 9(10)-EpOME in dogs (15 mg/kg) caused an immediate decrease in aortic flow and cardiac failures and death in <1 h.⁵⁵ This depressed cardiac function in dogs in a dose dependent manner. Similarly in cats, injection of 2.5–25 ng of 9(10)- and 12(13)-EpOMEs into isolated papillary muscles reduced myocardial contractility, while administration to carotid arteries resulted in vasoconstriction.⁷¹⁰ While injection of OA, LA, and SA at 10 mg/kg had no significant hemodynamic changes, injection of 50 mg/kg LA exhibited cardiotoxic effects (although less than those observed with 9(10)-EpOME).⁷¹¹ The same effect was observed in guinea pig papillary muscles after acute administration of the gut bacteria metabolite 12(Z)-10-HOME at concentrations 30–300 μM.⁷⁸ At a concentration of 10 nM, 13-HODE and its precursor 13-HpODE relaxed canine coronary artery segments with endothelium after PGF_2α-induced contraction.⁷¹² This vasodilatation activity is due to stimulation of prostacyclin (PGI₂) biosynthesis and activation of the thromboxane receptor.⁷¹³ This effect was also observed with 13-HpODE (10 μM) in human pulmonary arteries after PGF_2α-induced contraction.⁷¹² Treatment with 9,10-DiHOME (250 μM) increased the coronary resistance in mouse heart after ischemia-reperfusion injury, significantly impairing the heart functional recovery.⁷¹⁴

Impacts on platelet function and cellular adhesivity are another critical site of octadecanoid action. 13-HODE reduces thrombin-induced platelet aggregation,^715,716 and platelet adherence to monolayers of cultured pulmonary artery endothelial cells.⁷¹⁵ Moreover, high density lipoproteins enriched in 13-HODE containing phosphatidylcholine dose-dependently inhibited platelet aggregation.⁷¹⁷ Notably, in endothelial cells, the intracellular association of 13-HODE with the vitronectin receptor stabilizes it in the intracellular space. Dissociation of this complex appears to allow vitronectin to translocate to the cell surface where it promotes adhesivity.^544,646,647 The 9(R),16(S)- and 9(S),16(S)-diastereoisomers of 9,16-DiHOTrE have also been shown to inhibit collagen-stimulated platelet aggregation through COX-2 inhibition.¹⁰¹

Nitro lipids are also potent vasodilators with antihypertensive effects resulting from either the direct modulation of enzyme and/or receptor function, or by influencing the expression of key gene products. As mentioned in the immune modulation and mitochondrial metabolism sections, the sEH is inhibited by nitroalkylation of a redox-sensitive cysteine residue. In mice fed CLA and sodium nitrate, the antihypertensive effects and associated increase in plasma concentrations were ablated by deletion of this redox-sensitive cysteine.¹⁶¹ In addition, nitroalkylation of the angiotensin 1 receptor reduces its responsiveness to angiotensin II stimulation,⁷¹⁸ and the ability of NOMEs to enhance endothelial nitric oxide synthase and hemeoxygenase-1 expression further contribute to the antihypertensive influences of the nitro-octadecanoids.⁷¹⁹ Therefore, as in other systems, the impact of NO₂-FAs in the vascular system are pleotropic and dependent on their soft electrophilic characteristics.

11. Octadecanoids as Biomarkers

Alteration in octadecanoid levels has been observed in multiple pathologies including neurodegenerative, cardiometabolic, hepatic, and respiratory diseases as well as with chronic pain. Given that octadecanoid fatty acid precursors are generally present in high concentration, and that octadecanoid formation can proceed as a consequence of both oxidative stress and enzymatic activity, monitoring these compounds in the context of disease risk and progression may yield useful biomarkers; however, the specificity of these compounds as independent markers of any given malady is unlikely. The following section provides an overview of the literature to date reporting the potential uses of octadecanoids as biomarkers of diseases. Much work remains to establish the specificity and sensitivity of octadecanoids as biomarkers, either alone or in combination with other metabolites.

11.1. Neurological Diseases

The study of octadecanoids in neurological diseases has to date focused primarily on Alzheimer’s disease (AD), with a few studies investigating cerebral ischemia (e.g., stroke) and multiple sclerosis. Yoshida et al. reported a higher level of total-HODEs (defined as 9-(E,Z)-HODE, 13-(Z,E)-HODE), 9-(E,E)-HODE, and 13-(E,E)-HODE) in plasma and erythrocytes of individuals with AD compared to those with vascular dementia and healthy controls.⁷²⁰ Furthermore, the levels of total-HODEs increased with the clinical severity of dementia. The diagnosis of AD was based upon probable AD (according to Diagnostic and Statistical Manual of Mental Disorders IV (DSM-IV) criteria), whereas work by Kurano et al. reported that 13-HpODE levels in brain tissue from autopsies that included histopathological examination correlated with AD clinical phenotype.⁷²¹ Unfortunately, the absolute levels of 13-HpODE, as well as other octadecanoids potentially measured, were not reported. In a larger study of 150 AD patients and 139 healthy controls, Borkowski et al. reported that CSF levels of 9(10)- and 12(13)-EpOMEs were strong predictors of AD.⁷²² Morris et al. compared oxylipin profiles in individuals with AD, both with and without T2D, and reported that plasma levels of 9,10-DiHODE and 15,16-DiHODE contributed significantly to the PLS model of AD patients with T2D; however, the associations were weak and not univariately significant.⁷²³ Shen et al. recently reported a decrease in 13-HODE in male 10-month old AD TgF344 transgenic rats in brain neutral lipids compared to wild-type controls. In AD transgenic females, 13-HODE slightly increased in brain neutral lipids and the anti-inflammatory metabolite 13-HOTrE decreased in phospholipids. Neutral lipid-bound 13-HODE was reduced in AD males at 10 months, but not at 15 months. Compared to wildtype controls, the AD transgenic group of 15 months rats had significantly lower concentrations of esterified 13-oxo-ODE, 9-oxo-ODE and 9(10)-EpOME (decreased by 31%–40%) in brain phospholipids. The reported increases in HODEs further support the involvement of oxidative stress in AD pathology.⁷²⁴ The role of oxylipins in AD has been reviewed,^725,726 with suggestions that ALA-derived octadecanoids may mediate the observed beneficial effects of ALA in AD.⁷²⁶

In 2017, Hennebelle et al.⁷²⁷ showed the accumulation of LA-oxygenated metabolites in several rat brain regions during CO₂-induced ischemia. In particular 13-HODE was the most abundant metabolite in the hippocampus and its concentration increased 1.7-fold in cortex and brainstem following ischemia. 9-HODE and 13-oxo-ODE were also increased in cortex by 1.8-fold and 5.6-fold, respectively, in the ischemic CO₂-group compared to controls, whereas 13-oxo-ODE and 12,13-DiHOME were increased 3.2-fold and 1.4-fold, respectively in brainstem. Additionally 12(13)-EpOME and 9(10)-EpOME increased in both hippocampus (5.7-fold and 2.8-fold, respectively) and cerebellum (2.7-fold and 2.8-fold, respectively) compared to controls.⁷²⁷ Follow-up work in 2019 further reported that hypercapnia/ischemia increases brain oxylipin concentrations with observed increases in multiple LA-and ALA-derived octadecanoids.⁷²⁸ Interestingly Szczuko et al. in 2020 reported a significant decrease in the levels of 9- and 13-HODEs in the plasma of patients after ischemic stroke.⁷²⁹

Alteration in the levels of octadecanoids in brain has been used to predict the insurgence and severity of neurological disorders and age-related cognitive impairment. Swardfager and co-workers were able to correlate the ratios between 9,10-DiHOME and 9(10)-EpOME as well as between 12,13-DiHOME and 12(13)-EpOME (an index of sEH activity) with white matter hyperintensities (WMH) and proposed this value as a biomarker for vascular cognitive impairment in patients with WMH.⁷³⁰ The increased diols and reduced epoxides were related to increased sEH activity, which was reflected in increased levels of sEH-derived oxylipins in serum, implicating CYP/sEH-dependent metabolism in the etiology of WMH by injury to the periventricular subcortical white matter. A growing body of literature has supported this observation, and both plasma and cerebrospinal fluid levels of EpOMEs and DiHOMEs have proven to be valuable components of prediction models of neurological disorders. Increased CYP/sEH metabolites (as well as ethanolamides) were found to be strong predictors of AD with ROC curves ranging from 0.82 to 0.92 in CSF and plasma, respectively.⁷²² In 2020, Shinto et al. also showed that a higher ratio of 9,10-DiHOME/9(10)-EpOME is associated with increased WMH and poorer performance on the cognitive test Trails-B, and found a positive association between 9-HODE and WMH, and a negative association between 9-HODE and gray matter volume in nondemented people with controlled hypertension (mean age 65 ± 7.1 years), supporting the potential vasoconstrictive effects of 9-HODE.⁷³¹ Moreover, investigations into age related cognitive decline found a negative association between perceptual speed and the 12,13-DiHOME/12(13)-EpOME ratio after adjusting for the omega-3 diol and bile acid components.⁷³² More recently, Anita et al. reported that the 12(13)-EpOME was associated with poorer executive function and verbal memory scores in individuals with T2DM, while the 12,13-DiHOME was associated with lower executive function scores.⁷³³ Of interest, the authors reported an interaction between obesity and the ratio of 12,13-DiHOME/12(13)-EpOME, suggesting that BMI may play a role in the putative effects of these octadecanoids upon cognitive performance.

Multiple sclerosis is a severe inflammatory disorder resulting from an immunological attack on nerve fiber myelination throughout the body.⁷³⁴ Villoslada and colleagues identified 13-HODE as a biomarker of multiple sclerosis severity, its level being higher in serum samples of patient with active disease (associated with relapses and increase in EDSS, expanded disability status scale) compared to patient with a stable disease. (The paper refers to 13(S)-HODE although chirality was not measured in the analyses).⁷³⁵ Håkansson et al. found significantly higher levels of 9-HODE (average = 380 nM) and 13-HODE (average = 930 nM) in CSF of patients suffering from clinically isolated syndrome or relapsing remitting multiple sclerosis compared to healthy controls (average = 290 nM and 690 nM, respectively). However, the levels of the 9- and 13-HODE did not differ between patients with signs of disease activity during one, two and four years of follow-up and patients without, suggesting that increased HODE levels in patients may be an unspecific sign of neuroinflammation.⁷³⁶ Recently, it has been shown that the concentration of esterified 9(10)-EpOME was 206% higher in the neutral lipid pool of multiple sclerosis patients compared to controls. Because 9(10)-EpOME is known to be pro-inflammatory, increased esterification of this octadecanoid may reduce its availability as the free-form, resulting in decreased chronic inflammation associated with multiple sclerosis pathogenesis.⁷³⁷

Collectively, these findings suggest that there may be utility in monitoring octadecanoid levels in individuals with neurological disorders, but it is likely that the 9- and 13-HODE associations observed are due to nonspecific inflammation. The use of chiral-based methods will be important to delineate enzymatic vs. nonenzymatic formation of octadecanoids. Further research into other octadecanoid pathways including ALA would be of interest as well as investigations into other neurological diseases including Lewy body dementia and Parkinson’s. For example, the sEH pathway via epoxy-fatty acids has been proposed as a therapeutic target for neuropsychiatric disorders including Parkinson’s.⁷³⁸ While the literature suggests that alterations in these oxylipin pathways associate with neurological disorders, a comprehensive analysis of longitudinal changes in octadecanoid levels in relation to cognition is warranted, with particular focus on the putative role of ω6- vs. ω3-derived mediators.

11.2. Atherosclerosis

The analysis of oxidized linoleate in arterial atheromatous plaques dates from the early 1970s. C.J.W. Brooks and colleagues first noted the appearance of two classes of polar sterol esters in advanced aortic atheroma, one they identified as polar sterols esterified with normal fatty acids, and the second as cholesterol esterified with 9-HODE, 10(E),12(E)-9-HODE, or 13-HODE.⁷³⁹ They went on to show that the levels of oxidized linoleates were undetectable in “early” lesions (<1 μg/g of lipid), with an increase from 89 μg/g in fibrous plaques and atheromas, to 335 μg/g in more advanced lesions such as thrombotic or ulcerated plaques.⁷⁴⁰ Brooks further extended the analyses in two important ways: the sterol-HODE esters were shown to originate from cholesterol-linoleate hydroperoxides, and stereochemical analysis of the 13-HODE revealed it to be racemic.⁷⁴¹ The ratio 13(S)/13(R)-HODEs has been further investigated by Kühn et al., who determined that 15-LOX oxidation contributes to lipid peroxidation mainly in young human atherosclerotic lesions, where the ratio of 13(S)/13(R)-HODEs is (54 ± 3.2)/(45 ± 3.2). However, in more advanced human lesions, the ratio (50.7 ± 3.5)/(49.3 ± 3.5) is not significantly different to the one obtained by copper treated LDL, suggesting that most of the oxidized lipids of these lesions come from nonenzymatic oxidation.⁷⁴² The results pointed to lipid peroxidation as a factor in the development of atherosclerosis, a finding that has been followed up extensively in the subsequent decades.

By the 1990s, the oxidation of LDL became a major focus of research regarding its involvement in the initiation of atherosclerosis.^743,744 LA makes up approximately 40–45% of the PUFAs of low-density lipoproteins, which are the main constituent of atherosclerotic plaques. While further studies confirmed the findings of Brooks on the structures, relative abundance, and racemic stereochemistry of the oxidized linoleates in arterial plaques,^745,746 a key question was the possible involvement of a 15-LOX in initiating the oxidation of LDL and promoting the development of atherosclerosis.⁷⁴⁷ 15-LOX is capable of oxygenating LDL in vitro,⁷⁴⁸ and is implicated in the cellular oxidation of LDL.^749,750 This was examined using an in vivo model in which rabbits were fed a cholesterol-rich diet. The esterified 13-HODE in the initial arterial plaques exhibited 74% 13(S) chirality, which is evidence of oxygenation by 15-LOX. However, as the atheroma progressed, the 13-HODE became close to racemic, suggesting lipid peroxidation was predominant in the later stages of plaque development.⁷⁴⁷ Mouse knockout of the 12/15-LOX gene tends to support involvement of the enzyme in the early stages of atherogenesis, at least in animal models.⁷⁵¹ Further work in a rabbit hypercholesterolemia model reported that the 9- and 13-HODE were the most abundant quantified oxylipins in both plaques and plasma,⁷⁵² suggesting that these molecules would be translatable biomarkers of atherosclerosis. While the vast majority of studies have focused on the LA-derived HODEs, a recent review summarized the putative role of ALA-derived octadecanoids in cardiovascular diseases, suggesting potential immunomodulating effects.⁵⁷⁶ However, the reviewed studies were based upon mouse- and cell-based models and did not report utility as biomarkers of disease. In addition, Zhu et al. recently reported that the substitution of plant for animal protein in the diet was a promising strategy to modulate atherogenic lipids in a (apoE^–/–) mouse model.⁷⁵³ A 12-week high plant protein diet increased the abundance of microbiota from the Lachnospiraece family and resulted in a commensurate increase in circulatory levels of the 12,13-DiHOME, which was shown to inhibit lipid accumulation in vivo. The authors concluded that a high plant protein diet can alleviate hyperlipidemia via increased microbial production of the 12,13-DiHOME.

While there has been interest in the application of octadecanoids as indicators of atherosclerosis, the field has made few advances beyond the early identification of the 9- and 13-HODEs. This may be partially due to a paucity of studies investigating the octadecanoid content of the plaque in combination with screening for concomitant circulatory signatures. It would be of value to see studies performed in which the octadecanoid content of plaques was determined in conjunction with coronary angiography or other imaging techniques (e.g., CT angiography, ultrasound Doppler) to determine plaque composition and heterogeneity (e.g., calcified, necrotic core, fibroatheroma). This would enable determination of whether specific octadecanoid signatures, particularly as deposited cholesteryl esters, were associated with plaque pathology and/or stability, which could then be potentially linked with circulatory profiles to identify more accessible biomarkers.

11.3. Respiratory Diseases

There have been several investigations into the application of octadecanoids as an indicator of respiratory diseases. Most of these studies have concentrated on obstructive lung diseases (e.g., asthma, COPD) with a focus on the EpOMEs and DiHOMEs. The role of these compounds in the lung has been previously reviewed,⁷⁵⁴ with emphasis on the molecular and cellular mechanisms of EpOME-induced acute lung injury. An octadecanoid signature has been suggested to be a marker of the transition from healthy smokers with normal lung function to COPD, specifically driven by increases in EpOMEs, DiHOMEs, and TriHOMEs.⁷⁵⁵ Follow-up work on the TriHOMEs identified these compounds to be strongly increased in COPD;¹⁷² however, production was due to autoxidation likely in association with the oxidative stress in COPD patients. Levels of the 12,13-DiHOME were reported to increase in bronchoalveolar lavage fluid (BALF) of rats exposed to nitronaphthalene and ozone.⁷⁵⁶ Plasma levels of the 9,10-DiHOME and 9-HODE increased in healthy individuals following 1h exposure to biodiesel,⁷⁵⁷ while 12,13-DiHOME and 13-HODE were increased in BALF of healthy individuals exposed to biodiesel.⁷⁵⁸

The HODEs and oxo-ODEs have been reported to increase in several studies as primarily markers of oxidative stress. In a small pilot study, multiple octadecanoids including DiHOMEs, 13-HODE, 13-oxo-ODE, and 13-HOTrE were detected in the nasal epithelium of asthmatics and healthy controls. Only the 13-HOTrE evidenced a dysregulation, with decreases observed in asthmatics.⁷⁵⁹ The 9,10-DiHOME has been shown to correlate with lung function,⁷⁵⁵ and was reported to be at higher concentrations in circulation among a subgroup of asthmatics with the lowest lung function.⁷⁶⁰ Metabolomics analysis of sputum of asthmatics identified LA metabolism as the most significant pathway to discriminate between neutrophilic, eosinophilic and paucigranulocytic asthma, but did not identify any octadecanoid products.⁷⁶¹ Panda et al. reported that the 13-HODE partially leads to steroid-resistant asthma features through nuclear factor (NF)-κb.⁷⁶² Henricks et al. showed that 13-HODE (0.14 nmol) enhanced the increases in pulmonary resistance observed after administration of the contractile agents histamine or methacholine, in anesthetized, spontaneously breathing guinea pigs. These results indicate that 13-HODE may play an important role in the induction of airway hyperresponsiveness, a characteristic features of asthma, in vivo.⁷⁶³ It has been speculated that LA metabolism may have implications for the individualized treatment of neutrophilic asthma.

More recently, the DiHOMEs have been suggested to be biomarkers of COVID, with circulatory levels reported to associate with severe disease. A pilot study from Hammock and colleagues reported significant increases in the EpOMEs and DiHOMEs in plasma of 6 patients with laboratory-confirmed severe SARS-CoV-2 infection.⁷⁶⁴ Of the lipids analyzed, 18 had a > 4 fold-change and false discovery rate (p < 0.01), making a case for the “potential biomarker” claim in the title of the paper. The authors also noted that “incorporating diols in plasma multi-omics of patients could illuminate the COVID-19 pathological signature along with other lipid mediators and blood chemistry”. In a study of patients with varying COVID severity, 5 octadecanoids were elevated in plasma from intensive care unit patients, including 2–5-fold increases in both DiHOMEs as well as 12,13-DiHODE, while 13-HODE and 9-HOTrE increased 1–2-fold and there was surprisingly no change in 9-HODE levels.⁷⁶⁵ A study of individuals who had recovered from COVID reported decreased plasma levels of 13-oxo-ODE relative to individuals diagnosed with long COVID.⁷⁶⁶ There was no difference between healthy controls and individuals diagnosed with long COVID, indicating that 13-oxo-ODE would not be a useful biomarker of long COVID. This study explored potential mechanisms using a macrophage cell line to test the hypothesis that individuals with long COVID had alternatively polarized macrophages. They reported that the 12,13-DiHOME and a peak characterized as HpODEs were elevated in M2-like macrophages relative to M1 and concluded that system-wide alternative macrophage polarization is a key cell mechanism accounting for long COVID symptoms. A clinical trial treated hospitalized COVID patients with EPA for 3 days and found that the 9,10-DiHOME levels decreased relative to the placebo group.⁷⁶⁷ The uncontrolled inflammatory response in COVID-19 is characterized by a high neutrophil to lymphocyte ratio, which was decreased by EPA supplementation. Given that the 9,10-DiHOME is preferentially formed by neutrophils, there is potential for use of this compound as a biomarker for severe COVID. While promising, this clinical trial included a small number of patients (n = 22), who were of advanced age (81 ± 6.1 years). There is a need to perform additional studies in a broader cohort with inclusion of additional control groups to account for nonrespiratory viral infection. A recent study by Edin et al. examined oxylipins in a mouse model of COVID with human angiotensin-converting enzyme 2 (ACE2) expression.⁷⁶⁸ The study investigated the effect of sEH inhibition upon the host response to infection, focusing on the ensuring eicosanoid and cytokine storms. While circulatory levels of the DiHOMEs increased with infection and decreased with sEH inhibitor treatment, there was no overall effect on morbidity or mortality. A comprehensive multiomics study reported increases in the plasma levels of multiple LA-derived octadecanoids in adults with COVID-19 and in children with multisystem inflammatory syndrome in children (MIS-C) including the 12,13-DiHOME as well as 9-HpODE, 9-HODE and 13-KODE.⁷⁶⁹ Aside from COVID-19, there is less known about the role of octadecanoids in respiratory infections. The ratio of 13- to 9-HODE in bronchoalveolar lavage (BAL) was identified as a potential biomarker for immune status during an active influenza infection in a mouse model.⁷⁷⁰ The findings were replicated in nasopharyngeal lavage fluid of individuals with an active influenza infection, reporting an increased ratio in individuals with a high disease burden. The authors hypothesized that the ratio of 13-/9-HODE ratio reflected the balance of “anti-inflammatory” 13-HODE to “pro-inflammatory” 9-HODE, which is potentially an oversimplification of the inflammatory functions of these compounds.

The utility of octadecanoid profiles as biomarkers of respiratory diseases is promising. While early studies have demonstrated promise in circulatory profiles being associated with asthma severity and the transition from healthy smoker to a COPD smoker, additional investigations are needed to examine if these associations can be replicated in larger and more diverse cohorts. In particular, given the increasing focus on the role of the gut-lung axis in the etiology of respiratory diseases, emphasis should be placed on examining the utility of gut microbiome-derived octadecanoids as biomarkers of obstructive lung diseases. The association between COVID and circulatory DiHOME levels is of interest, and mechanistically plausible given the role of the DiHOMEs in ARDS and pulmonary damage. Further work should investigate the specificity of the DiHOME signature for COVID severity and particularly to determine if the levels decrease with the resolution of COVID symptoms.

11.4. Liver Diseases

There have been several investigations into the relationship between octadecanoid levels in liver diseases, with a focus on nonalcoholic fatty liver disease (NAFLD) and nonalcoholic steatohepatitis (NASH). Maciejewska et al. studied the role of 9-HODE in NAFLD in 24 patients (12 in the first stage and 12 in the second stage of NAFLD). Levels of HODEs were compared in plasma between the first and the second stage of hepatic steatosis. 9-HODE was present in higher concentrations in patients with grade II steatosis due to the greater exposure of liver cells to oxidative stress during the progression of the disease. After hepatic steatosis resolution by a six-month dietary intervention, a significant decrease in the concentrations of both 9- and 13-HODEs was observed.⁷⁷¹ In 2021, Mazi et al. reported elevations in a wide array of LA- and ALA-derived octadecanoids in NAFLD. Histology-matched Hispanic individuals had higher levels of TriHOMEs and EpOMEs than those of European ancestry, and DiHOME/EpOME ratios were important discriminators of ethnicity in those with full nonalcoholic steatohepatitis (i.e., NASH).⁷⁷² Kirpich and colleagues have published several studies on the interaction between alcohol-associated liver diseases and bioactive lipid mediators.⁷⁷³ Using a mouse model of alcoholic steatohepatitis, Warner et al. showed that the levels of 13-HODE, 9,10-DiHOME, and 12,13-DiHOME increased in plasma.⁷⁷⁴ A follow-up study examined the combined effects of ethanol and a diet high in LA (based upon corn oil) in the same mouse model, and reported that plasma and liver levels of 9- and 13-HODE increased in response to the combined feeding.⁶⁷⁶ The authors concluded that LA-derived octadecanoid induction of a pro-inflammatory response in macrophages is a potential mechanism driving the progression from alcohol-induced steatosis to alcoholic steatohepatitis. Of interest though, Liang et al. published that administering a high fat (corn oil) diet did not increase the liver concentration of octadecanoids (or other oxylipins), suggesting that these compounds do not accumulate in the liver.¹¹⁷ Further work in 2021 reported that the levels of 13-HODE and 13-oxo-ODE as well as 9,10- and 12,13-DiHOMEs were elevated in heavy drinkers suffering from moderate alcohol-associated hepatitis (mAH) compared to patients with mild alcohol-associated liver disease. However, 9(10)-EpOME and 12(13)-EpOME were decreased in heavy drinkers regardless of the presence or absence of liver injury.⁷⁷⁵

There have been several studies investigating associations between circulatory octadecanoid levels and different forms of liver diseases. In 2010, Thum et al. determined the levels of 9(10)-cis-EpODA in plasma of patients suffering from chronic liver diseases, a condition that often displays impaired liver CYP enzyme activities. 9(10)-cis-EpODA plasma concentrations were significantly repressed in patients with hepatic disease compared with healthy subjects. Thus, 9(10)-cis-EpODA was proposed as a biomarker to assess liver function.²⁰⁵ The profile of circulating lipid mediators has been characterized in individuals with acute decompensation of cirrhosis; 59 lipid mediators (out of 100 screened) were detected in plasma from cirrhotic patients and were significantly associated with disease severity. Among them, 11(E)-13-oxo-9(10)-EpOME was associated with short-term mortality and was a marker of coagulation and liver failure.⁷⁷⁶ Investigations in patients with hepatitis infection observed increases in circulatory octadecanoids, with Yoshida et al. showing in 2008 that the levels of total-HODEs (including 13-HODE, 9(E),11(E)-13-HODE, 9-HODE, 10(E),12(E)-9-HODE, 10-HODE, and 12-HODE) significantly increased in plasma and liver of patients infected with hepatitis B and C viruses.⁷⁷⁷ Later, in 2018, a study showed that 13-HODE, 9,10- and 12,13-DiHOME increased in patients with liver cirrhosis and hepatocellular carcinoma and the diols correlated with the levels of α-fetoprotein (AFP), a marker of hepatocellular carcinoma.⁷⁷⁸

There is a clear relationship between circulatory octadecanoid profiles and liver diseases, but liver levels do not appear to correlate with dietary input. Elevated octadecanoids have been suggested to have a causal role in NALFD and reported to be a biomarker, with circulatory levels being predictive of acute liver failure. Interestingly, some of these effects may occur via TRPV1, with deficiency protecting against experimental alcohol-associated liver disease. The sEH has been proposed as a therapeutic target in liver diseases, with therapeutic efficacy reported in NAFLD, liver fibrosis, and portal hypertension,⁷⁷⁹ potentially in combination with farnesoid X receptor (FXR) modulation.⁷⁸⁰ These findings collectively suggest that dietary octadecanoids are involved with liver diseases and merit further investigation to their utility as markers of both chronic and acute diseases, and represent a potential treatment route via reduction in dietary LA.

11.5. Inflammatory Pain

In 2018, Jensen et al. investigated the effect of chronic inflammatory pain on oxylipin concentrations in a mouse model of amygdala and periaqueductal gray (PAG) chronic inflammatory pain. Twelve LA-derived octadecanoids were detected in both the PAG and amygdala. Interestingly, two hydroxy-epoxide compounds known to be present in the skin of rodents and humans, 9(Z)-11-OH-12(13)-EpOME, and 11(E)-13-OH-9(10)-EpOME, were detected for the first time in brain tissue.⁶²⁷ When chronic inflammatory pain was induced by Complete Freund’s Adjuvant treatment, the concentration of the most detected compounds decreased in the amygdala, while the concentrations of 9(10)-EpOME, 12(13)-EpOME, and 9,10,11-TriHOME were reduced in PAG compared to the placebo controls.⁶²⁷ This octadecanoid reduction in mice suffering from chronic inflammatory pain is surprising, because an increase of concentrations of most octadecanoids (HODEs, oxo-ODEs, diols, keto-epoxides and hydroxy-epoxides) was previously observed in peripheral tissues, DRG, trigeminal ganglia (TG) and the dorsal horn of the spinal cord in animal models of inflammatory pain by lipidomic analyses.^61,222,617 Moreover, the levels of 9- and 13-HODEs as well as the related ketones increased in rodents suffering from heat and burn related pain of peripheral tissues.¹⁹⁵ The implications of these findings are as of yet unclear, but warrant further investigation into the specificity of octadecanoids as markers of inflammatory pain.

11.6. Sepsis

Lipid metabolism is closely associated with sepsis, with reported correlations with disease severity and systemic inflammation.^781−783 The role of oxylipins, particularly eicosanoids,^784,785 has been examined in sepsis with a number of reviews.^786−788 While the eicosanoids and SPMs have been investigated, there is significantly less known about octadecanoid involvement in this condition. Dalli et al. proposed that lipid mediator profiles from sepsis patients (n = 22) could serve as biomarkers of survival as well as development of ARDS.⁷⁸⁹ Sepsis nonsurvivors had greater plasma levels of inflammatory lipid mediators relative to sepsis survivors. However, no octadecanoids were included in these analyses. Bergmann et al. performed a study focusing on the DiHOMEs and EpOMEs in a burn-injured mouse model. They reported that DiHOME serum concentrations were significantly elevated, these increases could be ablated by administration of a sEH inhibitor, and that DiHOMEs rather than EpOMEs were the key driver of immune cell dysfunction through hyperinflammatory neutrophilic and impaired monocytic actions.⁶⁸ These findings are consistent with Hamaguchi et al., who reported that 9,10- and 12,13-DiHOME, as well as 9- and 13-HODE, were elevated in plasma from a single sepsis patient with a fatal Sequential Organ Failure Assessment (SOFA) score of 12.⁷⁹⁰ In a dog model, circulatory levels of the 9-HODE and 12,13-DiHOME were suggested as biomarkers of sepsis, with circulatory levels increasing 13.2- and 15.0-fold, respectively in sepsis.⁷⁹¹ Conversely, Sulaimin et al. reported in a study of sepsis patients (n = 274) that while AA-derived eicosanoid levels in plasma were elevated in poor outcome patients with sepsis compared to those with a more sustained and rapid recovery, there were no statistical differences in either 9- or 13-HODE levels.⁷⁹² While there is general consistency in the literature that octadecanoids increase in sepsis patients, further investigation is necessary, particularly in terms of establishing the mechanism. For example, it has been shown that phospholipase activity affects the observed eicosanoid signature in sepsis, with increased secretory phospholipase A2 group IIA (sPLA2-IIA) levels associating with eicosanoid metabolism in patients with bacterial sepsis syndrome.^793,794 Regardless, in the context of sepsis, monitoring changes in oxylipin profiles over time may provide an indication of the developing prognosis of a patient allowing adaptive therapeutic interventions.

11.7. Atopy

There have been multiple reports about linkages between dietary PUFA consumption and atopy, with reviews concluding that there is no relationship between atopy and exposure to ω6 PUFAs.⁷⁹⁵ Rucci et al. reported that in a population-based prospective cohort study of 4976 mothers in the second trimester, fatty acid levels in glycerophospholipids were associated with an increased risk of childhood eczema (1.21-fold) at age 6-years.⁷⁹⁶ The associations were primarily driven by LA, leading to the conclusion that higher maternal ω6 PUFA levels during pregnancy influence the risk of atopic diseases in childhood. While Rucci et al. did not observe a relationship with LA levels in the children and onset of atopy, the Ryukyus Child Health Study of 23 888 children in Japan (ages 6–15 years) identified a positive relationship between dietary LA and eczema (OR = 1.27).⁷⁹⁷ However, few studies have investigated the relationship between octadecanoids and allergic sensitization or atopy. Lundström et al. examined the effect of birch allergen provocation in asthmatics and found that multiple LA- and ALA-derived compounds were increased in the BALF relative to healthy individuals.⁷⁹⁸ While the majority of these compounds were increased in asthmatic controls relative to healthy controls, they further increased following provocation. In particular, the TriHOMEs, EpOMEs and DiHOMEs increased 1.7–2.6-fold. Nontargeted metabolomics showed that the LA-derived metabolites (13-HODE, 13-oxo-ODE) were significantly associated with moderate-to-severe atopic dermatitis in fecal samples from 6-month-old infants.⁷⁹⁹ The levels of 13-oxo-ODE were significantly reduced in the moderate-to-severe atopic dermatitis group compared to those in the healthy control and mild atopic dermatitis groups. In addition, 13-oxo-ODE negatively correlated with the SCORAD index (r = −0.595, p < 0.01), and 13-HODE negatively correlated with egg white–specific IgE at 12 months of age (r = −0.411, p = 0.016). Treatment of individuals with allergic rhinitis with either single- or double-species mite subcutaneous immunotherapy (SM-SCIT and DM-SCIT) for 36 weeks showed that the downstream products of LA metabolism (e.g., 9-HpODE, 13-HODE) decreased in serum; however, there was no significant difference between the SM-SCIT and DM-SCIT groups.⁸⁰⁰ These metabolites, as well as a number of eicosanoids (e.g., 11-HETE) were suggested as potential biological indicators for monitoring the desensitization effect on house dust mite (HDM) SCIT.

An elevated ratio in serum of LA to total fatty acids in 12-month-old infants was reported to be associated with onset of allergy in infants; however, the study employed an NMR method that did not measure the octadecanoids. The study included data from 438 infants, of which 48 had reported food allergy.⁸⁰¹ An earlier study by Yen and colleagues examined atopic dermatitis in children ages 2–17 years, with results reporting that atopic children had higher serum levels of LA, but lower levels of GLA and DGLA.⁸⁰² The authors concluded that the pathogenesis of atopic dermatitis is related to deficiency in ω6 essential fatty acids, which are required for normal skin barrier function and protection against inflammatory changes in the skin.⁸⁰² However, an earlier clinical trial in 102 subjects reported no effect of essential fatty acid supplementation upon atopic dermatitis.⁸⁰³ A pilot study in 20 atopic dermatitis patients reported that levels of 9,10,13-TriHOME in stratum corneum were elevated relative to healthy controls.⁸⁰⁴ This study used a tape-stripping method to sample the skin. Following correction with corneum total protein, the mean concentrations of total TriHOMEs in the atopic dermatitis group were 2.5- and 9.3-fold greater in the forehead skin and forearm skin, respectively, relative to normal controls. The study concluded that the noninvasive tape-stripping sampling may be useful for using TriHOME levels to monitor barrier function in atopic dermatitis. The antiallergic and anti-inflammatory effects of 12(Z)-10-HOME were investigated in a murine model of human atopic dermatitis. Addition of 12(Z)-10-HOME in the diet (0.01%, w/w) for 6 weeks decreased plasma IgE levels and skin infiltration of mast cells with a concomitant decrease in dermatitis score.⁸⁰⁵ While interesting, this study was relatively small in scope with 5 mice in each group, future work should expand the study and consider the role of parent LA in the mouse chow. Interestingly, an earlier study using a hairless mouse model of atopic dermatitis found that the development of atopic dermatitis-like symptoms was prevented by dietary supplementation with LA, but not with ALA.⁸⁰⁶ The study concluded that dietary-induced atopic dermatitis is mainly caused by deficiency of ω6 PUFAs, suggesting a potential therapeutic role of LA in atopic dermatitis.

Beyond the diet, environmental exposure is a significant octadecanoid source. While a number of groups have examined these effects, a limitation is that all of the reported environmental investigations presented here are cell based-studies. For example, pollen is rich in octadecanoids that have been demonstrated to exert a functional role in T2-mediated allergy. Plötz et al. examined the octadecanoid content of aqueous extracts of Phleum pratense L. (Timothy grass) and Betula alba L. (birch) pollen grains and measured the eosinophilic chemotaxic activity.⁸⁰⁷ Multiple LA-derived HODEs and ALA-derived HOTrEs were measured in the pollen, with chiral analyses demonstrating that the compounds were formed by autoxidation (racemic mixtures). The authors suggested that exposure to these lipid mediators may contribute to the elicitation and aggravation of eosinophil-associated inflammatory reactions by generating a T2-promoting local environment. Similar work from the group examined the biological activity of the same pollen fractions on polymorphonuclear granulocytes (PMNs), reporting that both LA- and ALA-derived monohydroxylated octadecanoids induced migratory responses as well as PMN activation.⁸⁰⁸ Traidl-Hoffmann and colleagues reported that the E₁-PhytoPs from birch pollen grains modulate human dendritic cell function in a fashion that favors T2 cell polarization in a fashion similar to PGE₂.⁸⁰⁹ Work by Mariani and colleagues identified the E₁-PhytoPs to increase the capacity of LPS-stimulated dendritic cells to attract T2 cells, whereas the capacity to recruit T1 cells was reduced.⁸¹⁰ Follow-up work concluded that the pollen-derived E₁-PhytoPs modulate dendritic cell function via PPARα dependent pathways that lead to T2 polarization.⁸¹¹ The ability of pollen-associated E₁-PhytoPs to impact cytokine secretion and maturation of dendritic cells was examined, with inhibition of IL-6 observed, but no effect upon LPS-induced surface expression of the maturation markers.⁸¹² More recently, the octadecanoid content of birch pollen was examined in more detail and reported multiple phytoprostanes as well as phytofurans.⁸¹³ Bet v 1 is the main allergen in birch pollen, and E₁-PhytoPs have been identified as novel ligands for this protein. These pollen-derived ligands enhance the proteolytic resistance of Bet v 1 and exhibit a dual role by stabilizing Bet v 1 and inhibiting cathepsin protease activity.⁸¹⁴ The presence of phytoprostanes and phytofurans has also been investigated in date palm, Phoenix dactylifera, edible parts and byproducts. The highest concentrations of phytoPs (11375 ± 2201 ng/100 g dry weight (dw)) have been reported in pollen, and PhytoFs concentration is elevated in skin (329.15 ± 70.19 ng/100 g dw) and pollen (188.41 ± 28.53 ng/100 g dw).⁸¹⁵ In addition, no HODEs were detected contrary to earlier reports.⁸⁰⁸

Given the hypothesis that dietary PUFAs are associated with atopy, there is a need to further explore the relationship between octadecanoids and onset of allergic sensitization. The role of octadecanoid lipid mediators in pollen in allergy has been previously reviewed.^816,817 The data to date are varied, rendering it difficult to make a definitive assessment of the relationship. Further studies will need to address the timing of the dietary exposure (maternal vs. child exposure), the route of exposure (inhalation, dietary, contact), and the dose/response nature of the exposure as well as the putative biological activity of the parent PUFA relative to the octadecanoid metabolite.

11.8. Diuresis/Antidiuresis

Urinary levels of DiHOMEs and TriHOMEs are influenced by salt loading and depletion. Dreisbach et al. showed that intravenous salt loading increased, and salt depletion with oral furosemide decreased, the urinary excretion of these linoleate metabolites measured after glucuronidase treatment.⁸¹⁸ Arachidonic acid epoxides are known to reduce the residence time of the furosemide sensitive Na⁺/K⁺/Cl^– cotransporter (NKCC) in the renal proximal tubule epithelial sodium channel to promote sodium clearance.^819,820 In addition, arginine vasopressin system activation increases soluble epoxide hydrolase activity, shifting tissue DiHOME/EpOME ratios.⁸²¹ Therefore, urinary levels of DiHOMEs, and possibly TriHOMEs, appear to be reporters of diuresis/antidiuresis processes in the kidney. This is of particular interest since such urinary octadecanoid profiles could complement classic measures of renal tubular function like electrolyte levels, osmolarity, or a furosemide stress test^822,823 by providing measures of the kidney’s adaptive ability to respond to a physiological demand, rather than simply measuring the outcome of the response.

11.9. Type 2 Diabetes Mellitus and Metabolic Syndrome

In 2012, Grapov et al. demonstrated that diabetes was associated with increases in plasma EpOMEs and EpODEs in weight-matched obese women, with the 9(10)-EpODE/12(13)-EpODE ratio being an important indicator of diabetes status.⁸²⁴ In 2013, Umeno et al. reported that the singlet oxygen products 10- and 12-HODEs are potential biomarkers of early Type 2 diabetes melitus.⁸²⁵ Specifically, they reported that after saponification, fasting plasma levels of the 10- and 12-HODEs/LA ratio increased with other clinical markers of diabetes and could enable the identification of borderline diabetes without the use of an oral glucose tolerance test. More recently, esterified oxylipin profiling was found to provide a powerful discriminate model of metabolic syndrome, with shifts in a host of HODEs, oxo-OMEs, DiHOMEs, EpOMEs, and EpODEs detected.⁸²⁶ In 2022, Fedorova and colleagues published an elegant workflow to analyze the levels of esterified oxidized lipids in plasma in lean, obese, and Type 2 diabetic people. The results evidenced significant differences between groups of samples for 20 oxidized phosphatidylcholines, 26 oxidized triglycerides and 26 oxidized cholesteryl esters. Of these 72 lipids, 56 contained octadecanoids. Interestingly, 25 species were significantly increased in lean individuals. All of the species contained octadecanoids in their side chains.⁸²⁷ Therefore, octadecanoid profiling in the context of diabetes and cardiometabolic syndrome may provide insights into disease progression. A particularly relevant point from this work is the focus on the esterified species, demonstrating the importance of not only focusing on the nonesterified oxylipin pool.³⁸⁰ Hateley et al. investigated oxylipins in the omental WAT, liver biopsies and plasma of patients undergoing bariatric surgery. They reported that increases in the sEH activity index of 12(13)-EpOME:12,13-DiHOME in WAT and liver were a marker of worsening metabolic syndrome in patients with obesity.⁸²⁸

12. Conclusion

The formation of octadecanoids in mammalian systems has been known for decades, but often overlooked relative to the eicosanoids and more recently the docosanoids. The longer chain PUFAs have understandably taken the lead due to their potency, exquisite regulation, and demonstrated actions in diverse systems. On the other hand, the octadecanoids have often been dismissed as simply oxidized fat and a marker of lipid oxidation. However, as our understanding of regulatory lipid metabolism has been enriched from decades of research, interest in this class of molecules has begun to increase as we recognize that many octadecanoids are in fact potent contextual lipid mediators. While there has been a long-standing question as to whether the octadecanoids are bioactive, it is now clear that numerous members of this class are indeed potent lipid mediators. In fact, some octadecanoids are considered to be lipokines or serve as epigenetic modifiers. The expanding interest in these compounds necessitates a concomitant response from the scientific community to develop the necessary tools to investigate octadecanoid biology. There is to date a paucity of analytical methods and chemical standards from the octadecanoid field, but this is changing and should result in a concomitant increase in studies investigating octadecanoids. We also see a need to standardize the nomenclature and reporting of these compounds and to work with repositories such as LIPID MAPS and HMDB to develop our understanding of their endogenous concentrations and tissue distributions. It is of particular importance to ascertain which aspects of octadecanoid biology are well established (i.e., reproduced) vs. proposed, especially in the context of describing the biology, biochemistry and signaling actions of these compounds. Ultimately, many important challenges remain in the octadecanoid field.

As is often the case with lipid mediators, there is little knowledge with regards to specific octadecanoid receptors. There is a need to perform much of the basic biochemical characterization of these compounds that has been done for other fatty acid cascades. We need to determine which cell types produce which octadecanoids under which conditions. There is a necessity to determine the range of their production, specifically in terms of triggers of biosynthesis (e.g., calcium ionophore, anti-IgE, and LPS) as well as the effects of cytokines and growth factors that affect the fidelity of the system (e.g., alarmins, IL-25, IL-33). As a component of understanding the mechanisms of formation, the effects of pharmacological interventions with enzyme (e.g., COX, LOX, CYP, sEH) inhibitors on lipid mediator formation should be examined. These efforts need to be augmented with investigations of the putative biological activity, which remains unknown for the vast majority of octadecanoids. In particular, identifying specific receptors will be key in developing an understanding of their biology. A comprehensive evaluation of octadecanoids as effectors of the entire TRP superfamily would be extremely valuable and may unravel potential mechanisms of action in various cell systems.

A limitation in the field is that the majority of studies to date have focused on the canonical LA-derived octadecanoids (e.g., HODEs, EpOMEs, DiHOMEs). In particular, there is a need to investigate the bacterial and fungal derived octadecanoids which constitute a rich source of unique structures whose biological activity has not been investigated. In addition, the low abundant and lesser-known octadecanoids should be characterized. For example, SDA is likely absent or present at very low concentrations in the general population; however, this lipid is present in the food supply (as well as in dietary supplements). The oxidative metabolism of this fatty acid has not been investigated, likely due to a lack of analytical standards and the general emphasis on more abundant precursors. As awareness increases of the richness of the full octadecanoid cascade from multiple C18-FAs, there is a need to cast a broader net and investigate these other octadecanoids, with plant species rich in the parent oils being an appropriate target to begin such an exploration. There is the potential that even if some compounds are not present endogenously, they may possess pharmacologic activity that could become relevant for individuals taking supplements and may even add value to some commodity crops. This work needs to include analytical characterization to establish endogenous levels under varying conditions as well as to characterize potential pharmacological activity. The application of chiral analyses will be useful to determine formation mechanisms. This is particularly relevant given that the majority of proposed octadecanoid biomarkers to date are HODEs, which readily form from oxidation of LA. While we have summarized multiple studies reporting octadecanoids as putative biomarkers of various diseases and pathologies, assigning mechanisms associated with the reported associations will be challenging. The most common associations reported are with the LA-derived 9- and 13-HODE, and may simply reflect shifts in systemic oxidation stress associated with general disease pathology rather than specific disease mechanisms. Demonstrating the enantiomeric ratios of formation will add specificity to any proposed biomarker and immediately enhance our ability to interpret the implications of the associated changes. With the advent of new technologies including gas-phase chiral separations by ion mobility mass spectrometry, we believe that research in this area over the next decade could revolutionize our understanding of the role of octadecanoids in complex biological systems.

While only addressed briefly in this review, it is relevant to highlight that the vast majority of fatty acids, including the C18 octadecanoid precursors, are esterified and that the octadecanoids themselves primarily exist as esterified products. The free forms are presumably the biologically active oxylipins, but the functions of those found esterified within phospholipids are not known and should not be ignored. It has been demonstrated that HODE incorporation into inositol phosphates proceeds the formation of HODE-diacyl glycerols and that they have participated in intercellular signaling cascades.⁸²⁹ It is also a strong possibility that they could alter membrane properties, thereby influencing the function of membrane proteins, and/or act as a storage reservoir participating in the Lands cycle of lipolytic release and re-esterification as demonstrated for eicosanoids.

There is also a need to characterize the octadecanoid component of different food oils and high fat food stuffs particularly under different storage and preparation conditions. It is likely that these greatly affect dietary octadecanoids levels. In addition, when examining the biological activity of octadecanoids, it is important to place their dietary and subsequent endogenous level within context. Eicosanoid lipid mediators have been demonstrated to exert potent biological activity at low concentrations. However, levels of AA are estimated to be 0.051% of total energy intake vs. 0.72% for ALA and 7.2% for LA in the United States.⁵¹ Octadecanoids are subsequently likely to be present at endogenous concentrations that are orders of magnitude greater than eicosanoids.

The field of octadecanoid research is promising; however, there is much work required to adequately characterize this class of lipids in systems that directly impact human biology, including both host and microbiota biochemistry. Only through the concerted efforts of the lipid community will we accumulate sufficient knowledge regarding these compounds to establish their rightful place alongside their more well-known cousins, and establish the importance of octadecanoids to animals, as has been so well established in plants.^830−833

Acknowledgments

We thank Jeff Johnson for multiple discussions on the bioactivity of octadecanoids and in-depth reading of this manuscript as well as three anonymous reviewers who provided helpful feedback that significantly improved the paper. J.W.N. acknowledges support provided by USDA project 2032-51530-025-00D. The USDA is an equal opportunity provider and employer. A.R.B. acknowledges support by NIH grants RO1-GM134548 and R35-GM152031, and the content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. C.E.W. acknowledges support from the Swedish Heart Lung Foundation (HLF 20230463 and HLF 20210519) and the Swedish Research Council (2022-00796).

Glossary

Abbreviations

AA: arachidonic acid
ACSL: acyl-CoA synthase
AD: Alzheimer’s disease
AdA: adrenic acid
ADP: adenosine diphosphate
AFP: α-fetoprotein
AKT: A serine/threonine protein kinase
ALA: α-linolenic acid
AMPP: N-(4-aminomethylphenyl)pyrimidium
ANT1: adenine nucleotide translocase 1
AOS: allene oxide synthase
ARDS: acute respiratory distress syndrome
ATP: adenosine 5′-triphosphate
BALF: bronchoalveolar lavage fluid
BAT: brown adipose tissue
CCS: collision cross section
CFA: complete Freund’s adjuvant
CGRP: calcitonin gene-related peptide
CLA: conjugated linoleic acid
CLA-DC: CLA-decarboxylase
CLA-DH: CLA-dehydrogenase
CLA-ER: CLA-enoyl reductase
CLA-HY: CLA-hydrolase
CLnA: conjugated linolenic acid
COPD: chronic obstructive pulmonary disease
COX: cyclooxygenase
CSF: cerebrospinal fluid
CYP: cytochrome P
Cys: cysteine
DESI: desorption electrospray ionization
DGLA: dihomo γ-linolenic acid
DHA: docosahexaenoic acid
DiHODA: dihydroxy-octadecanoic acid
DiHODE: dihydroxy-octadecadienoic acid
DiHOME: dihydroxy-octadecenoic acid
DiHOTE: dihydroxy-octadecatetraenoic acid
DiHOTrE: dihydroxy-octadecatrienoic acid
DMS: differencial mobility spectrometry
DM-SCIT: double-species mite subcutaneous immunotherapy
DNA: deoxyribonucleic acid
DOX: dioxygenase
DPA: docosapentaenoic acid
DRG: dorsal root ganglia
DSM-IV: Diagnostic and Statistical Manual of Mental Disorders IV
DSS: dextran sulfate sodium
dw: dry weight
EAS: epoxy alcohol synthase
ECD: electron capture detector
EDSS: expanded disability status scale
EET: epoxy-eicosatrienoic acid
EFA: essential fatty acid
EKODE: epoxy-keto-octadecadienoic acid
EKOME: epoxy-keto-octadecenoic acid
ELISA: enzyme linked immunoassay
ELOVL: elongation of very long chain fatty acids
eLOX: epidermal lipoxygenase
EOS: esterified omega-hydroxy sphingosine
Ep: epoxide
EPA: eicosapentaenoic acid
EPHX3: epoxide hydrolase EH3
EpODA: epoxy-octadecanoic acid
EpODE: epoxy-octadecadienoic acid
EpOME: epoxy-octadecenoic acid
EpOTrE: epoxy-octadecatrienoic acid
ESI: electron spray ionization
ETA: 8,11,14,17-eicosatetraenoic acid
FA: fatty acid
FABP4: fatty acid binding protein 4
FAD: flavin adenine dinucleotide
FADS: fatty acid desaturases
FA-HY: fatty acid hydratase
Fg-cat: fungal catalase
FID: flame ionization detector
fMLP: formyl-methionyl-leucylphenylalanine
FMN: flavin mononucleotide
FXR: farnesoid X receptor
GC: gas chromatography
GLA: γ-linolenic acid
GLP-1: glucagon-like peptide-1
GM-CSF: granulocyte-macrophage colony-stimulating factor
GPCR: G protein-coupled receptor
HDM: house dust mite
HETE: hydroxy-eicosatetraenoic acid
HFD: high fat diet
HODA: hydroxy-octadecanoic acid
HODE: hydroxy-octadecadienoic acid
HOME: hydroxy-octadecenoic acid
HOTE: hydroxy-octadecatetraenoic acid
HOTrE: hydroxy-octadecatrienoic acid
Hp: hydroperoxide
HpETE: hydroperoxy-eicosatetraenoic acid
HPLC: high-performance liquid chromatography
HpODA: hydroperoxy-octadecanoic acid
HpODE: hydroperoxy-octadecadienoic acid
HpOME: hydroperoxy-octadecenoic acid
HpOTE: hydroperoxy-octadecatetraenoic acid
HpOTrE: hydroperoxy-octadecatrienoic acid
HRMS: high resolution mass spectrometry
HWE: Horner–Wadsworth–Emmons reaction
IAC: immunoaffinity chromatography
IL: interleukin
IMS: ion mobility spectrometry
IsoF: isofuran
IsoP: isoprostane
JNK: c-Jun N-terminal kinase
Keap: Kelch-like ECH-associated protein
KETE: keto-eicosatetraenoic acid
LA: linoleic acid
LC-MS/MS: liquid chromatography tandem-mass spectrometry
LC-PUFA: long chain PUFA
LDL: low-density lipoprotein
LDS: linoleate diol synthase
LLOD: low limit of detection
LLOQ: low limit of quantification
LOX: lipoxygenase
LPS: lipopolysaccharide
LTB₄: leukotriene B4
MA: Mead acid
mAH: moderate alcohol-associated hepatitis
MCP-1: monocyte chemoattractant protein-1
mDC: matured dendritic cell
MRM: multiple reaction monitoring
MSI: mass spectrometry imaging
MUFA: monounsaturated fatty acid
MW: microwave
NAD: nicotinamide adenine dinucleotide
NADPH: nicotinamide adenine dinucleotide phosphate
NAFLD: nonalcoholic fatty liver disease
NASH: nonalcoholic steatohepatitis
NF-E2: nuclear factor erythroid 2
NKCC: Na⁺/K⁺/Cl^– cotransporter
NLRP3: NLR family pyrin domain containing 3
NMR: nuclear magnetic resonance
NO₂-FA: nitrated fatty acids
NODE: nitro-octadecadienoic acid
NOME: nitro-octadecenoic acid
NOTrE: nitro-octadecatrienoic acid
NRF2: nuclear factor erythroid 2-related factor-2
OA: oleic acid
ODA: octadecanoic acid
ODE: octadecadienoic acid
OME: octadecenoic acid
OPDA: 12-oxo-phytodienoic acid
OTE: octadecatetraenoic acid
OTrE: octadecatrienoic acid
PAG: periaqueductal gray
PCET: proton-coupled electron transfer
PG: prostaglandin
PhytoF: phytofuran
PhytoP: phytoprostane
PLA₂: phospholipase A2
PLS: partial least-squares
PMN: polymorphonuclear granulocyte
PPAR: peroxisome proliferator-activated receptor
PUFA: polyunsaturated fatty acid
RIA: radio immunoassay
RNS: reactive nitrogen species
ROS: reactive oxygen species
SDA: stearidonic acid
sEH: soluble epoxide hydrolase
SM-SCIT: single-species mite subcutaneous immunotherapy
SPM: specialized pro-resolving mediator
T2D: type-2 diabetes
TG: trigeminal ganglia
THFA: tetrahydrofuranyl fatty acids
TLC: thin layer chromatography
TLR4: toll-like receptor 4
TNF-α: tumor necrosis factor alpha
TPP: 5,10,15,20-tetraphenyl-21H,23H-porphyrin
TPPU: 1-(1-propanoylpiperidin-4-yl)-3-[4-(trifluoromethoxy)phenyl]urea
Treg: regulatory T cell
TriHODA: trihydroxy-octadecanoic acid
TriHODE: trihydroxy-octadecadienoic acid
TriHOME: trihydroxy-octadecenoic acid
TriHOTrE: trihydroxy-octadecatrienoic acid
TRPV: transient receptor potential vanilloid
Tyr: tyrosine
UPLC: uultraperformance liquid chromatography
UV: ultraviolet
WMH: white matter hyperintensities

Biographies

Johanna Revol-Cavalier is a staff chemist at Larodan AB in Stockholm (Sweden). She holds a Bachelor’s degree and a Master’s degree from the University of Montpellier (France). From 2017 to 2020, she performed her doctoral research at the University of Montpellier with Jean-Marie Galano and Camille Oger, where she developed multistep synthetic strategies of cyclic, nonenzymatic metabolites of polyunsaturated fatty acids. From 2020 to 2022, she worked as a Postdoctoral Researcher in the laboratory of Craig Wheelock at the Karolinska Institute (Stockholm, Sweden), where her research focused on the development of synthetic routes to obtain various oxidized metabolites of C18 polyunsaturated fatty acids.

Alessandro Quaranta is a bioanalytical scientist at Menarini Ricerche in Pomezia (Italy). He holds a Bachelor’s degree in Chemistry and a Master’s degree in Analytical Chemistry from Sapienza University in Rome (Italy). From 2014 to 2019, he performed his doctoral research at Stockholm University with Gunnar Thorsén and Leopold L. Ilag, where he developed analytical methods for the comprehensive characterization of protein glycosylation and investigated the alteration of post-translational modifications as a consequence of multiple diseases. From 2019 to 2023, he joined Craig Wheelock’s group at the Karolinska Institute (Stockholm, Sweden) as a Postdoctoral Researcher. There he developed the analytical platform for the characterization of octadecanoids and applied it to investigate the role of these compounds in the mediation of inflammation and in respiratory diseases.

John W. Newman has been a Research Chemist with the United States Department of Agriculture, Agricultural Research Service, at the Western Human Nutrition Research Center in Davis, California, since 2005. He is also an Adjunct Professor with the University of California Davis, Department of Nutrition, and Director of the Lipid Mediators Research Laboratory of the UC Davis West Coast Metabolomics Center. After receiving his Bachelor’s degree from the University of California Santa Cruz with research focused on the analysis, fate, and transport of trace contaminants in the marine ecosystems, he obtained a Ph.D. in Pharmacology and Toxicology from the University of California Davis, where he focused on the development and application of analytical tools for the study of the soluble epoxide hydrolase. Among many accomplishments, he codiscovered and reported the “vestigial” catalytic domain of the soluble epoxide hydrolase to be a functional lipid phosphatase. He is an expert in analytical chemistry, lipid metabolism, and metabolomics, with >30 years of experience and >200 peer reviewed manuscripts. He has broad domestic and international collaborations in human nutrition, emphasizing the impact of diet and dietary components on nutritionally sensitive disease risk mediated through bioactive lipids and other endogenous small molecules. He continues to explore lipoprotein-dependent bioactive lipid transport, and more recently, his research has turned to nutritional phenotyping, exploring the variability in metabolomic responses to dietary interventions.

Alan R. Brash studied medical sciences at Downing College, Cambridge University, and received his Ph.D. from the University of Edinburgh on the analysis of prostaglandins and their metabolites. After a postdoctoral fellowship at the Royal Postgraduate Medical School, he moved to Vanderbilt University in the USA, where he rose through the ranks to full Professor in 1989. From initial analytical work on the quantitative analysis of prostaglandin and prostacyclin metabolites in humans and the effects of anti-inflammatory drugs, the focus moved on to the mechanisms of leukotriene biosynthesis and mechanistic aspects of lipoxygenase catalysis. He later developed a particular interest in the rare occurrence of lipoxygenase enzymes producing the mirror image R-configuration products and in the mechanistic basis underlying this unusual catalysis, a theme he has developed throughout his career. He continues to pursue the enzymology of lipoxygenases and especially the role of 12R-lipoxygenase and related enzymes in formation of the mammalian skin permeability barrier. In 2013, he was elected as an AAAS Fellow ”as one of the world’s leading authorities in the field of the biosynthesis of prostaglandins and related eicosanoids”.

Mats Hamberg is a senior professor in the Institute of Environmental Medicine, Karolinska Institute, Stockholm (Sweden). After retirement, he joined Larodan AB in Stockholm, where he now works as a senior chemist. He has published >350 papers, most of them dealing with fatty acid dioxygenases and their products. Highly cited papers include those from the 1970s–1980s describing the discovery of prostaglandin endoperoxides, thromboxanes, lipoxins, and the animal 12- and 5-lipoxygenases. Subsequent work was focused on lipoxygenases and related enzymes in higher plants and include discovery of the enzyme allene oxide cyclase required for jasmonate biosynthesis. Together with co-workers, he discovered the first manganese-dependent lipoxygenase and the enzyme α-dioxygenase involved in plant α-oxidation. Notable compounds encountered during these studies include allene oxides, α-hydroperoxides, epoxy alcohols, as well as a number of new plant natural products.

Craig Wheelock is Head of the Unit of Integrative Metabolomics in the Institute of Environmental Medicine at the Karolinska Institute, where he also leads the Integrative Molecular Phenotyping Laboratory and serves as Director of the Small Molecule Mass Spectrometry core facility (KI-SMMS). He is a Distinguished Visiting Professor of Metabolomics at Gunma University, Japan, a Fellow of the European Respiratory Society (FERS), and a member of the Swedish Fulbright Commission. He served on the Board of Directors for the International Metabolomics Society (2016–2020) as well as the Nordic Metabolomics Society (2017–2021). Following pre- and postdoctoral work on lipid mediators at the University of California Davis with Bruce Hammock, he conducted postdoctoral studies at the KEGG laboratory in Kyoto University. In 2006, he was awarded a Marie Curie Fellowship to relocate to the Karolinska Institute where his research has focused on oxylipins. He has published >250 papers dealing with a diverse range of topics centering on the role of bioactive lipids in inflammation. Research in his laboratory focuses on molecular phenotyping of respiratory diseases, with a particular interest in investigating the role of lipid mediators in disease pathophysiology.

The authors declare no competing financial interest.

Dedication

This work is dedicated to Dr. Bruce Hammock, whose pioneering work in leukotoxins set the stage for this report of the octadecanoids.

References

Chevreul M. E.; de Bouffe J.. Ou recueil des mémoires concernant la chimie et les arts qui en dépendent. Annales de Chimie 1813.881 [Google Scholar]
Burdge G. C.; Calder P. C.. Introduction to Fatty Acids and Lipids. Intravenous Lipid Emulsions; World Review of Nutrition and Dietetics; Karger, 2015; Vol. 112, pp 1–16. 10.1159/000365423 [DOI] [PubMed] [Google Scholar]
Benatti P.; Peluso G.; Nicolai R.; Calvani M. Polyunsaturated Fatty Acids: Biochemical, Nutritional and Epigenetic Properties. J. Am. Coll. Nutr. 2004, 23 (4), 281–302. 10.1080/07315724.2004.10719371. [DOI] [PubMed] [Google Scholar]
Vance D. E.; Vance J. E.. Biochemistry of Lipids, Lipoproteins and Membranes; Elsevier Science, 2008. [Google Scholar]
Burr G. O.; Burr M. M.; Miller E. S. On the Fatty Acids Essential in Nutrition. III. J. Biol. Chem. 1932, 97, 1–9. 10.1016/S0021-9258(18)76213-3. [DOI] [Google Scholar]
Tocher D. R.; Leaver M. J.; Hodgson P. A. Recent Advances in the Biochemistry and Molecular Biology of Fatty Acyl Desaturases. Prog. Lipid Res. 1998, 37 (2), 73–117. 10.1016/S0163-7827(98)00005-8. [DOI] [PubMed] [Google Scholar]
Holman R. T.; Mohrhauer H.; Porwit-Bobr Z.; Baptist J. N. A Hypothesis Involving Competitive Inhibitions in the Metabolism of Polyunsaturated Fatty Acids. Acta Chemica Scandinavia. 1963, S84–S90. 10.3891/acta.chem.scand.17s-0084. [DOI] [Google Scholar]
Nakamura M. T.; Nara T. Y. Essential Fatty Acid Synthesis and Its Regulation in Mammals. Prostaglandins Leukot. Essent. Fatty Acids 2003, 68 (2), 145–150. 10.1016/S0952-3278(02)00264-8. [DOI] [PubMed] [Google Scholar]
Cormier H.; Rudkowska I.; Lemieux S.; Couture P.; Julien P.; Vohl M.-C. Effects of FADS and ELOVL Polymorphisms on Indexes of Desaturase and Elongase Activities: Results from a Pre-Post Fish Oil Supplementation. Genes Nutr. 2014, 9 (6), 437. 10.1007/s12263-014-0437-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
Glick N. R.; Fischer M. H. The Role of Essential Fatty Acids in Human Health. J. Evid.-Based Complement. Altern. Med. 2013, 18 (4), 268–289. 10.1177/2156587213488788. [DOI] [Google Scholar]
Gerwick W. H.; Moghaddam M.; Hamberg M. Oxylipin Metabolism in the Red Alga Gracilariopsis lemaneiformis: Mechanism of Formation of Vicinal Dihydroxy Fatty Acids. Arch. Biochem. Biophys. 1991, 290 (2), 436–444. 10.1016/0003-9861(91)90563-X. [DOI] [PubMed] [Google Scholar]
Dyall S. C.; Balas L.; Bazan N. G.; Brenna J. T.; Chiang N.; da Costa Souza F.; Dalli J.; Durand T.; Galano J.-M.; Lein P. J.; Serhan C. N.; Taha A. Y. Polyunsaturated Fatty Acids and Fatty Acid-Derived Lipid Mediators: Recent Advances in the Understanding of Their Biosynthesis, Structures, and Functions. Prog. Lipid Res. 2022, 86, 101165. 10.1016/j.plipres.2022.101165. [DOI] [PMC free article] [PubMed] [Google Scholar]
Harizi H.; Corcuff J.-B.; Gualde N. Arachidonic-Acid-Derived Eicosanoids: Roles in Biology and Immunopathology. Trends Mol. Med. 2008, 14 (10), 461–469. 10.1016/j.molmed.2008.08.005. [DOI] [PubMed] [Google Scholar]
Calder P. C. Eicosanoids. Essays Biochem. 2020, 64 (3), 423–441. 10.1042/EBC20190083. [DOI] [PubMed] [Google Scholar]
Samuelsson B.; Dahlén S.-E.; Lindgren J. Å.; Rouzer C. A.; Serhan C. N. Leukotrienes and Lipoxins: Structures, Biosynthesis, and Biological Effects. Science 1987, 237 (4819), 1171–1176. 10.1126/science.2820055. [DOI] [PubMed] [Google Scholar]
Tourdot B.; Ahmed I.; Holinstat M. The Emerging Role of Oxylipins in Thrombosis and Diabetes. Front. Pharmacol. 2014, 4, 176. 10.3389/fphar.2013.00176. [DOI] [PMC free article] [PubMed] [Google Scholar]
Misheva M.; Johnson J.; McCullagh J. Role of Oxylipins in the Inflammatory-Related Diseases NAFLD, Obesity, and Type 2 Diabetes. Metabolites 2022, 12 (12), 1238. 10.3390/metabo12121238. [DOI] [PMC free article] [PubMed] [Google Scholar]
Barquissau V.; Ghandour R. A.; Ailhaud G.; Klingenspor M.; Langin D.; Amri E.-Z.; Pisani D. F. Control of Adipogenesis by Oxylipins, GPCRs and PPARs. Biochimie 2017, 136, 3–11. 10.1016/j.biochi.2016.12.012. [DOI] [PubMed] [Google Scholar]
Liput K. P.; Lepczyński A.; Ogłuszka M.; Nawrocka A.; Poławska E.; Grzesiak A.; Ślaska B.; Pareek C. S.; Czarnik U.; Pierzchała M. Effects of Dietary n-3 and n-6 Polyunsaturated Fatty Acids in Inflammation and Cancerogenesis. Int. J. Mol. Sci. 2021, 22 (13), 6965. 10.3390/ijms22136965. [DOI] [PMC free article] [PubMed] [Google Scholar]
Edin M. L.; Duval C.; Zhang G.; Zeldin D. C. Role of Linoleic Acid-Derived Oxylipins in Cancer. Cancer Metastasis Rev. 2020, 39 (3), 581–582. 10.1007/s10555-020-09904-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lundstrom S. L.; Balgoma D.; Wheelock A. M.; Haeggstrom J. Z.; Dahlen S.-E.; Wheelock C. E. Lipid Mediator Profiling in Pulmonary Disease. Curr. Pharm. Biotechnol. 2011, 12, 1026–1052. 10.2174/138920111795909087. [DOI] [PubMed] [Google Scholar]
Swardfager W.; Hennebelle M.; Yu D.; Hammock B.; Levitt A.; Hashimoto K.; Taha A. Metabolic/Inflammatory/Vascular Comorbidity in Psychiatric Disorders; Soluble Epoxide Hydrolase (SEH) as a Possible New Target. Neurosci. Biobehav. Rev. 2018, 87, 56–66. 10.1016/j.neubiorev.2018.01.010. [DOI] [PMC free article] [PubMed] [Google Scholar]
Duan J.; Song Y.; Zhang X.; Wang C. Effect of ω-3 Polyunsaturated Fatty Acids-Derived Bioactive Lipids on Metabolic Disorders. Front. Physiol. 2021, 12, 646491. 10.3389/fphys.2021.646491. [DOI] [PMC free article] [PubMed] [Google Scholar]
Guimarães R. C.; Gonçalves T. T.; Leiria L. O. Exploiting Oxidized Lipids and the Lipid-Binding GPCRs against Cardiometabolic Diseases. Br. J. Pharmacol. 2021, 178 (3), 531–549. 10.1111/bph.15321. [DOI] [PubMed] [Google Scholar]
Nayeem M. A. Role of Oxylipins in Cardiovascular Diseases. Acta Pharmacol. Sin. 2018, 39 (7), 1142–1154. 10.1038/aps.2018.24. [DOI] [PMC free article] [PubMed] [Google Scholar]
He J.; Ma C.; Tang D.; Zhong S.; Yuan X.; Zheng F.; Zeng Z.; Chen Y.; Liu D.; Hong X.; Dai W.; Yin L.; Dai Y. Absolute Quantification and Characterization of Oxylipins in Lupus Nephritis and Systemic Lupus Erythematosus. Front. Immunol. 2022, 13, 964901. 10.3389/fimmu.2022.964901. [DOI] [PMC free article] [PubMed] [Google Scholar]
Khanapure S. P.; Garvey D. S.; Janero D. R.; Letts L. G. Eicosanoids in Inflammation: Biosynthesis, Pharmacology, and Therapeutic Frontiers. Curr. Top. Med. Chem. 2007, 7, 311–340. 10.2174/156802607779941314. [DOI] [PubMed] [Google Scholar]
Wang B.; Wu L.; Chen J.; Dong L.; Chen C.; Wen Z.; Hu J.; Fleming I.; Wang D. W. Metabolism Pathways of Arachidonic Acids: Mechanisms and Potential Therapeutic Targets. Signal Transduct. Target. Ther. 2021, 6 (1), 94. 10.1038/s41392-020-00443-w. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bazan N. G. Docosanoids and Elovanoids from Omega-3 Fatty Acids Are pro-Homeostatic Modulators of Inflammatory Responses, Cell Damage and Neuroprotection. Mol. Aspects Med. 2018, 64, 18–33. 10.1016/j.mam.2018.09.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
Serhan C. N. Novel Eicosanoid and Docosanoid Mediators: Resolvins, Docosatrienes, and Neuroprotectins. Curr. Opin. Clin. Nutr. Metab. Care 2005, 8 (2), 115–121. 10.1097/00075197-200503000-00003. [DOI] [PubMed] [Google Scholar]
Hajeyah A. A.; Griffiths W. J.; Wang Y.; Finch A. J.; O’Donnell V. B. The Biosynthesis of Enzymatically Oxidized Lipids. Front. Endocrinol. 2020, 11, 591819. 10.3389/fendo.2020.591819. [DOI] [PMC free article] [PubMed] [Google Scholar]
Gabbs M.; Leng S.; Devassy J. G.; Monirujjaman M.; Aukema H. M. Advances in Our Understanding of Oxylipins Derived from Dietary PUFAs. Adv. Nutr. 2015, 6 (5), 513–540. 10.3945/an.114.007732. [DOI] [PMC free article] [PubMed] [Google Scholar]
Christie W. W.; Harwood J. L. Oxidation of Polyunsaturated Fatty Acids to Produce Lipid Mediators. Essays Biochem. 2020, 64 (3), 401–421. 10.1042/EBC20190082. [DOI] [PMC free article] [PubMed] [Google Scholar]
Dennis E. A.; Norris P. C. Eicosanoid Storm in Infection and Inflammation. Nat. Rev. Immunol. 2015, 15 (8), 511–523. 10.1038/nri3859. [DOI] [PMC free article] [PubMed] [Google Scholar]
Serhan C. N. Pro-Resolving Lipid Mediators Are Leads for Resolution Physiology. Nature 2014, 510 (7503), 92–101. 10.1038/nature13479. [DOI] [PMC free article] [PubMed] [Google Scholar]
Chiang N.; Serhan C. N. Specialized Pro-Resolving Mediator Network: An Update on Production and Actions. Essays Biochem. 2020, 64 (3), 443–462. 10.1042/EBC20200018. [DOI] [PMC free article] [PubMed] [Google Scholar]
Calder P. C. Eicosapentaenoic and Docosahexaenoic Acid Derived Specialised Pro-Resolving Mediators: Concentrations in Humans and the Effects of Age, Sex, Disease and Increased Omega-3 Fatty Acid Intake. Biochimie 2020, 178, 105–123. 10.1016/j.biochi.2020.08.015. [DOI] [PubMed] [Google Scholar]
Vidar Hansen T.; Serhan C. N. Protectins: Their Biosynthesis, Metabolism and Structure-Functions. Biochem. Pharmacol. 2022, 206, 115330. 10.1016/j.bcp.2022.115330. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schebb N. H.; Kühn H.; Kahnt A. S.; Rund K. M.; O’Donnell V. B.; Flamand N.; Peters-Golden M.; Jakobsson P.-J.; Weylandt K. H.; Rohwer N.; Murphy R. C.; Geisslinger G.; FitzGerald G. A.; Hanson J.; Dahlgren C.; Alnouri M. W.; Offermanns S.; Steinhilber D. Formation, Signaling and Occurrence of Specialized Pro-Resolving Lipid Mediators—What Is the Evidence so Far?. Front. Pharmacol. 2022, 13, 838782. 10.3389/fphar.2022.838782. [DOI] [PMC free article] [PubMed] [Google Scholar]
Andreou A.; Brodhun F.; Feussner I. Biosynthesis of Oxylipins in Non-Mammals. Prog. Lipid Res. 2009, 48 (3–4), 148–170. 10.1016/j.plipres.2009.02.002. [DOI] [PubMed] [Google Scholar]
Parchem K.; Letsiou S.; Petan T.; Oskolkova O.; Medina I.; Kuda O.; O’Donnell V. B.; Nicolaou A.; Fedorova M.; Bochkov V.; Gladine C. Oxylipin Profiling for Clinical Research: Current Status and Future Perspectives. Prog. Lipid Res. 2024, 95, 101276. 10.1016/j.plipres.2024.101276. [DOI] [PubMed] [Google Scholar]
Schaller F. Enzymes of the Biosynthesis of Octadecanoid-Derived Signalling Molecules. J. Exp. Bot. 2001, 52 (354), 11–23. 10.1093/jexbot/52.354.11. [DOI] [PubMed] [Google Scholar]
Agrawal G. K.; Tamogami S.; Han O.; Iwahashi H.; Rakwal R. Rice Octadecanoid Pathway. Biochem. Biophys. Res. Commun. 2004, 317 (1), 1–15. 10.1016/j.bbrc.2004.03.020. [DOI] [PubMed] [Google Scholar]
Carlson S. E.; Carver J. D.; House S. G. High Fat Diets Varying in Ratios of Polyunsaturated to Saturated Fatty Acid and Linoleic to Linolenic Acid: A Comparison of Rat Neural and Red Cell Membrane Phospholipids. J. Nutr. 1986, 116 (5), 718–725. 10.1093/jn/116.5.718. [DOI] [PubMed] [Google Scholar]
Whelan J.; Fritsche K. Linoleic Acid. Adv. Nutr. 2013, 4 (3), 311–312. 10.3945/an.113.003772. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hutchins P. M.; Murphy R. C. Cholesteryl Ester Acyl Oxidation and Remodeling in Murine Macrophages: Formation of Oxidized Phosphatidylcholine. J. Lipid Res. 2012, 53 (8), 1588–1597. 10.1194/jlr.M026799. [DOI] [PMC free article] [PubMed] [Google Scholar]
Liu G.-Y.; Moon S. H.; Jenkins C. M.; Sims H. F.; Guan S.; Gross R. W. Synthesis of Oxidized Phospholipids by Sn-1 Acyltransferase Using 2–15-HETE Lysophospholipids. J. Biol. Chem. 2019, 294 (26), 10146–10159. 10.1074/jbc.RA119.008766. [DOI] [PMC free article] [PubMed] [Google Scholar]
Burke J. E.; Dennis E. A. Phospholipase A2 Biochemistry. Cardiovasc. Drugs Ther. 2009, 23 (1), 49–59. 10.1007/s10557-008-6132-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wang L.; Gill R.; Pedersen T. L.; Higgins L. J.; Newman J. W.; Rutledge J. C. Triglyceride-Rich Lipoprotein Lipolysis Releases Neutral and Oxidized FFAs That Induce Endothelial Cell Inflammation. J. Lipid Res. 2009, 50 (2), 204–213. 10.1194/jlr.M700505-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Dennis E. A.; Cao J.; Hsu Y.-H.; Magrioti V.; Kokotos G. Phospholipase A2 Enzymes: Physical Structure, Biological Function, Disease Implication, Chemical Inhibition, and Therapeutic Intervention. Chem. Rev. 2011, 111 (10), 6130–6185. 10.1021/cr200085w. [DOI] [PMC free article] [PubMed] [Google Scholar]
Blasbalg T. L.; Hibbeln J. R.; Ramsden C. E.; Majchrzak S. F.; Rawlings R. R. Changes in Consumption of Omega-3 and Omega-6 Fatty Acids in the United States during the 20th Century. Am. J. Clin. Nutr. 2011, 93 (5), 950–962. 10.3945/ajcn.110.006643. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wasternack C. Jasmonates: An Update on Biosynthesis, Signal Transduction and Action in Plant Stress Response, Growth and Development. Ann. Bot. 2007, 100 (4), 681–697. 10.1093/aob/mcm079. [DOI] [PMC free article] [PubMed] [Google Scholar]
Feussner I.; Wasternack C. The Lipoxygenase Pathway. Annu. Rev. Plant Biol. 2002, 53 (1), 275–297. 10.1146/annurev.arplant.53.100301.135248. [DOI] [PubMed] [Google Scholar]
Mosblech A.; Feussner I.; Heilmann I. Oxylipins: Structurally Diverse Metabolites from Fatty Acid Oxidation. Plant Physiol. Biochem. 2009, 47 (6), 511–517. 10.1016/j.plaphy.2008.12.011. [DOI] [PubMed] [Google Scholar]
Sugiyama S.; Hayakawa M.; Nagai S.; Ajioka M.; Ozawa T. Leukotoxin, 9, 10-Epoxy-12-Octadecenoate, Causes Cardiac Failure in Dogs. Life Sci. 1987, 40 (3), 225–231. 10.1016/0024-3205(87)90336-5. [DOI] [PubMed] [Google Scholar]
Greene J. F.; Hammock B. D. Toxicity of Linoleic Acid Metabolites. Adv. Exp. Med. Biol. 1999, 469, 471–477. 10.1007/978-1-4615-4793-8_69. [DOI] [PubMed] [Google Scholar]
Newman J. W.; Morisseau C.; Hammock B. D. Epoxide Hydrolases: Their Roles and Interactions with Lipid Metabolism. Prog. Lipid Res. 2005, 44 (1), 1–51. 10.1016/j.plipres.2004.10.001. [DOI] [PubMed] [Google Scholar]
Hildreth K.; Kodani S. D.; Hammock B. D.; Zhao L. Cytochrome P450-Derived Linoleic Acid Metabolites EpOMEs and DiHOMEs: A Review of Recent Studies. J. Nutr. Biochem. 2020, 86, 108484. 10.1016/j.jnutbio.2020.108484. [DOI] [PMC free article] [PubMed] [Google Scholar]
Chiba T.; Thomas C. P.; Calcutt M. W.; Boeglin W. E.; O’Donnell V. B.; Brash A. R. The Precise Structures and Stereochemistry of Trihydroxy-Linoleates Esterified in Human and Porcine Epidermis and Their Significance in Skin Barrier Function. J. Biol. Chem. 2016, 291 (28), 14540–14554. 10.1074/jbc.M115.711267. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ramsden C. E.; Ringel A.; Majchrzak-Hong S. F.; Yang J.; Blanchard H.; Zamora D.; Loewke J. D.; Rapoport S. I.; Hibbeln J. R.; Davis J. M.; Hammock B. D.; Taha A. Y. Dietary Linoleic Acid-Induced Alterations in pro- and Anti-Nociceptive Lipid Autacoids: Implications for Idiopathic Pain Syndromes?. Mol. Pain 2016, 12, 174480691663638. 10.1177/1744806916636386. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ramsden C. E.; Domenichiello A. F.; Yuan Z.-X.; Sapio M. R.; Keyes G. S.; Mishra S. K.; Gross J. R.; Majchrzak-Hong S.; Zamora D.; Horowitz M. S.; Davis J. M.; Sorokin A. V.; Dey A.; LaPaglia D. M.; Wheeler J. J.; Vasko M. R.; Mehta N. N.; Mannes A. J.; Iadarola M. J. A Systems Approach for Discovering Linoleic Acid Derivatives That Potentially Mediate Pain and Itch. Sci. Signal. 2017, 10 (493), eaal5241 10.1126/scisignal.aal5241. [DOI] [PMC free article] [PubMed] [Google Scholar]
Domenichiello A. F.; Sapio M. R.; Loydpierson A. J.; Maric D.; Goto T.; Horowitz M. S.; Keyes G. S.; Yuan Z.-X.; Majchrzak-Hong S. F.; Mannes A. J.; Iadarola M. J.; Ramsden C. E. Molecular Pathways Linking Oxylipins to Nociception in Rats. J. Pain 2021, 22, 275–299. 10.1016/j.jpain.2020.09.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
Stanford K. I.; Lynes M. D.; Takahashi H.; Baer L. A.; Arts P. J.; May F. J.; Lehnig A. C.; Middelbeek R. J. W.; Richard J. J.; So K.; Chen E. Y.; Gao F.; Narain N. R.; Distefano G.; Shettigar V. K.; Hirshman M. F.; Ziolo M. T.; Kiebish M. A.; Tseng Y.-H.; Coen P. M.; Goodyear L. J. 12,13-DiHOME: An Exercise-Induced Lipokine That Increases Skeletal Muscle Fatty Acid Uptake. Cell Metab. 2018, 27 (5), 1111–1120. 10.1016/j.cmet.2018.03.020. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lynes M. D.; Leiria L. O.; Lundh M.; Bartelt A.; Shamsi F.; Huang T. L.; Takahashi H.; Hirshman M. F.; Schlein C.; Lee A.; Baer L. A.; May F. J.; Gao F.; Narain N. R.; Chen E. Y.; Kiebish M. A.; Cypess A. M.; Blüher M.; Goodyear L. J.; Hotamisligil G. S.; Stanford K. I.; Tseng Y.-H. The Cold-Induced Lipokine 12,13-DiHOME Promotes Fatty Acid Transport into Brown Adipose Tissue. Nat. Med. 2017, 23 (5), 631–637. 10.1038/nm.4297. [DOI] [PMC free article] [PubMed] [Google Scholar]
Cabral M.; Martín-Venegas R.; Moreno J. J. Differential Cell Growth/Apoptosis Behavior of 13-Hydroxyoctadecadienoic Acid Enantiomers in a Colorectal Cancer Cell Line. Am. J. Physiol.-Gastrointest. Liver Physiol. 2014, 307 (6), G664–G671. 10.1152/ajpgi.00064.2014. [DOI] [PubMed] [Google Scholar]
Vangaveti V. N.; Jansen H.; Kennedy R. L.; Malabu U. H. Hydroxyoctadecadienoic Acids: Oxidised Derivatives of Linoleic Acid and Their Role in Inflammation Associated with Metabolic Syndrome and Cancer. Eur. J. Pharmacol. 2016, 785, 70–76. 10.1016/j.ejphar.2015.03.096. [DOI] [PubMed] [Google Scholar]
Kumar N.; Gupta G.; Anilkumar K.; Fatima N.; Karnati R.; Reddy G. V.; Giri P. V.; Reddanna P. 15-Lipoxygenase Metabolites of α-Linolenic Acid, [13-(S)-HPOTrE and 13-(S)-HOTrE], Mediate Anti-Inflammatory Effects by Inactivating NLRP3 Inflammasome. Sci. Rep. 2016, 6 (1), 31649. 10.1038/srep31649. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bergmann C. B.; McReynolds C. B.; Wan D.; Singh N.; Goetzman H.; Caldwell C. C.; Supp D. M.; Hammock B. D. SEH-Derived Metabolites of Linoleic Acid Drive Pathologic Inflammation While Impairing Key Innate Immune Cell Function in Burn Injury. Proc. Natl. Acad. Sci. U. S. A. 2022, 119 (13), e2120691119 10.1073/pnas.2120691119. [DOI] [PMC free article] [PubMed] [Google Scholar]
Gilroy D. W.; Edin M. L.; De Maeyer R. P. H.; Bystrom J.; Newson J.; Lih F. B.; Stables M.; Zeldin D. C.; Bishop-Bailey D. CYP450-Derived Oxylipins Mediate Inflammatory Resolution. Proc. Natl. Acad. Sci. U. S. A. 2016, 113 (23), E3240. 10.1073/pnas.1521453113. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fujimura K. E.; Sitarik A. R.; Havstad S.; Lin D. L.; Levan S.; Fadrosh D.; Panzer A. R.; LaMere B.; Rackaityte E.; Lukacs N. W.; Wegienka G.; Boushey H. A.; Ownby D. R.; Zoratti E. M.; Levin A. M.; Johnson C. C.; Lynch S. V. Neonatal Gut Microbiota Associates with Childhood Multisensitized Atopy and T Cell Differentiation. Nat. Med. 2016, 22 (10), 1187–1191. 10.1038/nm.4176. [DOI] [PMC free article] [PubMed] [Google Scholar]
Levan S. R.; Stamnes K. A.; Lin D. L.; Panzer A. R.; Fukui E.; McCauley K.; Fujimura K. E.; McKean M.; Ownby D. R.; Zoratti E. M.; Boushey H. A.; Cabana M. D.; Johnson C. C.; Lynch S. V. Elevated Faecal 12,13-DiHOME Concentration in Neonates at High Risk for Asthma Is Produced by Gut Bacteria and Impedes Immune Tolerance. Nat. Microbiol. 2019, 4 (11), 1851–1861. 10.1038/s41564-019-0498-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bell M. V.; Dick J. R.; Pond D. W. Octadecapentaenoic Acid in a Raphidophyte Alga, Heterosigma akashiwo. Phytochemistry 1997, 45 (2), 303–306. 10.1016/S0031-9422(96)00867-9. [DOI] [Google Scholar]
Trostchansky A.; Bonilla L.; González-Perilli L.; Rubbo H. Nitro-Fatty Acids: Formation, Redox Signaling, and Therapeutic Potential. Antioxid. Redox Signal. 2013, 19 (11), 1257–1265. 10.1089/ars.2012.5023. [DOI] [PubMed] [Google Scholar]
Mata-Pérez C.; Sánchez-Calvo B.; Padilla M. N.; Begara-Morales J. C.; Valderrama R.; Corpas F. J.; Barroso J. B. Nitro-Fatty Acids in Plant Signaling: New Key Mediators of Nitric Oxide Metabolism. Redox Biol. 2017, 11, 554–561. 10.1016/j.redox.2017.01.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Delmastro-Greenwood M.; Freeman B. A.; Wendell S. G. Redox-Dependent Anti-Inflammatory Signaling Actions of Unsaturated Fatty Acids. Annu. Rev. Physiol. 2014, 76 (1), 79–105. 10.1146/annurev-physiol-021113-170341. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schopfer F. J.; Cipollina C.; Freeman B. A. Formation and Signaling Actions of Electrophilic Lipids. Chem. Rev. 2011, 111 (10), 5997–6021. 10.1021/cr200131e. [DOI] [PMC free article] [PubMed] [Google Scholar]
Quaranta A.; Revol-Cavalier J.; Wheelock C. E. The Octadecanoids: An Emerging Class of Lipid Mediators. Biochem. Soc. Trans. 2022, 50 (6), 1569–1582. 10.1042/BST20210644. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yamada Y.; Uemura H.; Nakaya H.; Sakata K.; Takatori T.; Nagao M.; Iwase H.; Iwadate K. Production of Hydroxy Fatty Acid (10-Hydroxy-12(Z)-Octadecenoic Acid) by Lactobacillus plantarum from Linoleic Acid and Its Cardiac Effects to Guinea Pig Papillary Muscles. Biochem. Biophys. Res. Commun. 1996, 226 (2), 391–395. 10.1006/bbrc.1996.1366. [DOI] [PubMed] [Google Scholar]
Miyamoto J.; Igarashi M.; Watanabe K.; Karaki S.; Mukouyama H.; Kishino S.; Li X.; Ichimura A.; Irie J.; Sugimoto Y.; Mizutani T.; Sugawara T.; Miki T.; Ogawa J.; Drucker D. J.; Arita M.; Itoh H.; Kimura I. Gut Microbiota Confers Host Resistance to Obesity by Metabolizing Dietary Polyunsaturated Fatty Acids. Nat. Commun. 2019, 10 (1), 4007. 10.1038/s41467-019-11978-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
Tang L.; Shang J.; Song C.; Yang R.; Shang X.; Mao W.; Bao D.; Tan Q. Untargeted Metabolite Profiling of Antimicrobial Compounds in the Brown Film of Lentinula edodes Mycelium via LC-MS/MS Analysis. ACS Omega 2020, 5 (13), 7567–7575. 10.1021/acsomega.0c00398. [DOI] [PMC free article] [PubMed] [Google Scholar]
Takahashi H.; Kamakari K.; Goto T.; Hara H.; Mohri S.; Suzuki H.; Shibata D.; Nakata R.; Inoue H.; Takahashi N.; Kawada T. 9-Oxo-10(E),12(Z),15(Z)-Octadecatrienoic Acid Activates Peroxisome Proliferator-Activated Receptor α in Hepatocytes. Lipids 2015, 50 (11), 1083–1091. 10.1007/s11745-015-4071-3. [DOI] [PubMed] [Google Scholar]
Petta T.; Secatto A.; Faccioli L. H.; Moraes L. A. B. Inhibition of Inflammatory Response in LPS Induced Macrophages by 9-KOTE and 13-KOTE Produced by Biotransformation. Enzyme Microb. Technol. 2014, 58–59, 36–43. 10.1016/j.enzmictec.2014.02.011. [DOI] [PubMed] [Google Scholar]
Fahy E.; Subramaniam S.; Murphy R. C.; Nishijima M.; Raetz C. R. H.; Shimizu T.; Spener F.; van Meer G.; Wakelam M. J. O.; Dennis E. A. Update of the LIPID MAPS Comprehensive Classification System for Lipids. J. Lipid Res. 2009, 50, S9–S14. 10.1194/jlr.R800095-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sud M.; Fahy E.; Cotter D.; Brown A.; Dennis E. A.; Glass C. K.; Merrill A. H. Jr; Murphy R. C.; Raetz C. R. H.; Russell D. W.; Subramaniam S. LMSD: LIPID MAPS Structure Database. Nucleic Acids Res. 2007, 35, D527–D532. 10.1093/nar/gkl838. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wishart D. S.; Guo A.; Oler E.; Wang F.; Anjum A.; Peters H.; Dizon R.; Sayeeda Z.; Tian S.; Lee B. L.; Berjanskii M.; Mah R.; Yamamoto M.; Jovel J.; Torres-Calzada C.; Hiebert-Giesbrecht M.; Lui V. W.; Varshavi D.; Varshavi D.; Allen D.; Arndt D.; Khetarpal N.; Sivakumaran A.; Harford K.; Sanford S.; Yee K.; Cao X.; Budinski Z.; Liigand J.; Zhang L.; Zheng J.; Mandal R.; Karu N.; Dambrova M.; Schiöth H. B.; Greiner R.; Gautam V. HMDB 5.0: The Human Metabolome Database for 2022. Nucleic Acids Res. 2022, 50 (D1), D622–D631. 10.1093/nar/gkab1062. [DOI] [PMC free article] [PubMed] [Google Scholar]
Smith D. L.; Willis A. L. A Suggested Shorthand Nomenclature for the Eicosanoids. Lipids 1987, 22 (12), 983–986. 10.1007/BF02536436. [DOI] [PubMed] [Google Scholar]
Taber D. F.; Morrow J. D.; Roberts L. J II. A Nomenclature System for the Isoprostanes. Prostaglandins 1997, 53, 63–67. 10.1016/S0090-6980(97)00005-1. [DOI] [PubMed] [Google Scholar]
Taber D. F.; Roberts L. J. Nomenclature Systems for the Neuroprostanes and for the Neurofurans. Prostaglandins Other Lipid Mediat. 2005, 78 (1–4), 14–18. 10.1016/j.prostaglandins.2005.07.002. [DOI] [PubMed] [Google Scholar]
Taber D. F.; Fessel J. P.; Roberts L. J. A Nomenclature System for Isofurans. Prostaglandins Other Lipid Mediat. 2004, 73 (1), 47–50. 10.1016/j.prostaglandins.2003.11.004. [DOI] [PubMed] [Google Scholar]
Vick B. A.; Zimmerman D. C. Biosynthesis of Jasmonic Acid by Several Plant Species. Plant Physiol. 1984, 75 (2), 458–461. 10.1104/pp.75.2.458. [DOI] [PMC free article] [PubMed] [Google Scholar]
Grechkin A. N.; Hamberg M. The “Heterolytic Hydroperoxide Lyase” Is an Isomerase Producing a Short-Lived Fatty Acid Hemiacetal. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2004, 1636 (1), 47–58. 10.1016/j.bbalip.2003.12.003. [DOI] [PubMed] [Google Scholar]
Hamberg M. A Pathway for Biosynthesis of Divinyl Ether Fatty Acids in Green Leaves. Lipids 1998, 33 (11), 1061–1071. 10.1007/s11745-998-0306-7. [DOI] [PubMed] [Google Scholar]
Toporkova Y. Y.; Smirnova E. O.; Lantsova N. V.; Mukhtarova L. S.; Grechkin A. N. Detection of the First Epoxyalcohol Synthase/Allene Oxide Synthase (CYP74 Clan) in the Lancelet (Branchiostoma belcheri, Chordata). Int. J. Mol. Sci. 2021, 22 (9), 4737. 10.3390/ijms22094737. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lee M.; Lenman M.; Banaś A.; Bafor M.; Singh S.; Schweizer M.; Nilsson R.; Liljenberg C.; Dahlqvist A.; Gummeson P.-O.; Sjödahl S.; Green A.; Stymne S. Identification of Non-Heme Diiron Proteins That Catalyze Triple Bond and Epoxy Group Formation. Science 1998, 280 (5365), 915–918. 10.1126/science.280.5365.915. [DOI] [PubMed] [Google Scholar]
Laneuville O.; Breuer D. K.; Xu N.; Huang Z. H.; Gage D. A.; Watson J. T.; Lagarde M.; DeWitt D. L.; Smith W. L. Fatty Acid Substrate Specificities of Human Prostaglandin-Endoperoxide H Synthase-1 and – 2: FORMATION OF 12-HYDROXY-(9Z,13E/Z,15Z)-OCTADECATRIENOIC ACIDS FROM α-LINOLENIC ACID. J. Biol. Chem. 1995, 270 (33), 19330–19336. 10.1074/jbc.270.33.19330. [DOI] [PubMed] [Google Scholar]
Brash A. R.; Boeglin W. E.; Chang M. S. Discovery of a Second 15S-Lipoxygenase in Humans. Proc. Natl. Acad. Sci. U. S. A. 1997, 94 (12), 6148–6152. 10.1073/pnas.94.12.6148. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bylund J.; Ericsson J.; Oliw E. H. Analysis of Cytochrome P450 Metabolites of Arachidonic and Linoleic Acids by Liquid Chromatography-Mass Spectrometry with Ion Trap MS2. Anal. Biochem. 1998, 265 (1), 55–68. 10.1006/abio.1998.2897. [DOI] [PubMed] [Google Scholar]
Oliw E. H.; Bylund J.; Herman C. Bisallylic Hydroxylation and Epoxidation of Polyunsaturated Fatty Acids by Cytochrome P450. Lipids 1996, 31 (10), 1003–1021. 10.1007/BF02522457. [DOI] [PubMed] [Google Scholar]
Schuchardt J. P.; Schmidt S.; Kressel G.; Dong H.; Willenberg I.; Hammock B. D.; Hahn A.; Schebb N. H. Comparison of Free Serum Oxylipin Concentrations in Hyper- vs. Normolipidemic Men. Prostaglandins Leukot. Essent. Fatty Acids 2013, 89 (1), 19–29. 10.1016/j.plefa.2013.04.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yu Z.; Schneider C.; Boeglin W. E.; Marnett L. J.; Brash A. R. The Lipoxygenase Gene ALOXE3 Implicated in Skin Differentiation Encodes a Hydroperoxide Isomerase. Proc. Natl. Acad. Sci. U. S. A. 2003, 100 (16), 9162–9167. 10.1073/pnas.1633612100. [DOI] [PMC free article] [PubMed] [Google Scholar]
Liu M.; Chen P.; Véricel E.; Lelli M.; Béguin L.; Lagarde M.; Guichardant M. Characterization and Biological Effects of Di-Hydroxylated Compounds Deriving from the Lipoxygenation of ALA. J. Lipid Res. 2013, 54 (8), 2083–2094. 10.1194/jlr.M035139. [DOI] [PMC free article] [PubMed] [Google Scholar]
Jahn U.; Galano J.-M.; Durand T. Beyond Prostaglandins—Chemistry and Biology of Cyclic Oxygenated Metabolites Formed by Free-Radical Pathways from Polyunsaturated Fatty Acids. Angew. Chem., Int. Ed. 2008, 47 (32), 5894–5955. 10.1002/anie.200705122. [DOI] [PubMed] [Google Scholar]
Leung K. S.; Oger C.; Guy A.; Bultel-Poncé V.; Vigor C.; Durand T.; Gil-Izquierdo A.; Medina S.; Galano J.-M.; Lee J. C.-Y.. Alpha-Linolenic Acid, Phytoprostanes and Phytofurans in Plant, Algae and Food. In Lipids in Plants and Algae: From Fundamental Science to Industrial Applications; Advances in Botanical Research; Elsevier, 2022; Vol. 101, p 437. 10.1016/bs.abr.2021.09.005. [DOI] [Google Scholar]
Mata-Pérez C.; Sánchez-Calvo B.; Padilla M. N.; Begara-Morales J. C.; Luque F.; Melguizo M.; Jiménez-Ruiz J.; Fierro-Risco J.; Peñas-Sanjuán A.; Valderrama R.; Corpas F. J.; Barroso J. B. Nitro-Fatty Acids in Plant Signaling: Nitro-Linolenic Acid Induces the Molecular Chaperone Network in Arabidopsis. Plant Physiol. 2016, 170 (2), 686–701. 10.1104/pp.15.01671. [DOI] [PMC free article] [PubMed] [Google Scholar]
Koutoulogenis G. S.; Kokotos G. Nitro Fatty Acids (NO₂-FAs): An Emerging Class of Bioactive Fatty Acids. Molecules 2021, 26 (24), 7536. 10.3390/molecules26247536. [DOI] [PMC free article] [PubMed] [Google Scholar]
Diczfalusy U. Beta-Oxidation of Eicosanoids. Prog. Lipid Res. 1994, 33 (4), 403–428. 10.1016/0163-7827(94)90025-6. [DOI] [PubMed] [Google Scholar]
Misheva M.; Kotzamanis K.; Davies L. C.; Tyrrell V. J.; Rodrigues P. R. S.; Benavides G. A.; Hinz C.; Murphy R. C.; Kennedy P.; Taylor P. R.; Rosas M.; Jones S. A.; McLaren J. E.; Deshpande S.; Andrews R.; Schebb N. H.; Czubala M. A.; Gurney M.; Aldrovandi M.; Meckelmann S. W.; Ghazal P.; Darley-Usmar V.; White D. A.; O’Donnell V. B. Oxylipin Metabolism Is Controlled by Mitochondrial β-Oxidation during Bacterial Inflammation. Nat. Commun. 2022, 13 (1), 139. 10.1038/s41467-021-27766-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
Whelan J. The Health Implications of Changing Linoleic Acid Intakes. Prostaglandins Leukot. Essent. Fatty Acids 2008, 79 (3), 165–167. 10.1016/j.plefa.2008.09.013. [DOI] [PubMed] [Google Scholar]
Montani J.-P. Ancel Keys: The Legacy of a Giant in Physiology, Nutrition, and Public Health. Obes. Rev. 2021, 22, e13196 10.1111/obr.13196. [DOI] [PubMed] [Google Scholar]
Mercola J.; D’Adamo C. R. Linoleic Acid: A Narrative Review of the Effects of Increased Intake in the Standard American Diet and Associations with Chronic Disease. Nutrients 2023, 15 (14), 3129. 10.3390/nu15143129. [DOI] [PMC free article] [PubMed] [Google Scholar]
Marangoni F.; Agostoni C.; Borghi C.; Catapano A. L.; Cena H.; Ghiselli A.; La Vecchia C.; Lercker G.; Manzato E.; Pirillo A.; Riccardi G.; Risé P.; Visioli F.; Poli A. Dietary Linoleic Acid and Human Health: Focus on Cardiovascular and Cardiometabolic Effects. Atherosclerosis 2020, 292, 90–98. 10.1016/j.atherosclerosis.2019.11.018. [DOI] [PubMed] [Google Scholar]
Li J.; Guasch-Ferré M.; Li Y.; Hu F. B. Dietary Intake and Biomarkers of Linoleic Acid and Mortality: Systematic Review and Meta-Analysis of Prospective Cohort Studies. Am. J. Clin. Nutr. 2020, 112 (1), 150–167. 10.1093/ajcn/nqz349. [DOI] [PMC free article] [PubMed] [Google Scholar]
Taha A. Y. Linoleic Acid-Good or Bad for the Brain?. NPJ. Sci. Food 2020, 4, 1. 10.1038/s41538-019-0061-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lankinen M. A.; Fauland A.; Shimizu B.-I.; Ågren J.; Wheelock C. E.; Laakso M.; Schwab U.; Pihlajamäki J. Inflammatory Response to Dietary Linoleic Acid Depends on FADS1 Genotype. Am. J. Clin. Nutr. 2019, 109 (1), 165–175. 10.1093/ajcn/nqy287. [DOI] [PubMed] [Google Scholar]
Ramsden C. E.; Hennebelle M.; Schuster S.; Keyes G. S.; Johnson C. D.; Kirpich I. A.; Dahlen J. E.; Horowitz M. S.; Zamora D.; Feldstein A. E.; McClain C. J.; Muhlhausler B. S.; Makrides M.; Gibson R. A.; Taha A. Y. Effects of Diets Enriched in Linoleic Acid and Its Peroxidation Products on Brain Fatty Acids, Oxylipins, and Aldehydes in Mice. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2018, 1863 (10), 1206–1213. 10.1016/j.bbalip.2018.07.007. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ramsden C. E.; Ringel A.; Feldstein A. E.; Taha A. Y.; MacIntosh B. A.; Hibbeln J. R.; Majchrzak-Hong S. F.; Faurot K. R.; Rapoport S. I.; Cheon Y.; Chung Y.-M.; Berk M.; Mann J. D. Lowering Dietary Linoleic Acid Reduces Bioactive Oxidized Linoleic Acid Metabolites in Humans. Prostaglandins Leukot. Essent. Fatty Acids 2012, 87, 135–141. 10.1016/j.plefa.2012.08.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
Liang N.; Hennebelle M.; Gaul S.; Johnson C. D.; Zhang Z.; Kirpich I. A.; McClain C. J.; Feldstein A. E.; Ramsden C. E.; Taha A. Y. Feeding Mice a Diet High in Oxidized Linoleic Acid Metabolites Does Not Alter Liver Oxylipin Concentrations. Prostaglandins Leukot. Essent. Fatty Acids 2021, 172, 102316. 10.1016/j.plefa.2021.102316. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schuster S.; Johnson C. D.; Hennebelle M.; Holtmann T.; Taha A. Y.; Kirpich I. A.; Eguchi A.; Ramsden C. E.; Papouchado B. G.; McClain C. J.; Feldstein A. E. Oxidized Linoleic Acid Metabolites Induce Liver Mitochondrial Dysfunction, Apoptosis, and NLRP3 Activation in Mice. J. Lipid Res. 2018, 59 (9), 1597–1609. 10.1194/jlr.M083741. [DOI] [PMC free article] [PubMed] [Google Scholar]
Caligiuri S. P. B.; Parikh M.; Stamenkovic A.; Pierce G. N.; Aukema H. M. Dietary Modulation of Oxylipins in Cardiovascular Disease and Aging. Am. J. Physiol. Heart Circ. Physiol. 2017, 313 (5), H903–H918. 10.1152/ajpheart.00201.2017. [DOI] [PubMed] [Google Scholar]
Ostermann A. I.; Schebb N. H. Effects of Omega-3 Fatty Acid Supplementation on the Pattern of Oxylipins: A Short Review about the Modulation of Hydroxy-, Dihydroxy-, and Epoxy-Fatty Acids. Food Funct. 2017, 8 (7), 2355–2367. 10.1039/C7FO00403F. [DOI] [PubMed] [Google Scholar]
Innes J. K.; Calder P. C. Omega-6 Fatty Acids and Inflammation. Prostaglandins Leukot. Essent. Fatty Acids 2018, 132, 41–48. 10.1016/j.plefa.2018.03.004. [DOI] [PubMed] [Google Scholar]
Hamilton J. S.; Klett E. L. Linoleic Acid and the Regulation of Glucose Homeostasis: A Review of the Evidence. Prostaglandins Leukot. Essent. Fatty Acids 2021, 175, 102366. 10.1016/j.plefa.2021.102366. [DOI] [PMC free article] [PubMed] [Google Scholar]
Belury M. A.; Cole R. M.; Bailey B. E.; Ke J.-Y.; Andridge R. R.; Kiecolt-Glaser J. K. Erythrocyte Linoleic Acid, but Not Oleic Acid, Is Associated with Improvements in Body Composition in Men and Women. Mol. Nutr. Food Res. 2016, 60 (5), 1206–1212. 10.1002/mnfr.201500744. [DOI] [PMC free article] [PubMed] [Google Scholar]
Shahidi F.; Ambigaipalan P. Omega-3 Polyunsaturated Fatty Acids and Their Health Benefits. Annu. Rev. Food Sci. Technol. 2018, 9 (1), 345–381. 10.1146/annurev-food-111317-095850. [DOI] [PubMed] [Google Scholar]
Spector A. A.; Kim H.-Y. Emergence of Omega-3 Fatty Acids in Biomedical Research. Prostaglandins Leukot. Essent. Fatty Acids 2019, 140, 47–50. 10.1016/j.plefa.2018.11.017. [DOI] [PMC free article] [PubMed] [Google Scholar]
Galli C.; Marangoni F. N-3 Fatty Acids in the Mediterranean Diet. Prostaglandins Leukot. Essent. Fatty Acids 2006, 75 (3), 129–133. 10.1016/j.plefa.2006.05.007. [DOI] [PubMed] [Google Scholar]
Naghshi S.; Aune D.; Beyene J.; Mobarak S.; Asadi M.; Sadeghi O. Dietary Intake and Biomarkers of Alpha Linolenic Acid and Risk of All Cause, Cardiovascular, and Cancer Mortality: Systematic Review and Dose-Response Meta-Analysis of Cohort Studies. BMJ. 2021, 375, n2213. 10.1136/bmj.n2213. [DOI] [PMC free article] [PubMed] [Google Scholar]
Seidel U.; Eberhardt K.; Wiebel M.; Luersen K.; Ipharraguerre I. R.; Haegele F. A.; Winterhalter P.; Bosy-Westphal A.; Schebb N. H.; Rimbach G. Stearidonic Acid Improves Eicosapentaenoic Acid Status: Studies in Humans and Cultured Hepatocytes. Front. Nutr. 2024, 11, 1359958. 10.3389/fnut.2024.1359958. [DOI] [PMC free article] [PubMed] [Google Scholar]
Benjamin S.; Spener F. Conjugated Linoleic Acids as Functional Food: An Insight into Their Health Benefits. Nutr. Metab. 2009, 6 (1), 36. 10.1186/1743-7075-6-36. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yuan G.-F.; Chen X.-E.; Li D. Conjugated Linolenic Acids and Their Bioactivities: A Review. Food Funct. 2014, 5 (7), 1360–1368. 10.1039/c4fo00037d. [DOI] [PubMed] [Google Scholar]
Paul D.; Dey T. K.; Chakraborty A.; Dhar P.. Chapter 13 - Promising Functional Lipids for Therapeutic Applications. In Role of Materials Science in Food Bioengineering; Handbook of Food Bioengineering; Grumezescu A. M., Holban A. M., Eds.; Academic Press, 2018; pp 413–449. 10.1016/B978-0-12-811448-3.00013-9. [DOI] [Google Scholar]
Yuan G.; Sinclair A. J.; Xu C.; Li D. Incorporation and Metabolism of Punicic Acid in Healthy Young Humans. Mol. Nutr. Food Res. 2009, 53 (10), 1336–1342. 10.1002/mnfr.200800520. [DOI] [PubMed] [Google Scholar]
Shabbir M. A.; Khan M. R.; Saeed M.; Pasha I.; Khalil A. A.; Siraj N. Punicic Acid: A Striking Health Substance to Combat Metabolic Syndromes in Humans. Lipids Health Dis. 2017, 16 (1), 99. 10.1186/s12944-017-0489-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
De Caterina R. N-3 Fatty Acids in Cardiovascular Disease. N. Engl. J. Med. 2011, 364 (25), 2439–2450. 10.1056/NEJMra1008153. [DOI] [PubMed] [Google Scholar]
Cao M.; Yang F.; McClements D. J.; Guo Y.; Liu R.; Chang M.; Wei W.; Jin J.; Wang X. Impact of Dietary N-6/n-3 Fatty Acid Ratio of Atherosclerosis Risk: A Review. Prog. Lipid Res. 2024, 95, 101289. 10.1016/j.plipres.2024.101289. [DOI] [PubMed] [Google Scholar]
Risérus U.; Willett W. C.; Hu F. B. Dietary Fats and Prevention of Type 2 Diabetes. Prog. Lipid Res. 2009, 48 (1), 44–51. 10.1016/j.plipres.2008.10.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Erkkilä A.; de Mello V. D. F.; Risérus U.; Laaksonen D. E. Dietary Fatty Acids and Cardiovascular Disease: An Epidemiological Approach. Prog. Lipid Res. 2008, 47 (3), 172–187. 10.1016/j.plipres.2008.01.004. [DOI] [PubMed] [Google Scholar]
Wendell S. G.; Baffi C.; Holguin F. Fatty Acids, Inflammation, and Asthma. J. Allergy Clin. Immunol. 2014, 133 (5), 1255–1264. 10.1016/j.jaci.2013.12.1087. [DOI] [PMC free article] [PubMed] [Google Scholar]
Brühl L.; Weisshaar R.; Matthäus B. Epoxy Fatty Acids in Used Frying Fats and Oils, Edible Oils and Chocolate and Their Formation in Oils during Heating. Eur. J. Lipid Sci. Technol. 2016, 118 (3), 425–434. 10.1002/ejlt.201500235. [DOI] [Google Scholar]
Emami S.; Zhang Z.; Taha A. Y. Quantitation of Oxylipins in Fish and Algae Oil Supplements Using Optimized Hydrolysis Procedures and Ultra-High Performance Liquid Chromatography Coupled to Tandem Mass-Spectrometry. J. Agric. Food Chem. 2020, 68 (35), 9329–9344. 10.1021/acs.jafc.0c02461. [DOI] [PubMed] [Google Scholar]
Koch E.; Wiebel M.; Löwen A.; Willenberg I.; Schebb N. H. Characterization of the Oxylipin Pattern and Other Fatty Acid Oxidation Products in Freshly Pressed and Stored Plant Oils. J. Agric. Food Chem. 2022, 70 (40), 12935–12945. 10.1021/acs.jafc.2c04987. [DOI] [PubMed] [Google Scholar]
Richardson C. E.; Hennebelle M.; Otoki Y.; Zamora D.; Yang J.; Hammock B. D.; Taha A. Y. Lipidomic Analysis of Oxidized Fatty Acids in Plant and Algae Oils. J. Agric. Food Chem. 2017, 65 (9), 1941–1951. 10.1021/acs.jafc.6b05559. [DOI] [PMC free article] [PubMed] [Google Scholar]
Devassy J. G.; Yamaguchi T.; Monirujjaman M.; Gabbs M.; Ravandi A.; Zhou J.; Aukema H. M. Distinct Effects of Dietary Flax Compared to Fish Oil, Soy Protein Compared to Casein, and Sex on the Renal Oxylipin Profile in Models of Polycystic Kidney Disease. Prostaglandins Leukot. Essent. Fatty Acids 2017, 123, 1–13. 10.1016/j.plefa.2017.07.002. [DOI] [PubMed] [Google Scholar]
Domínguez-Perles R.; Abellán Á.; León D.; Ferreres F.; Guy A.; Oger C.; Galano J. M.; Durand T.; Gil-Izquierdo Á. Sorting out the Phytoprostane and Phytofuran Profile in Vegetable Oils. Food Res. Int. 2018, 107, 619–628. 10.1016/j.foodres.2018.03.013. [DOI] [PubMed] [Google Scholar]
Mubiru E.; Jacxsens L.; Papastergiadis A.; Lachat C.; Shrestha K.; Mozumder N. H. M. R.; De Meulenaer B. Exposure Assessment of Epoxy Fatty Acids through Consumption of Specific Foods Available in Belgium. Food Addit. Contam. Part A 2017, 34 (6), 1000–1011. 10.1080/19440049.2017.1310399. [DOI] [PubMed] [Google Scholar]
Koch E.; Löwen A.; Schebb N. H. Do Meals Contain a Relevant Amount of Oxylipins? LC-MS-Based Analysis of Oxidized Fatty Acids in Food. Food Chem. 2024, 438, 137941. 10.1016/j.foodchem.2023.137941. [DOI] [PubMed] [Google Scholar]
Abbattista R.; Feinberg N. G.; Snodgrass I. F.; Newman J. W.; Dandekar A. M. Unveiling the “Hidden Quality” of the Walnut Pellicle: A Precious Source of Bioactive Lipids. Front. Plant Sci. 2024, 15, 1395543. 10.3389/fpls.2024.1395543. [DOI] [PMC free article] [PubMed] [Google Scholar]
Nourooz-Zadeh J.; Tajaddini-Sarmadi J.; Wolff S. P. Measurement of Hydroperoxides in Edible Oils Using the Ferrous Oxidation in Xylenol Orange Assay. J. Agric. Food Chem. 1995, 43 (1), 17–21. 10.1021/jf00049a005. [DOI] [Google Scholar]
Choe E.; Min D. B. Mechanisms and Factors for Edible Oil Oxidation. Compr. Rev. Food Sci. Food Saf. 2006, 5 (4), 169–186. 10.1111/j.1541-4337.2006.00009.x. [DOI] [Google Scholar]
Domínguez R.; Pateiro M.; Gagaoua M.; Barba F. J.; Zhang W.; Lorenzo J. M. A Comprehensive Review on Lipid Oxidation in Meat and Meat Products. Antioxidants 2019, 8 (10), 429. 10.3390/antiox8100429. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kanner J.; Lapidot T. The Stomach as a Bioreactor: Dietary Lipid Peroxidation in the Gastric Fluid and the Effects of Plant-Derived Antioxidants. Free Radic. Biol. Med. 2001, 31 (11), 1388–1395. 10.1016/S0891-5849(01)00718-3. [DOI] [PubMed] [Google Scholar]
Márquez-Ruiz G.; Holgado F.; Ruiz-Méndez M. V.; Velasco J. Chemical Changes of Hydroperoxy-, Epoxy-, Keto- and Hydroxy-Model Lipids under Simulated Gastric Conditions. Foods 2021, 10 (9), 2035. 10.3390/foods10092035. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zhang Z.; Emami S.; Hennebelle M.; Morgan R. K.; Lerno L. A.; Slupsky C. M.; Lein P. J.; Taha A. Y. Linoleic Acid-Derived 13-Hydroxyoctadecadienoic Acid Is Absorbed and Incorporated into Rat Tissues. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2021, 1866 (3), 158870. 10.1016/j.bbalip.2020.158870. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kanazawa K.; Ashida H. Catabolic Fate of Dietary Trilinoleoylglycerol Hydroperoxides in Rat Gastrointestines. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1998, 1393 (2–3), 336–348. 10.1016/S0005-2760(98)00088-5. [DOI] [PubMed] [Google Scholar]
Ringseis R.; Eder K. Regulation of Genes Involved in Lipid Metabolism by Dietary Oxidized Fat. Mol. Nutr. Food Res. 2011, 55 (1), 109–121. 10.1002/mnfr.201000424. [DOI] [PubMed] [Google Scholar]
Zaunschirm M.; Pignitter M.; Kopic A.; Keßler C.; Hochkogler C.; Kretschy N.; Somoza M. M.; Somoza V. Exposure of Human Gastric Cells to Oxidized Lipids Stimulates Pathways of Amino Acid Biosynthesis on a Genomic and Metabolomic Level. Molecules 2019, 24 (22), 4111. 10.3390/molecules24224111. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fazzari M.; Trostchansky A.; Schopfer F. J.; Salvatore S. R.; Sánchez-Calvo B.; Vitturi D.; Valderrama R.; Barroso J. B.; Radi R.; Freeman B. A.; Rubbo H. Olives and Olive Oil Are Sources of Electrophilic Fatty Acid Nitroalkenes. PLoS One 2014, 9 (1), e84884 10.1371/journal.pone.0084884. [DOI] [PMC free article] [PubMed] [Google Scholar]
Khoo N. K.; Schopfer F. J. Nitrated Fatty Acids: From Diet to Disease. Curr. Opin. Physiol. 2019, 9, 67–72. 10.1016/j.cophys.2019.04.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
Karwowska M.; Kononiuk A. Nitrates/Nitrites in Food—Risk for Nitrosative Stress and Benefits. Antioxidants 2020, 9 (3), 241. 10.3390/antiox9030241. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bonacci G.; Baker P. R. S.; Salvatore S. R.; Shores D.; Khoo N. K. H.; Koenitzer J. R.; Vitturi D. A.; Woodcock S. R.; Golin-Bisello F.; Cole M. P.; Watkins S.; St Croix C.; Batthyany C. I.; Freeman B. A.; Schopfer F. J. Conjugated Linoleic Acid Is a Preferential Substrate for Fatty Acid Nitration. J. Biol. Chem. 2012, 287, 44071–44082. 10.1074/jbc.M112.401356. [DOI] [PMC free article] [PubMed] [Google Scholar]
Charles R. L.; Rudyk O.; Prysyazhna O.; Kamynina A.; Yang J.; Morisseau C.; Hammock B. D.; Freeman B. A.; Eaton P. Protection from Hypertension in Mice by the Mediterranean Diet Is Mediated by Nitro Fatty Acid Inhibition of Soluble Epoxide Hydrolase. Proc. Natl. Acad. Sci. U. S. A. 2014, 111 (22), 8167–8172. 10.1073/pnas.1402965111. [DOI] [PMC free article] [PubMed] [Google Scholar]
Duarte S.; Melo T.; Domingues R.; de Dios Alché J.; Pérez-Sala D. Insight into the Cellular Effects of Nitrated Phospholipids: Evidence for Pleiotropic Mechanisms of Action. Free Radic. Biol. Med. 2019, 144, 192–202. 10.1016/j.freeradbiomed.2019.06.003. [DOI] [PubMed] [Google Scholar]
O’Neill G. P.; Ford-Hutchinson A. W. Expression of MRNA for Cyclooxygenase-1 and Cyclooxygenase-2 in Human Tissues. FEBS Lett. 1993, 330 (2), 157–160. 10.1016/0014-5793(93)80263-T. [DOI] [PubMed] [Google Scholar]
Zidar N.; Odar K.; Glavac D.; Jerse M.; Zupanc T.; Stajer D. Cyclooxygenase in Normal Human Tissues - Is COX-1 Really a Constitutive Isoform, and COX-2 an Inducible Isoform?. J. Cell. Mol. Med. 2009, 13 (9b), 3753–3763. 10.1111/j.1582-4934.2008.00430.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kirkby N. S.; Chan M. V.; Zaiss A. K.; Garcia-Vaz E.; Jiao J.; Berglund L. M.; Verdu E. F.; Ahmetaj-Shala B.; Wallace J. L.; Herschman H. R.; Gomez M. F.; Mitchell J. A. Systematic Study of Constitutive Cyclooxygenase-2 Expression: Role of NF-ΚB and NFAT Transcriptional Pathways. Proc. Natl. Acad. Sci. U. S. A. 2016, 113 (2), 434–439. 10.1073/pnas.1517642113. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ringbom T.; Huss U.; Stenholm Å.; Flock S.; Skattebøl L.; Perera P.; Bohlin L. COX-2 Inhibitory Effects of Naturally Occurring and Modified Fatty Acids. J. Nat. Prod. 2001, 64 (6), 745–749. 10.1021/np000620d. [DOI] [PubMed] [Google Scholar]
Laneuville O.; Breuer D. K.; Xu N.; Huang Z. H.; Gage D. A.; Watson J. T.; Lagarde M.; DeWitt D. L.; Smith W. L. Fatty Acid Substrate Specificities of Human Prostaglandin-Endoperoxide H Synthase-1 and – 2. J. Biol. Chem. 1995, 270 (33), 19330–19336. 10.1074/jbc.270.33.19330. [DOI] [PubMed] [Google Scholar]
Marnett L. J. Cyclooxygenase Mechanisms. Curr. Opin. Chem. Biol. 2000, 4 (5), 545–552. 10.1016/S1367-5931(00)00130-7. [DOI] [PubMed] [Google Scholar]
Hamberg M. Stereochemistry of Oxygenation of Linoleic Acid Catalyzed by Prostaglandin-Endoperoxide H Synthase-2. Arch. Biochem. Biophys. 1998, 349 (2), 376–380. 10.1006/abbi.1997.0443. [DOI] [PubMed] [Google Scholar]
Altmann R.; Hausmann M.; Spöttl T.; Gruber M.; Bull A. W.; Menzel K.; Vogl D.; Herfarth H.; Schölmerich J.; Falk W.; Rogler G. 13-Oxo-ODE Is an Endogenous Ligand for PPARγ in Human Colonic Epithelial Cells. Biochem. Pharmacol. 2007, 74 (4), 612–622. 10.1016/j.bcp.2007.05.027. [DOI] [PubMed] [Google Scholar]
Funk C. D.; Powell W. S. Metabolism of Linoleic Acid by Prostaglandin Endoperoxide Synthase from Adult and Fetal Blood Vessels. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1983, 754 (1), 57–71. 10.1016/0005-2760(83)90082-6. [DOI] [PubMed] [Google Scholar]
Fuchs D.; Tang X.; Johnsson A.-K.; Dahlén S.-E.; Hamberg M.; Wheelock C. E. Eosinophils Synthesize Trihydroxyoctadecenoic Acids (TriHOMEs) via a 15-Lipoxygenase Dependent Process. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2020, 1865 (4), 158611. 10.1016/j.bbalip.2020.158611. [DOI] [PubMed] [Google Scholar]
Kuhn H.; Banthiya S.; van Leyen K. Mammalian Lipoxygenases and Their Biological Relevance. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2015, 1851 (4), 308–330. 10.1016/j.bbalip.2014.10.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wuest S. J. A.; Crucet M.; Gemperle C.; Loretz C.; Hersberger M. Expression and Regulation of 12/15-Lipoxygenases in Human Primary Macrophages. Atherosclerosis 2012, 225 (1), 121–127. 10.1016/j.atherosclerosis.2012.07.022. [DOI] [PubMed] [Google Scholar]
Jisaka M.; Kim R. B.; Boeglin W. E.; Nanney L. B.; Brash A. R. Molecular Cloning and Functional Expression of a Phorbol Ester-Inducible 8S-Lipoxygenase from Mouse Skin. J. Biol. Chem. 1997, 272 (39), 24410–24416. 10.1074/jbc.272.39.24410. [DOI] [PubMed] [Google Scholar]
Lehnert N.; Solomon E. I. Density-Functional Investigation on the Mechanism of H-Atom Abstraction by Lipoxygenase. J. Biol. Inorg. Chem. 2003, 8 (3), 294–305. 10.1007/s00775-002-0415-6. [DOI] [PubMed] [Google Scholar]
Gardner H. W.; Kleiman R.; Weisleder D. Homolytic Decomposition of Linoleic Acid Hydroperoxide: Identification of Fatty Acid Products. Lipids 1974, 9 (9), 696–706. 10.1007/BF02532178. [DOI] [Google Scholar]
Soberman R. J.; Harper T. W.; Betteridge D.; Lewis R. A.; Austen K. F. Characterization and Separation of the Arachidonic Acid 5-Lipoxygenase and Linoleic Acid Omega-6 Lipoxygenase (Arachidonic Acid 15-Lipoxygenase) of Human Polymorphonuclear Leukocytes. J. Biol. Chem. 1985, 260 (7), 4508–4515. 10.1016/S0021-9258(18)89293-6. [DOI] [PubMed] [Google Scholar]
Baer A. N.; Costello P. B.; Green F. A. Stereospecificity of the Products of the Fatty Acid Oxygenases Derived from Psoriatic Scales. J. Lipid Res. 1991, 32 (2), 341–347. 10.1016/S0022-2275(20)42094-2. [DOI] [PubMed] [Google Scholar]
Zheng Y.; Yin H.; Boeglin W. E.; Elias P. M.; Crumrine D.; Beier D. R.; Brash A. R. Lipoxygenases Mediate the Effect of Essential Fatty Acid in Skin Barrier Formation: A Proposed Role In Releasing Omega-Hydroxyceramide For Construction Of The Corneocyte Lipid Envelope. J. Biol. Chem. 2011, 286 (27), 24046–24056. 10.1074/jbc.M111.251496. [DOI] [PMC free article] [PubMed] [Google Scholar]
Haeggström J. Z.; Funk C. D. Lipoxygenase and Leukotriene Pathways: Biochemistry, Biology, and Roles in Disease. Chem. Rev. 2011, 111 (10), 5866–5898. 10.1021/cr200246d. [DOI] [PubMed] [Google Scholar]
Reinaud O.; Delaforge M.; Boucher J. L.; Rocchiccioli F.; Mansuy D. Oxidative Metabolism of Linoleic Acid by Human Leukocytes. Biochem. Biophys. Res. Commun. 1989, 161 (2), 883–891. 10.1016/0006-291X(89)92682-X. [DOI] [PubMed] [Google Scholar]
Kühn H.; Belkner J.; Wiesner R.; Alder L. Occurrence of 9- and 13-Keto-Octadecadienoic Acid in Biological Membranes Oxygenated by the Reticulocyte Lipoxygenase. Arch. Biochem. Biophys. 1990, 279 (2), 218–224. 10.1016/0003-9861(90)90484-G. [DOI] [PubMed] [Google Scholar]
Bull A. W.; Earles S. M.; Bronstein J. C. Metabolism of Oxidized Linoleic Acid: Distribution of Activity for the Enzymatic Oxidation of 13-Hydroxyoctadecadienoic Acid to 13-Oxooctadecadienoic Acid in Rat Tissues. Prostaglandins 1991, 41 (1), 43–50. 10.1016/0090-6980(91)90103-M. [DOI] [PubMed] [Google Scholar]
Goodfriend T. L.; Ball D. L.; Egan B. M.; Campbell W. B.; Nithipatikom K. Epoxy-Keto Derivative of Linoleic Acid Stimulates Aldosterone Secretion. Hypertension 2004, 43 (2), 358–363. 10.1161/01.HYP.0000113294.06704.64. [DOI] [PubMed] [Google Scholar]
Brash A. R.; Noguchi S.; Boeglin W. E.; Calcutt M. W.; Stec D. F.; Schneider C.; Meyer J. M. Two C18 Hydroxy-Cyclohexenone Fatty Acids from Mammalian Epidermis: Potential Relation to 12R-Lipoxygenase and Covalent Binding of Ceramides. J. Biol. Chem. 2023, 299 (6), 104739. 10.1016/j.jbc.2023.104739. [DOI] [PMC free article] [PubMed] [Google Scholar]
Edin M. L.; Yamanashi H.; Boeglin W. E.; Graves J. P.; DeGraff L. M.; Lih F. B.; Zeldin D. C.; Brash A. R. Epoxide Hydrolase 3 (Ephx3) Gene Disruption Reduces Ceramide Linoleate Epoxide Hydrolysis and Impairs Skin Barrier Function. J. Biol. Chem. 2021, 296, 100198. 10.1074/jbc.RA120.016570. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hamberg M. An Epoxy Alcohol Synthase Pathway in Higher Plants: Biosynthesis of Antifungal Trihydroxy Oxylipins in Leaves of Potato. Lipids 1999, 34 (11), 1131–1142. 10.1007/s11745-999-0464-7. [DOI] [PubMed] [Google Scholar]
Collins J. F.; Hu Z.; Ranganathan P. N.; Feng D.; Garrick L. M.; Garrick M. D.; Browne R. W. Induction of Arachidonate 12-Lipoxygenase (Alox15) in Intestine of Iron-Deficient Rats Correlates with the Production of Biologically Active Lipid Mediators. Am. J. Physiol.-Gastrointest. Liver Physiol. 2008, 294 (4), G948–G962. 10.1152/ajpgi.00274.2007. [DOI] [PubMed] [Google Scholar]
Caligiuri S. P. B.; Love K.; Winter T.; Gauthier J.; Taylor C. G.; Blydt-Hansen T.; Zahradka P.; Aukema H. M. Dietary Linoleic Acid and α-Linolenic Acid Differentially Affect Renal Oxylipins and Phospholipid Fatty Acids in Diet-Induced Obese Rats. J. Nutr. 2013, 143 (9), 1421–1431. 10.3945/jn.113.177360. [DOI] [PubMed] [Google Scholar]
Cossette C.; Patel P.; Anumolu J. R.; Sivendran S.; Lee G. J.; Gravel S.; Graham F. D.; Lesimple A.; Mamer O. A.; Rokach J.; Powell W. S. Human Neutrophils Convert the Sebum-Derived Polyunsaturated Fatty Acid Sebaleic Acid to a Potent Granulocyte Chemoattractant. J. Biol. Chem. 2008, 283 (17), 11234–11243. 10.1074/jbc.M709531200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Arterburn L. M.; Barclay W.; Dangi B.; Flatt J.; Lee J.; Vinjamoori D.. Oxylipins From Stearidonic Acid and Gamma-Linolenic Acid and Methods Of Making and Using The Same. US Patent US 2007/0248586 A1, 2007.
Danielson P. B. The Cytochrome P450 Superfamily: Biochemistry, Evolution and Drug Metabolism in Humans. Curr. Drug Metab. 2002, 3, 561–597. 10.2174/1389200023337054. [DOI] [PubMed] [Google Scholar]
Nebert D. W.; Wikvall K.; Miller W. L. Human Cytochromes P450 in Health and Disease. Philos. Trans. R. Soc. B Biol. Sci. 2013, 368 (1612), 20120431. 10.1098/rstb.2012.0431. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kells P. M.; Ouellet H.; Santos-Aberturas J.; Aparicio J. F.; Podust L. M. Structure of Cytochrome P450 PimD Suggests Epoxidation of the Polyene Macrolide Pimaricin Occurs via a Hydroperoxoferric Intermediate. Chem. Biol. 2010, 17 (8), 841–851. 10.1016/j.chembiol.2010.05.026. [DOI] [PMC free article] [PubMed] [Google Scholar]
Meunier B.; de Visser S. P.; Shaik S. Mechanism of Oxidation Reactions Catalyzed by Cytochrome P450 Enzymes. Chem. Rev. 2004, 104 (9), 3947–3980. 10.1021/cr020443g. [DOI] [PubMed] [Google Scholar]
Morisseau C.; Kodani S. D.; Kamita S. G.; Yang J.; Lee K. S. S.; Hammock B. D. Relative Importance of Soluble and Microsomal Epoxide Hydrolases for the Hydrolysis of Epoxy-Fatty Acids in Human Tissues. Int. J. Mol. Sci. 2021, 22 (9), 4993. 10.3390/ijms22094993. [DOI] [PMC free article] [PubMed] [Google Scholar]
Moran J. H.; Mitchell L. A.; Bradbury J. A.; Qu W.; Zeldin D. C.; Schnellmann R. G.; Grant D. F. Analysis of the Cytotoxic Properties of Linoleic Acid Metabolites Produced by Renal and Hepatic P450s. Toxicol. Appl. Pharmacol. 2000, 168 (3), 268–279. 10.1006/taap.2000.9053. [DOI] [PubMed] [Google Scholar]
Draper A. J.; Hammock B. D. Identification of CYP2C9 as a Human Liver Microsomal Linoleic Acid Epoxygenase. Arch. Biochem. Biophys. 2000, 376 (1), 199–205. 10.1006/abbi.2000.1705. [DOI] [PubMed] [Google Scholar]
Bylund J.; Kunz T.; Valmsen K.; Oliw E. H. Cytochromes P450 with Bisallylic Hydroxylation Activity on Arachidonic and Linoleic Acids Studied with Human Recombinant Enzymes and with Human and Rat Liver Microsomes. J. Pharmacol. Exp. Ther. 1998, 284, 51–60. [PubMed] [Google Scholar]
Laethem R. M.; Balazy M.; Koop D. R. Epoxidation of C18 Unsaturated Fatty Acids by Cytochromes P4502C2 and P4502CAA. Drug Metab. Dispos. 1996, 24, 664–668. [PubMed] [Google Scholar]
Williamson K. C.; Morisseau C.; Maxwell J. E.; Hammock B. D. Regio- and Enantioselective Hydrolysis of Phenyloxiranes Catalyzed by Soluble Epoxide Hydrolase. Tetrahedron Asymmetry 2000, 11 (22), 4451–4462. 10.1016/S0957-4166(00)00437-7. [DOI] [Google Scholar]
Teder T.; Boeglin W. E.; Brash A. R. Lipoxygenase-Catalyzed Transformation of Epoxy Fatty Acids to Hydroxy-Endoperoxides: A Potential P450 and Lipoxygenase Interaction. J. Lipid Res. 2014, 55 (12), 2587–2596. 10.1194/jlr.M054072. [DOI] [PMC free article] [PubMed] [Google Scholar]
Tsikas D.; Sawa M.; Brunner G.; Gutzki F.-M.; Meyer H. H.; Frölich J. C. Gas Chromatography-Mass Spectrometry of cis-9,10-Epoxyoctadecanoic Acid (cis-EODA): I. Direct Evidence for cis-EODA Formation from Oleic Acid Oxidation by Liver Microsomes and Isolated Hepatocytes. J. Chromatogr. B 2003, 784 (2), 351–365. 10.1016/S1570-0232(02)00821-8. [DOI] [PubMed] [Google Scholar]
Thum T.; Batkai S.; Malinski P. G.; Becker T.; Mevius I.; Klempnauer J.; Meyer H. H.; Frölich J. C.; Borlak J.; Tsikas D. Measurement and Diagnostic Use of Hepatic Cytochrome P450 Metabolism of Oleic Acid in Liver Disease. Liver Int. 2010, 30 (8), 1181–1188. 10.1111/j.1478-3231.2010.02310.x. [DOI] [PubMed] [Google Scholar]
Oliw E. H. Bis-Allylic Hydroxylation of Linoleic Acid and Arachidonic Acid by Human Hepatic Monooxygenases. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1993, 1166 (2), 258–263. 10.1016/0005-2760(93)90106-J. [DOI] [PubMed] [Google Scholar]
Adas F.; Salaün J. P.; Berthou F.; Picart D.; Simon B.; Amet Y. Requirement for ω and (ω-1)-Hydroxylations of Fatty Acids by Human Cytochromes P450 2E1 and 4A11. J. Lipid Res. 1999, 40 (11), 1990–1997. 10.1016/S0022-2275(20)32422-6. [DOI] [PubMed] [Google Scholar]
Nguyen X.; Wang M.-H.; Reddy K. M.; Falck J. R.; Schwartzman M. L. Kinetic Profile of the Rat CYP4A Isoforms: Arachidonic Acid Metabolism and Isoform-Specific Inhibitors. Am. J. Physiol.-Regul. Integr. Comp. Physiol. 1999, 276 (6), R1691–R1700. 10.1152/ajpregu.1999.276.6.R1691. [DOI] [PubMed] [Google Scholar]
Chuang S. S.; Helvig C.; Taimi M.; Ramshaw H. A.; Collop A. H.; Amad M.; White J. A.; Petkovich M.; Jones G.; Korczak B. CYP2U1, a Novel Human Thymus- and Brain-Specific Cytochrome P450, Catalyzes ω- and (ω-1)-Hydroxylation of Fatty Acids. J. Biol. Chem. 2004, 279 (8), 6305–6314. 10.1074/jbc.M311830200. [DOI] [PubMed] [Google Scholar]
Bjorkhem I.; Danielsson H. Omega- and (Omega-1)-Oxidation of Fatty Acids by Rat Liver Microsomes. Eur. J. Biochem. 1970, 17 (3), 450–459. 10.1111/j.1432-1033.1970.tb01186.x. [DOI] [PubMed] [Google Scholar]
Le Quéré V.; Plée-Gautier E.; Potin P.; Madec S.; Salaün J.-P. Human CYP4F3s Are the Main Catalysts in the Oxidation of Fatty Acid Epoxides. J. Lipid Res. 2004, 45 (8), 1446–1458. 10.1194/jlr.M300463-JLR200. [DOI] [PubMed] [Google Scholar]
Dhar M.; Sepkovic D. W.; Hirani V.; Magnusson R. P.; Lasker J. M. Omega Oxidation of 3-Hydroxy Fatty Acids by the Human CYP4F Gene Subfamily Enzyme CYP4F11. J. Lipid Res. 2008, 49 (3), 612–624. 10.1194/jlr.M700450-JLR200. [DOI] [PubMed] [Google Scholar]
Aggarwal N.; Kitano S.; Puah G. R. Y.; Kittelmann S.; Hwang I. Y.; Chang M. W. Microbiome and Human Health: Current Understanding, Engineering, and Enabling Technologies. Chem. Rev. 2023, 123 (1), 31–72. 10.1021/acs.chemrev.2c00431. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schoeler M.; Caesar R. Dietary Lipids, Gut Microbiota and Lipid Metabolism. Rev. Endocr. Metab. Disord. 2019, 20 (4), 461–472. 10.1007/s11154-019-09512-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schoeler M.; Ellero-Simatos S.; Birkner T.; Mayneris-Perxachs J.; Olsson L.; Brolin H.; Loeber U.; Kraft J. D.; Polizzi A.; Martí-Navas M.; Puig J.; Moschetta A.; Montagner A.; Gourdy P.; Heymes C.; Guillou H.; Tremaroli V.; Fernández-Real J. M.; Forslund S. K.; Burcelin R.; Caesar R. The Interplay between Dietary Fatty Acids and Gut Microbiota Influences Host Metabolism and Hepatic Steatosis. Nat. Commun. 2023, 14 (1), 5329. 10.1038/s41467-023-41074-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zhang Q.; Zhang L.; Chen C.; Li P.; Lu B. The Gut Microbiota-Artery Axis: A Bridge between Dietary Lipids and Atherosclerosis?. Prog. Lipid Res. 2023, 89, 101209. 10.1016/j.plipres.2022.101209. [DOI] [PubMed] [Google Scholar]
Jonsson A. L.; Bäckhed F. Role of Gut Microbiota in Atherosclerosis. Nat. Rev. Cardiol. 2017, 14 (2), 79–87. 10.1038/nrcardio.2016.183. [DOI] [PubMed] [Google Scholar]
Fan C.; Xu J.; Tong H.; Fang Y.; Chen Y.; Lin Y.; Chen R.; Chen F.; Wu G. Gut-Brain Communication Mediates the Impact of Dietary Lipids on Cognitive Capacity. Food Funct. 2024, 15 (4), 1803–1824. 10.1039/D3FO05288E. [DOI] [PubMed] [Google Scholar]
Özçam M.; Lynch S. V. The Gut-Airway Microbiome Axis in Health and Respiratory Diseases. Nat. Rev. Microbiol. 2024, 22, 492. 10.1038/s41579-024-01048-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
Budden K. F.; Gellatly S. L.; Wood D. L. A.; Cooper M. A.; Morrison M.; Hugenholtz P.; Hansbro P. M. Emerging Pathogenic Links between Microbiota and the Gut-Lung Axis. Nat. Rev. Microbiol. 2017, 15 (1), 55–63. 10.1038/nrmicro.2016.142. [DOI] [PubMed] [Google Scholar]
Pinckard K. M.; Shettigar V. K.; Wright K. R.; Abay E.; Baer L. A.; Vidal P.; Dewal R. S.; Das D.; Duarte-Sanmiguel S.; Hernández-Saavedra D.; Arts P. J.; Lehnig A. C.; Bussberg V.; Narain N. R.; Kiebish M. A.; Yi F.; Sparks L. M.; Goodpaster B. H.; Smith S. R.; Pratley R. E.; Lewandowski E. D.; Raman S. V.; Wold L. E.; Gallego-Perez D.; Coen P. M.; Ziolo M. T.; Stanford K. I. A Novel Endocrine Role for the BAT-Released Lipokine 12,13-DiHOME to Mediate Cardiac Function. Circulation 2021, 143 (2), 145–159. 10.1161/CIRCULATIONAHA.120.049813. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zimmer B.; Angioni C.; Osthues T.; Toewe A.; Thomas D.; Pierre S. C.; Geisslinger G.; Scholich K.; Sisignano M. The Oxidized Linoleic Acid Metabolite 12,13-DiHOME Mediates Thermal Hyperalgesia during Inflammatory Pain. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2018, 1863 (7), 669–678. 10.1016/j.bbalip.2018.03.012. [DOI] [PubMed] [Google Scholar]
Kishino S.; Takeuchi M.; Park S.-B.; Hirata A.; Kitamura N.; Kunisawa J.; Kiyono H.; Iwamoto R.; Isobe Y.; Arita M.; Arai H.; Ueda K.; Shima J.; Takahashi S.; Yokozeki K.; Shimizu S.; Ogawa J. Polyunsaturated Fatty Acid Saturation by Gut Lactic Acid Bacteria Affecting Host Lipid Composition. Proc. Natl. Acad. Sci. U. S. A. 2013, 110 (44), 17808–17813. 10.1073/pnas.1312937110. [DOI] [PMC free article] [PubMed] [Google Scholar]
Polan C. E.; McNeill J. J.; Tove S. B. Biohydrogenation of Unsaturated Fatty Acids by Rumen Bacteria. J. Bacteriol. 1964, 88 (4), 1056–1064. 10.1128/jb.88.4.1056-1064.1964. [DOI] [PMC free article] [PubMed] [Google Scholar]
Mir P. S.; McAllister T. A.; Scott S.; Aalhus J.; Baron V.; McCartney D.; Charmley E.; Goonewardene L.; Basarab J.; Okine E.; Weselake R. J.; Mir Z. Conjugated Linoleic Acid-Enriched Beef Production. Am. J. Clin. Nutr. 2004, 79, 1207S–1211S. 10.1093/ajcn/79.6.1207S. [DOI] [PubMed] [Google Scholar]
Ferlay A.; Bernard L.; Meynadier A.; Malpuech-Brugère C. Production of Trans and Conjugated Fatty Acids in Dairy Ruminants and Their Putative Effects on Human Health: A Review. Biochimie 2017, 141, 107–120. 10.1016/j.biochi.2017.08.006. [DOI] [PubMed] [Google Scholar]
Lee K. W.; Lee H. J.; Cho H. Y.; Kim Y. J. Role of the Conjugated Linoleic Acid in the Prevention of Cancer. Crit. Rev. Food Sci. Nutr. 2005, 45 (2), 135–144. 10.1080/10408690490911800. [DOI] [PubMed] [Google Scholar]
Wahle K. W. J.; Heys S. D.; Rotondo D. Conjugated Linoleic Acids: Are They Beneficial or Detrimental to Health?. Prog. Lipid Res. 2004, 43 (6), 553–587. 10.1016/j.plipres.2004.08.002. [DOI] [PubMed] [Google Scholar]
Dhar Dubey K. K.; Sharma G.; Kumar A. Conjugated Linolenic Acids: Implication in Cancer. J. Agric. Food Chem. 2019, 67 (22), 6091–6101. 10.1021/acs.jafc.9b01379. [DOI] [PubMed] [Google Scholar]
Tanaka T.; Hosokawa M.; Yasui Y.; Ishigamori R.; Miyashita K. Cancer Chemopreventive Ability of Conjugated Linolenic Acids. Int. J. Mol. Sci. 2011, 12 (11), 7495–7509. 10.3390/ijms12117495. [DOI] [PMC free article] [PubMed] [Google Scholar]
Scollan N. D.; Price E. M.; Morgan S. A.; Huws S. A.; Shingfield K. J. Can We Improve the Nutritional Quality of Meat?. Proc. Nutr. Soc. 2017, 76 (4), 603–618. 10.1017/S0029665117001112. [DOI] [PubMed] [Google Scholar]
Ponnampalam E.; Mann N.; Sinclair A. Effect of Feeding Systems on Omega-3 Fatty Acids, Conjugated Linoleic Acid and Trans Fatty Acids in Australian Beef Cuts: Potential Impact on Humnan Health. Asia Pac. J. Clin. Nutr. 2006, 15, 21–29. [PubMed] [Google Scholar]
Vahmani P.; Ponnampalam E. N.; Kraft J.; Mapiye C.; Bermingham E. N.; Watkins P. J.; Proctor S. D.; Dugan M. E. R. Bioactivity and Health Effects of Ruminant Meat Lipids. Invited Review. Meat Sci. 2020, 165, 108114. 10.1016/j.meatsci.2020.108114. [DOI] [PubMed] [Google Scholar]
Hira T.; Ogasawara S.; Yahagi A.; Kamachi M.; Li J.; Nishimura S.; Sakaino M.; Yamashita T.; Kishino S.; Ogawa J.; Hara H. Novel Mechanism of Fatty Acid Sensing in Enteroendocrine Cells: Specific Structures in Oxo-Fatty Acids Produced by Gut Bacteria Are Responsible for CCK Secretion in STC-1 Cells via GPR40. Mol. Nutr. Food Res. 2018, 62 (19), 1800146. 10.1002/mnfr.201800146. [DOI] [PubMed] [Google Scholar]
Hirata A.; Kishino S.; Park S.-B.; Takeuchi M.; Kitamura N.; Ogawa J. A Novel Unsaturated Fatty Acid Hydratase toward C16 to C22 Fatty Acids from Lactobacillus acidophilus. J. Lipid Res. 2015, 56 (7), 1340–1350. 10.1194/jlr.M059444. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kishimoto N.; Yamamoto I.; Toraishi K.; Yoshioka S.; Saito K.; Masuda H.; Fujita T. Two Distinct Pathways for the Formation of Hydroxy FA from Linoleic Acid by Lactic Acid Bacteria. Lipids 2003, 38 (12), 1269–1274. 10.1007/s11745-003-1188-4. [DOI] [PubMed] [Google Scholar]
Takeuchi M.; Kishino S.; Tanabe K.; Hirata A.; Park S.-B.; Shimizu S.; Ogawa J. Hydroxy Fatty Acid Production by Pediococcus sp. Eur. J. Lipid Sci. Technol. 2013, 115 (4), 386–393. 10.1002/ejlt.201200414. [DOI] [Google Scholar]
Hudson J. A.; Morvan B.; Joblin K. N. Hydration of Linoleic Acid by Bacteria Isolated from Ruminants. FEMS Microbiol. Lett. 1998, 169 (2), 277–282. 10.1111/j.1574-6968.1998.tb13329.x. [DOI] [PubMed] [Google Scholar]
Koritala S.; Hou C. T.; Hesseltine C. W.; Bagby M. O. Microbial Conversion of Oleic Acid to 10-Hydroxystearic Acid. Appl. Microbiol. Biotechnol. 1989, 32 (3), 299–304. 10.1007/BF00184978. [DOI] [Google Scholar]
Hou C. T. Conversion of Linoleic Acid to 10-Hydroxy-12(Z)-Octadecenoic Acid by Flavobacterium sp. J. Am. Oil Chem. Soc. 1994, 71 (9), 975–978. 10.1007/BF02542264. [DOI] [Google Scholar]
Hou C. T. New Bioactive Fatty Acids. Asia Pac. J. Clin. Nutr. 2008, 17, 192–195. [PubMed] [Google Scholar]
Hou C. T. A Novel Compound, 12,13,17-Trihydroxy-9(Z)-Octadecenoic Acid, from Linoleic Acid by a New Microbial Isolate Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 1996, 73 (11), 1359–1362. 10.1007/BF02523497. [DOI] [Google Scholar]
Iwasaki Y.; Brown W.; Hou C. T. Biosynthetic Pathway of Diepoxy Bicyclic FA from Linoleic Acid by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2002, 79 (4), 369–372. 10.1007/s11746-002-0490-x. [DOI] [Google Scholar]
Gardner H. W.; Hou C. T.; Weisleder D.; Brown W. Biotransformation of Linoleic Acid by Clavibacter sp. ALA2: Heterocyclic and Heterobicyclic Fatty Acids. Lipids 2000, 35 (10), 1055–1060. 10.1007/s11745-000-0618-7. [DOI] [PubMed] [Google Scholar]
Hosokawa M.; Hou C. T.; Weisleder D.; Brown W. Biosynthesis of Tetrahydrofuranyl Fatty Acids from Linoleic Acid by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2003, 80 (2), 145–149. 10.1007/s11746-003-0667-3. [DOI] [Google Scholar]
Hou C. T.; Gardner H. W.; Brown W. 12,13,16-Trihydroxy-9(Z)-Octadecenoic Acid, a Possible Intermediate in the Bioconversion of Linoleic Acid to Tetrahydrofuranyl Fatty Acids by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2001, 78 (11), 1167–1169. 10.1007/s11746-001-0407-8. [DOI] [Google Scholar]
Hosokawa M.; Hou C. T.; Weisleder D. Production of Novel Tetrahydroxyfuranyl Fatty Acids from α-Linolenic Acid by Clavibacter sp. Strain ALA2. Appl. Environ. Microbiol. 2003, 69 (7), 3868–3873. 10.1128/AEM.69.7.3868-3873.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hosokawa M.; Hou C. T.; Weisleder D. Bioconversion of N–3 and N–6 PUFA by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2003, 80 (11), 1085–1091. 10.1007/s11746-003-0824-8. [DOI] [Google Scholar]
Andreou A.; Göbel C.; Hamberg M.; Feussner I. A Bisallylic Mini-Lipoxygenase from Cyanobacterium Cyanothece sp. That Has an Iron as Cofactor. J. Biol. Chem. 2010, 285 (19), 14178–14186. 10.1074/jbc.M109.094771. [DOI] [PMC free article] [PubMed] [Google Scholar]
Gao B.; Boeglin W. E.; Brash A. R. Omega-3 Fatty Acids Are Oxygenated at the n-7 Carbon by the Lipoxygenase Domain of a Fusion Protein in the Cyanobacterium Acaryochloris marina. Biochim. Biophys. Acta 2010, 1801 (1), 58–63. 10.1016/j.bbalip.2009.09.004. [DOI] [PMC free article] [PubMed] [Google Scholar]
el-Sharkawy S. H.; Yang W.; Dostal L.; Rosazza J. P. Microbial Oxidation of Oleic Acid. Appl. Environ. Microbiol. 1992, 58 (7), 2116–2122. 10.1128/aem.58.7.2116-2122.1992. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hou C. T. Production of Hydroxy Fatty Acids from Unsaturated Fatty Acids by Flavobacterium sp. DS5 Hydratase, a C-10 Positional- and Cis Unsaturation-Specific Enzyme. J. Am. Oil Chem. Soc. 1995, 72 (11), 1265–1270. 10.1007/BF02546197. [DOI] [Google Scholar]
Hou C. T. Production of 10-Ketostearic Acid from Oleic Acid by Flavobacterium sp. Strain DS5 (NRRL B-14859). Appl. Environ. Microbiol. 1994, 60 (10), 3760–3763. 10.1128/aem.60.10.3760-3763.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hou C. T.; Bagby M. O.; Plattner R. D.; Koritala S. A Novel Compound, 7,10-Dihydroxy-8(E)-Octadecenoic Acid from Oleic Acid by Bioconversion. J. Am. Oil Chem. Soc. 1991, 68 (2), 99–101. 10.1007/BF02662326. [DOI] [Google Scholar]
Hou C. T.; Bagby M. O. 10-Hydroxy-8(Z)-Octadecenoic Acid, an Intermediate in the Bioconversion of Oleic Acid to 7,10-Dihydroxy-8(E)-Octadecenoic Acid. J. Ind. Microbiol. 1992, 9 (2), 103–107. 10.1007/BF01569740. [DOI] [Google Scholar]
Guerrero A.; Casals I.; Busquets M.; Leon Y.; Manresa A. Oxydation of Oleic Acid to (E)-10-Hydroperoxy-8-Octadecenoic and (E)-10-Hydroxy-8-Octadecenoic Acids by Pseudomonas sp. 42A2. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1997, 1347 (1), 75–81. 10.1016/S0005-2760(97)00056-8. [DOI] [PubMed] [Google Scholar]
Busquets M.; Deroncelé V.; Vidal-Mas J.; Rodríguez E.; Guerrero A.; Manresa A. Isolation and Characterization of a Lipoxygenase from Pseudomonas 42A2 Responsible for the Biotransformation of Oleic Acid into (S)-(E)-10-Hydroxy-8-Octadecenoic Acid. Antonie Van Leeuwenhoek 2004, 85 (2), 129–139. 10.1023/B:ANTO.0000020152.15440.65. [DOI] [PubMed] [Google Scholar]
Bhattarai S.; Niraula N. P.; Sohng J. K.; Oh T.-J. In-Silico and In-Vitro Based Studies of Streptomyces peucetius CYP107N3 for Oleic Acid Epoxidation. BMB Rep. 2012, 45 (12), 736–741. 10.5483/BMBRep.2012.45.12.080. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hintikka E.; Holopainen R.; Asola A.; Jestoi M.; Peitzsch M.; Kalso S.; Larsson L.; Reijula K.; Tuomi T. Mycotoxins in the Ventilation Systems of Four Schools in Finland. World Mycotoxin J. 2009, 2 (4), 369–379. 10.3920/WMJ2009.1155. [DOI] [Google Scholar]
Niu M.; Keller N. P. Co-opting Oxylipin Signals in Microbial Disease. Cell. Microbiol. 2019, 21 (6), e13025 10.1111/cmi.13025. [DOI] [PMC free article] [PubMed] [Google Scholar]
Tsitsigiannis D. I.; Keller N. P. Oxylipins as Developmental and Host-Fungal Communication Signals. Trends Microbiol. 2007, 15 (3), 109–118. 10.1016/j.tim.2007.01.005. [DOI] [PubMed] [Google Scholar]
Brodhun F.; Feussner I. Oxylipins in Fungi. FEBS J. 2011, 278 (7), 1047–1063. 10.1111/j.1742-4658.2011.08027.x. [DOI] [PubMed] [Google Scholar]
Christensen S. A.; Kolomiets M. V. The Lipid Language of Plant-Fungal Interactions. Fungal Genet. Biol. 2011, 48 (1), 4–14. 10.1016/j.fgb.2010.05.005. [DOI] [PubMed] [Google Scholar]
Wurzenberger M.; Grosch W. Origin of the Oxygen in the Products of the Enzymatic Cleavage Reaction of Linoleic Acid to 1-Octen-3-Ol and 10-Oxo-Trans-8-Decenoic Acid in Mushrooms (Psalliota bispora). Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1984, 794 (1), 18–24. 10.1016/0005-2760(84)90292-3. [DOI] [Google Scholar]
Wurzenberger M.; Grosch W. Stereochemistry of the Cleavage of the 10-Hydroperoxide Isomer of Linoleic Acid to 1-Octen-3-Ol by a Hydroperoxide Lyase from Mushrooms (Psalliota bispora). Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1984, 795 (1), 163–165. 10.1016/0005-2760(84)90117-6. [DOI] [Google Scholar]
Wurzenberger M.; Grosch W. The Formation of 1-Octen-3-Ol from the 10-Hydroperoxide Isomer of Linoleic Acid by a Hydroperoxide Lyase in Mushrooms (Psalliota bispora). Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1984, 794 (1), 25–30. 10.1016/0005-2760(84)90293-5. [DOI] [Google Scholar]
Wurzenberger M.; Grosch W. Enzymic Oxidation of Linolenic Acid to 1,Z-5-Octadien-3-Ol, Z-2,Z-5-Octadien-1-Ol and 10-Oxo-E-8-Decenoic Acid by a Protein Fraction from Mushrooms (Psalliota bispora). Lipids 1986, 21 (4), 261–266. 10.1007/BF02536408. [DOI] [PubMed] [Google Scholar]
Brash A. R.; Niraula N. P.; Boeglin W. E.; Mashhadi Z. An Ancient Relative of Cyclooxygenase in Cyanobacteria Is a Linoleate 10S-Dioxygenase That Works in Tandem with a Catalase-Related Protein with Specific 10S-Hydroperoxide Lyase Activity. J. Biol. Chem. 2014, 289 (19), 13101–13111. 10.1074/jbc.M114.555904. [DOI] [PMC free article] [PubMed] [Google Scholar]
Teshima T.; Funai R.; Nakazawa T.; Ito J.; Utsumi T.; Kakumyan P.; Mukai H.; Yoshiga T.; Murakami R.; Nakagawa K.; Honda Y.; Matsui K. Coprinopsis Cinerea Dioxygenase Is an Oxygenase Forming 10(S)-Hydroperoxide of Linoleic Acid, Essential for Mushroom Alcohol, 1-Octen-3-Ol, Synthesis. J. Biol. Chem. 2022, 298 (11), 102507. 10.1016/j.jbc.2022.102507. [DOI] [PMC free article] [PubMed] [Google Scholar]
Oliw E. H. Fatty Acid Dioxygenase-Cytochrome P450 Fusion Enzymes of Filamentous Fungal Pathogens. Fungal Genet. Biol. 2021, 157, 103623. 10.1016/j.fgb.2021.103623. [DOI] [PubMed] [Google Scholar]
Wadman M. W.; van Zadelhoff G.; Hamberg M.; Visser T.; Veldink G. A.; Vliegenthart J. F. G. Conversion of Linoleic Acid into Novel Oxylipins by the Mushroom Agaricus bisporus. Lipids 2005, 40 (11), 1163–1170. 10.1007/s11745-005-1481-2. [DOI] [PubMed] [Google Scholar]
Hoffmann I.; Jernerén F.; Garscha U.; Oliw E. H. Expression of 5,8-LDS of Aspergillus fumigatus and Its Dioxygenase Domain: A Comparison with 7,8-LDS, 10-Dioxygenase, and Cyclooxygenase. Arch. Biochem. Biophys. 2011, 506 (2), 216–222. 10.1016/j.abb.2010.11.022. [DOI] [PubMed] [Google Scholar]
Garscha U.; Jernerén F.; Chung D.; Keller N. P.; Hamberg M.; Oliw E. H. Identification of Dioxygenases Required for Aspergillus Development. J. Biol. Chem. 2007, 282 (48), 34707–34718. 10.1074/jbc.M705366200. [DOI] [PubMed] [Google Scholar]
Su C.; Sahlin M.; Oliw E. H. A Protein Radical and Ferryl Intermediates Are Generated by Linoleate Diol Synthase, a Ferric Hemeprotein with Dioxygenase and Hydroperoxide Isomerase Activities. J. Biol. Chem. 1998, 273 (33), 20744–20751. 10.1074/jbc.273.33.20744. [DOI] [PubMed] [Google Scholar]
Brodowsky I. D.; Oliw E. H. Metabolism of 18:2(n – 6), 18:3(n – 3), 20:4(n – 6) and 20:5(n – 3) by the Fungus Gaeumannomyces graminis: Identification of Metabolites Formed by 8-Hydroxylation and by W2 and W3 Oxygenation. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1992, 1124 (1), 59–65. 10.1016/0005-2760(92)90126-G. [DOI] [PubMed] [Google Scholar]
Cristea M.; Osbourn A. E.; Oliw E. H. Linoleate Diol Synthase of the Rice Blast Fungus Magnaporthe grisea. Lipids 2003, 38 (12), 1275–1280. 10.1007/s11745-003-1189-3. [DOI] [PubMed] [Google Scholar]
Jernerén F.; Garscha U.; Hoffmann I.; Hamberg M.; Oliw E. H. Reaction Mechanism of 5,8-Linoleate Diol Synthase, 10R-Dioxygenase, and 8,11-Hydroperoxide Isomerase of Aspergillus clavatus. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2010, 1801 (4), 503–507. 10.1016/j.bbalip.2009.12.012. [DOI] [PubMed] [Google Scholar]
Shin K.-C.; Seo M.-J.; Oh D.-K. Characterization of a Novel 8R,11S-Linoleate Diol Synthase from Penicillium chrysogenum by Identification of Its Enzymatic Products. J. Lipid Res. 2016, 57 (2), 207–218. 10.1194/jlr.M061341. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hoffmann I.; Jernerén F.; Oliw E. H. Epoxy Alcohol Synthase of the Rice Blast Fungus Represents a Novel Subfamily of Dioxygenase-Cytochrome P450 Fusion Enzymes. J. Lipid Res. 2014, 55 (10), 2113–2123. 10.1194/jlr.M051755. [DOI] [PMC free article] [PubMed] [Google Scholar]
Su C.; Oliw E. H. Manganese Lipoxygenase: Purification And Characterization. J. Biol. Chem. 1998, 273 (21), 13072–13079. 10.1074/jbc.273.21.13072. [DOI] [PubMed] [Google Scholar]
Hamberg M.; Su C.; Oliw E. Manganese Lipoxygenase. Discovery Of A Bis-Allylic Hydroperoxide As Product And Intermediate In A Lipoxygenase Reaction. J. Biol. Chem. 1998, 273 (21), 13080–13088. 10.1074/jbc.273.21.13080. [DOI] [PubMed] [Google Scholar]
Wennman A.; Oliw E. H. Secretion of Two Novel Enzymes, Manganese 9S-Lipoxygenase and Epoxy Alcohol Synthase, by the Rice Pathogen Magnaporthe salvinii. J. Lipid Res. 2013, 54 (3), 762–775. 10.1194/jlr.M033787. [DOI] [PMC free article] [PubMed] [Google Scholar]
Brodhun F.; Cristobal-Sarramian A.; Zabel S.; Newie J.; Hamberg M.; Feussner I. An Iron 13S-Lipoxygenase with an α-Linolenic Acid Specific Hydroperoxidase Activity from Fusarium oxysporum. PLoS One 2013, 8 (5), e64919 10.1371/journal.pone.0064919. [DOI] [PMC free article] [PubMed] [Google Scholar]
Teder T.; Boeglin W. E.; Schneider C.; Brash A. R. A Fungal Catalase Reacts Selectively with the 13S Fatty Acid Hydroperoxide Products of the Adjacent Lipoxygenase Gene and Exhibits 13S-Hydroperoxide-Dependent Peroxidase Activity. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2017, 1862 (7), 706–715. 10.1016/j.bbalip.2017.03.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
González-Benjumea A.; Carro J.; Renau-Mínguez C.; Linde D.; Fernández-Fueyo E.; Gutiérrez A.; Martínez A. T. Fatty Acid Epoxidation by Collariella virescens Peroxygenase and Heme-Channel Variants. Catal. Sci. Technol. 2020, 10 (3), 717–725. 10.1039/C9CY02332A. [DOI] [Google Scholar]
Yin H.; Xu L.; Porter N. A. Free Radical Lipid Peroxidation: Mechanisms and Analysis. Chem. Rev. 2011, 111 (10), 5944–5972. 10.1021/cr200084z. [DOI] [PubMed] [Google Scholar]
Viswanathan S.; Hammock B. D.; Newman J. W.; Meerarani P.; Toborek M.; Hennig B. Involvement of CYP 2C9 in Mediating the Proinflammatory Effects of Linoleic Acid in Vascular Endothelial Cells. J. Am. Coll. Nutr. 2003, 22 (6), 502–510. 10.1080/07315724.2003.10719328. [DOI] [PubMed] [Google Scholar]
Kellogg E.; Fridovich I. Superoxide, Hydrogen Peroxide, and Singlet Oxygen in Lipid Peroxidation by a Xanthine Oxidase System. J. Biol. Chem. 1975, 250 (22), 8812–8817. 10.1016/S0021-9258(19)40745-X. [DOI] [PubMed] [Google Scholar]
Li Y.; Mouche S.; Sajic T.; Veyrat-Durebex C.; Supale R.; Pierroz D.; Ferrari S.; Negro F.; Hasler U.; Feraille E.; Moll S.; Meda P.; Deffert C.; Montet X.; Krause K.-H.; Szanto I. Deficiency in the NADPH Oxidase 4 Predisposes towards Diet-Induced Obesity. Int. J. Obes. 2012, 36 (12), 1503–1513. 10.1038/ijo.2011.279. [DOI] [PubMed] [Google Scholar]
Kurutas E. B. The Importance of Antioxidants Which Play the Role in Cellular Response against Oxidative/Nitrosative Stress: Current State. Nutr. J. 2015, 15 (1), 71. 10.1186/s12937-016-0186-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
Oh J. M.; Choi J. M.; Lee J. Y.; Oh S. J.; Kim B. H.; Kim S. K. Role of NADPH Oxidase-4 in Saturated Fatty Acid-Induced Insulin Resistance in SK-Hep-1 Cells. Food Chem. Toxicol. 2014, 63, 128–135. 10.1016/j.fct.2013.10.049. [DOI] [PubMed] [Google Scholar]
Hatanaka E.; Dermargos A.; Hirata A. E.; Vinolo M. A. R.; Carpinelli A. R.; Newsholme P.; Armelin H. A.; Curi R. Oleic, Linoleic and Linolenic Acids Increase ROS Production by Fibroblasts via NADPH Oxidase Activation. PLoS One 2013, 8 (4), e58626 10.1371/journal.pone.0058626. [DOI] [PMC free article] [PubMed] [Google Scholar]
Badwey J. A.; Curnutte J. T.; Karnovsky M. L. cis-Polyunsaturated Fatty Acids Induce High Levels of Superoxide Production by Human Neutrophils. J. Biol. Chem. 1981, 256 (24), 12640–12643. 10.1016/S0021-9258(18)42939-0. [DOI] [PubMed] [Google Scholar]
Tyurina Y. Y.; Poloyac S. M.; Tyurin V. A.; Kapralov A. A.; Jiang J.; Anthonymuthu T. S.; Kapralova V. I.; Vikulina A. S.; Jung M.-Y.; Epperly M. W.; Mohammadyani D.; Klein-Seetharaman J.; Jackson T. C.; Kochanek P. M.; Pitt B. R.; Greenberger J. S.; Vladimirov Y. A.; Bayır H.; Kagan V. E. A Mitochondrial Pathway for Biosynthesis of Lipid Mediators. Nat. Chem. 2014, 6 (6), 542–552. 10.1038/nchem.1924. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sies H.; Berndt C.; Jones D. P. Oxidative Stress. Annu. Rev. Biochem. 2017, 86 (1), 715–748. 10.1146/annurev-biochem-061516-045037. [DOI] [PubMed] [Google Scholar]
Gutteridge J. M. C.; Bannister J. V. Copper + Zinc and Manganese Superoxide Dismutases Inhibit Deoxyribose Degradation by the Superoxide-Driven Fenton Reaction at Two Different Stages. Implications for the Redox States of Copper and Manganese. Biochem. J. 1986, 234 (1), 225–228. 10.1042/bj2340225. [DOI] [PMC free article] [PubMed] [Google Scholar]
Pryor W. A. Oxy-Radicals and Related Species: Their Formation, Lifetimes, and Reactions. Annu. Rev. Physiol. 1986, 48, 657–667. 10.1146/annurev.ph.48.030186.003301. [DOI] [PubMed] [Google Scholar]
Wilcox A. L.; Marnett L. J. Polyunsaturated Fatty Acid Alkoxyl Radicals Exist as Carbon-Centered Epoxyallylic Radicals: A Key Step in Hydroperoxide-Amplified Lipid Peroxidation. Chem. Res. Toxicol. 1993, 6 (4), 413–416. 10.1021/tx00034a003. [DOI] [PubMed] [Google Scholar]
Gardner H. W. Oxygen Radical Chemistry of Polyunsaturated Fatty Acids. Free Radic. Biol. Med. 1989, 7 (1), 65–86. 10.1016/0891-5849(89)90102-0. [DOI] [PubMed] [Google Scholar]
Spiteller G. Linoleic Acid Peroxidation-the Dominant Lipid Peroxidation Process in Low Density Lipoprotein-and Its Relationship to Chronic Diseases. Chem. Phys. Lipids 1998, 95 (2), 105–162. 10.1016/S0009-3084(98)00091-7. [DOI] [PubMed] [Google Scholar]
Kaplan E.; Ansari K. Reduction of Polyunsaturated Fatty Acid Hydroperoxides by Human Brain Glutathione Peroxidase. Lipids 1984, 19 (10), 784–789. 10.1007/BF02534472. [DOI] [PubMed] [Google Scholar]
Christophersen B. O. Formation of Monohydroxy-Polyenic Fatty Acids from Lipid Peroxides by a Glutathione Peroxidase. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1968, 164 (1), 35–46. 10.1016/0005-2760(68)90068-4. [DOI] [PubMed] [Google Scholar]
Little C.; O’Brien P. J. An Intracellular GSH-Peroxidase with a Lipid Peroxide Substrate. Biochem. Biophys. Res. Commun. 1968, 31 (2), 145–150. 10.1016/0006-291X(68)90721-3. [DOI] [PubMed] [Google Scholar]
Yoshida Y.; Umeno A.; Akazawa Y.; Shichiri M.; Murotomi K.; Horie M. Chemistry of Lipid Peroxidation Products and Their Use as Biomarkers in Early Detection of Diseases. J. Oleo Sci. 2015, 64 (4), 347–356. 10.5650/jos.ess14281. [DOI] [PubMed] [Google Scholar]
Frankel E. N.; Neff W. E.; Rohwedder W. K.; Khambay B. P. S.; Garwood R. F.; Weedon B. C. L. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry: II. Methyl Linoleate. Lipids 1977, 12 (11), 908–913. 10.1007/BF02533310. [DOI] [PubMed] [Google Scholar]
Frankel E. N.; Neff W. E.; Rohwedder W. K.; Khambay B. P. S.; Garwood R. F.; Weedon B. C. L. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry: III. Methyl Linolenate. Lipids 1977, 12 (12), 1055–1061. 10.1007/BF02533334. [DOI] [PubMed] [Google Scholar]
Frankel E. N.; Neff W. E.; Rohwedder W. K.; Khambay B. P. S.; Garwood R. F.; Weedon B. C. L. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry: I. Methyl Oleate. Lipids 1977, 12 (11), 901–907. 10.1007/BF02533309. [DOI] [PubMed] [Google Scholar]
Watabe N.; Ishida Y.; Ochiai A.; Tokuoka Y.; Kawashima N. Oxidation Decomposition of Unsaturated Fatty Acids by Singlet Oxygen in Phospholipid Bilayer Membranes. J. Oleo Sci. 2007, 56 (2), 73–80. 10.5650/jos.56.73. [DOI] [PubMed] [Google Scholar]
Frimer A. A. The Reaction of Singlet Oxygen with Olefins: The Question of Mechanism. Chem. Rev. 1979, 79 (5), 359–387. 10.1021/cr60321a001. [DOI] [Google Scholar]
Minami Y.; Yokoyama K.; Bando N.; Kawai Y.; Terao J. Occurrence of Singlet Oxygen Oxygenation of Oleic Acid and Linoleic Acid in the Skin of Live Mice. Free Radic. Res. 2008, 42 (3), 197–204. 10.1080/10715760801948088. [DOI] [PubMed] [Google Scholar]
Frankel E. N.; Neff W. E.; Selke E. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry. IX. Homolytic vs. Heterolytic Cleavage of Primary and Secondary Oxidation Products. Lipids 1984, 19 (10), 790–800. 10.1007/BF02534473. [DOI] [Google Scholar]
Radi R.; Beckman J. S.; Bush K. M.; Freeman B. A. Peroxynitrite-Induced Membrane Lipid Peroxidation: The Cytotoxic Potential of Superoxide and Nitric Oxide. Arch. Biochem. Biophys. 1991, 288 (2), 481–487. 10.1016/0003-9861(91)90224-7. [DOI] [PubMed] [Google Scholar]
Rubbo H.; Parthasarathy S.; Barnes S.; Kirk M.; Kalyanaraman B.; Freeman B. A. Nitric Oxide Inhibition of Lipoxygenase-Dependent Liposome and Low-Density Lipoprotein Oxidation: Termination of Radical Chain Propagation Reactions and Formation of Nitrogen-Containing Oxidized Lipid Derivatives. Arch. Biochem. Biophys. 1995, 324 (1), 15–25. 10.1006/abbi.1995.9935. [DOI] [PubMed] [Google Scholar]
Freeman B. A.; Baker P. R. S.; Schopfer F. J.; Woodcock S. R.; Napolitano A.; d’Ischia M. Nitro-Fatty Acid Formation and Signaling. J. Biol. Chem. 2008, 283 (23), 15515–15519. 10.1074/jbc.R800004200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Cantu-Medellin N.; Kelley E. E. Xanthine Oxidoreductase-Catalyzed Reduction of Nitrite to Nitric Oxide: Insights Regarding Where, When and How. Nitric Oxide 2013, 34, 19–26. 10.1016/j.niox.2013.02.081. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kim-Shapiro D. B.; Schechter A. N.; Gladwin M. T. Unraveling the Reactions of Nitric Oxide, Nitrite, and Hemoglobin in Physiology and Therapeutics. Arterioscler. Thromb. Vasc. Biol. 2006, 26 (4), 697–705. 10.1161/01.ATV.0000204350.44226.9a. [DOI] [PubMed] [Google Scholar]
Spiteller P.; Spiteller G. 9-Hydroxy-10,12-Octadecadienoic Acid (9-HODE) and 13-Hydroxy-9,11-Octadecadienoic Acid (13-HODE): Excellent Markers for Lipid Peroxidation. Chem. Phys. Lipids 1997, 89 (2), 131–139. 10.1016/S0009-3084(97)00070-4. [DOI] [Google Scholar]
Bergström S.; Holman R. T. Total Conjugation of Linoleic Acid in Oxidation with Lipoxidase. Nature 1948, 161 (4080), 55. 10.1038/161055a0. [DOI] [PubMed] [Google Scholar]
Bergström S.; Holman R. T.. Lipoxidase and the Autoxidation of Unsaturated Fatty Acids. In Advances in Enzymology and Related Areas of Molecular Biology; John Wiley & Sons, 1948; pp 425–457. 10.1002/9780470122532.ch9 [DOI] [Google Scholar]
Russell G. A. Deuterium-Isotope Effects in the Autoxidation of Aralkyl Hydrocarbons. Mechanism of the Interaction of Peroxy Radicals. J. Am. Chem. Soc. 1957, 79 (14), 3871–3877. 10.1021/ja01571a068. [DOI] [Google Scholar]
Kanofsky J. R. Singlet Oxygen Production from the Reactions of Alkylperoxy Radicals. Evidence from 1268-nm Chemiluminescence. J. Org. Chem. 1986, 51 (17), 3386–3388. 10.1021/jo00367a032. [DOI] [Google Scholar]
Miyamoto S.; Martinez G. R.; Medeiros M. H. G.; Di Mascio P. Singlet Molecular Oxygen Generated from Lipid Hydroperoxides by the Russell Mechanism: Studies Using 18(O)-Labeled Linoleic Acid Hydroperoxide and Monomol Light Emission Measurements. J. Am. Chem. Soc. 2003, 125 (20), 6172–6179. 10.1021/ja029115o. [DOI] [PubMed] [Google Scholar]
Miyamoto S.; Martinez G. R.; Rettori D.; Augusto O.; Medeiros M. H. G.; Di Mascio P. Linoleic Acid Hydroperoxide Reacts with Hypochlorous Acid, Generating Peroxyl Radical Intermediates and Singlet Molecular Oxygen. Proc. Natl. Acad. Sci. U. S. A. 2006, 103 (2), 293–298. 10.1073/pnas.0508170103. [DOI] [PMC free article] [PubMed] [Google Scholar]
Takigawa Y.; Koshiishi I. Catalytic Production of Oxo-Fatty Acids by Lipoxygenases Is Mediated by the Radical-Radical Dismutation between Fatty Acid Alkoxyl Radicals and Fatty Acid Peroxyl Radicals in Fatty Acid Assembly. Chem. Pharm. Bull. (Tokyo) 2020, 68 (3), 258–264. 10.1248/cpb.c19-00975. [DOI] [PubMed] [Google Scholar]
Zhu X.; Tang X.; Anderson V. E.; Sayre L. M. Mass Spectrometric Characterization of Protein Modification by the Products of Nonenzymatic Oxidation of Linoleic Acid. Chem. Res. Toxicol. 2009, 22 (8), 1386–1397. 10.1021/tx9000072. [DOI] [PMC free article] [PubMed] [Google Scholar]
Labeque R.; Marnett L. J. Reaction of Hematin with Allylic Fatty Acid Hydroperoxides: Identification of Products and Implications for Pathways of Hydroperoxide-Dependent Epoxidation of 7,8-Dihydroxy-7,8-Dihydrobenzo[a]Pyrene. Biochemistry 1988, 27 (18), 7060–7070. 10.1021/bi00418a059. [DOI] [PubMed] [Google Scholar]
Dix T. A.; Marnett L. J. Conversion of Linoleic Acid Hydroperoxide to Hydroxy, Keto, Epoxyhydroxy, and Trihydroxy Fatty Acids by Hematin. J. Biol. Chem. 1985, 260 (9), 5351–5357. 10.1016/S0021-9258(18)89028-7. [DOI] [PubMed] [Google Scholar]
Neff W. E.; Frankel E. N.; Selke E.; Weisleder D. Photosensitized Oxidation of Methyl Linoleate Monohydroperoxides: Hydroperoxy Cyclic Peroxides, Dihydroperoxides, Keto Esters and Volatile Thermal Decomposition Products. Lipids 1983, 18 (12), 868–876. 10.1007/BF02534564. [DOI] [Google Scholar]
Hamberg M. A Novel Transformation of 13-Ls-Hydroperoxy-9,11-Octadecadienoic acid. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1983, 752 (2), 191–197. 10.1016/0005-2760(83)90112-1. [DOI] [PubMed] [Google Scholar]
Hamberg M. Autoxidation of Linoleic Acid: Isolation and Structure of Four Dihydroxyoctadecadienoic Acids. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1983, 752 (2), 353–356. 10.1016/0005-2760(83)90134-0. [DOI] [Google Scholar]
Niisuke K.; Boeglin W. E.; Murray J. J.; Schneider C.; Brash A. R. Biosynthesis of a Linoleic Acid Allylic Epoxide: Mechanistic Comparison with Its Chemical Synthesis and Leukotriene A Biosynthesis. J. Lipid Res. 2009, 50 (7), 1448–1455. 10.1194/jlr.M900025-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Frankel E. N.; Neff W. E.; Selke E.; Weisleder D. Photosensitized Oxidation of Methyl Linoleate: Secondary and Volatile Thermal Decomposition Products. Lipids 1982, 17 (1), 11–18. 10.1007/BF02535116. [DOI] [PubMed] [Google Scholar]
Neff W. E.; Frankel E. N.; Weisleder D. Photosensitized Oxidation of Methyl Linolenate. Secondary Products. Lipids 1982, 17 (11), 780–790. 10.1007/BF02535354. [DOI] [PubMed] [Google Scholar]
Hamberg M.; Gotthammar B. A New Reaction of Unsaturated Fatty Acid Hydroperoxides: Formation of 11-Hydroxy-12,13-Epoxy-9-Octadecenoic Acid from 13-Hydroperoxy-9,11-Octadecadienoic Acid. Lipids 1973, 8 (12), 737–744. 10.1007/BF02531842. [DOI] [Google Scholar]
Gardner H. W.; Kleiman R. Degradation of Linoleic Acid Hydroperoxides by a Cysteine FeCl3 Catalyst as a Model for Similar Biochemical Reactions: II. Specificity in Formation of Fatty Acid Epoxides. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1981, 665 (1), 113–125. 10.1016/0005-2760(81)90239-3. [DOI] [PubMed] [Google Scholar]
Gardner H. W.; Plattner R. D.; Weisleder D. The Epoxyallylic Radical from Homolysis and Rearrangement of Methyl Linoleate Hydroperoxide Combines with the Thiyl Radical of N-Acetylcysteine. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1985, 834 (1), 65–74. 10.1016/0005-2760(85)90177-8. [DOI] [Google Scholar]
Gardner H. W.; Weisleder D.; Kleiman R. Formation of trans-12,13-Epoxy-9-Hydroperoxy-trans-10-Octadecenoic Acid from 13-L-Hydroperoxy-cis-9, trans-11-Octadecadienoic Acid Catalyzed by Either a Soybean Extract or Cysteine-FeCl3. Lipids 1978, 13 (4), 246–252. 10.1007/BF02533664. [DOI] [PubMed] [Google Scholar]
Gardner H. W.; Selke E. Volatiles from Thermal Decomposition of Isomeric Methyl (12S, 13S)-(E)-12,13-Epoxy-9-Hydroperoxy-10-Octadecenoates. Lipids 1984, 19 (6), 375–380. 10.1007/BF02537398. [DOI] [Google Scholar]
Lin D.; Zhang J.; Sayre L. M. Synthesis of Six Epoxyketooctadecenoic Acid (EKODE) Isomers, Their Generation from Nonenzymatic Oxidation of Linoleic Acid, and Their Reactivity with Imidazole Nucleophiles. J. Org. Chem. 2007, 72 (25), 9471–9480. 10.1021/jo701373f. [DOI] [PubMed] [Google Scholar]
Szabó C.; Ischiropoulos H.; Radi R. Peroxynitrite: Biochemistry, Pathophysiology and Development of Therapeutics. Nat. Rev. Drug Discovery 2007, 6 (8), 662–680. 10.1038/nrd2222. [DOI] [PubMed] [Google Scholar]
Lundberg J. O.; Gladwin M. T.; Ahluwalia A.; Benjamin N.; Bryan N. S.; Butler A.; Cabrales P.; Fago A.; Feelisch M.; Ford P. C.; Freeman B. A.; Frenneaux M.; Friedman J.; Kelm M.; Kevil C. G.; Kim-Shapiro D. B.; Kozlov A. V.; Lancaster J. R.; Lefer D. J.; McColl K.; McCurry K.; Patel R. P.; Petersson J.; Rassaf T.; Reutov V. P.; Richter-Addo G. B.; Schechter A.; Shiva S.; Tsuchiya K.; van Faassen E. E.; Webb A. J.; Zuckerbraun B. S.; Zweier J. L.; Weitzberg E. Nitrate and Nitrite in Biology, Nutrition and Therapeutics. Nat. Chem. Biol. 2009, 5 (12), 865–869. 10.1038/nchembio.260. [DOI] [PMC free article] [PubMed] [Google Scholar]
Trujillo M.; Folkes L.; Bartesaghi S.; Kalyanaraman B.; Wardman P.; Radi R. Peroxynitrite-Derived Carbonate and Nitrogen Dioxide Radicals Readily React with Lipoic and Dihydrolipoic Acid. Free Radic. Biol. Med. 2005, 39 (2), 279–288. 10.1016/j.freeradbiomed.2005.03.014. [DOI] [PubMed] [Google Scholar]
Meli R.; Nauser T.; Latal P.; Koppenol W. H. Reaction of Peroxynitrite with Carbon Dioxide: Intermediates and Determination of the Yield of CO3•- and NO2•. J. Biol. Inorg. Chem. 2002, 7 (1), 31–36. 10.1007/s007750100262. [DOI] [PubMed] [Google Scholar]
Baker P. R. S.; Lin Y.; Schopfer F. J.; Woodcock S. R.; Groeger A. L.; Batthyany C.; Sweeney S.; Long M. H.; Iles K. E.; Baker L. M. S.; Branchaud B. P.; Chen Y. E.; Freeman B. A. Fatty Acid Transduction of Nitric Oxide Signaling: Multiple Nitrated Unsaturated Fatty Acid Derivatives Exist in Human Blood and Urine and Serve As Endogenous Peroxisome Proliferator-Activated Receptor Ligands. J. Biol. Chem. 2005, 280 (51), 42464–42475. 10.1074/jbc.M504212200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Baker P. R. S.; Schopfer F. J.; Sweeney S.; Freeman B. A. Red Cell Membrane and Plasma Linoleic Acid Nitration Products: Synthesis, Clinical Identification, and Quantitation. Proc. Natl. Acad. Sci. U. S. A. 2004, 101 (32), 11577–11582. 10.1073/pnas.0402587101. [DOI] [PMC free article] [PubMed] [Google Scholar]
Buchan G. J.; Bonacci G.; Fazzari M.; Salvatore S. R.; Gelhaus Wendell S. Nitro-Fatty Acid Formation and Metabolism. Nitric Oxide 2018, 79, 38–44. 10.1016/j.niox.2018.07.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
Napolitano A.; Camera E.; Picardo M.; d’Ischia M. Acid-Promoted Reactions of Ethyl Linoleate with Nitrite Ions: Formation and Structural Characterization of Isomeric Nitroalkene, Nitrohydroxy, and Novel 3-Nitro-1,5-Hexadiene and 1,5-Dinitro-1,3-Pentadiene Products. J. Org. Chem. 2000, 65 (16), 4853–4860. 10.1021/jo000090q. [DOI] [PubMed] [Google Scholar]
Onyango A. N. Endogenous Generation of Singlet Oxygen and Ozone in Human and Animal Tissues: Mechanisms, Biological Significance, and Influence of Dietary Components. Oxid. Med. Cell. Longev. 2016, 2016, 1–22. 10.1155/2016/2398573. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ahmed O. S.; Galano J.-M.; Pavlickova T.; Revol-Cavalier J.; Vigor C.; Lee J. C.-Y.; Oger C.; Durand T. Moving Forward with Isoprostanes, Neuroprostanes and Phytoprostanes: Where Are We Now?. Essays Biochem. 2020, 64 (3), 463–484. 10.1042/EBC20190096. [DOI] [PubMed] [Google Scholar]
Cuyamendous C.; de la Torre A.; Lee Y. Y.; Leung K. S.; Guy A.; Bultel-Poncé V.; Galano J.-M.; Lee J. C.-Y.; Oger C.; Durand T. The Novelty of Phytofurans, Isofurans, Dihomo-Isofurans and Neurofurans: Discovery, Synthesis and Potential Application. Biochimie 2016, 130, 49–62. 10.1016/j.biochi.2016.08.002. [DOI] [PubMed] [Google Scholar]
Samuelsson B. On the Incorporation of Oxygen in the Conversion of 8,11,14-Eicosatrienoic Acid to Prostaglandin E ₁. J. Am. Chem. Soc. 1965, 87 (13), 3011–3013. 10.1021/ja01091a043. [DOI] [PubMed] [Google Scholar]
Hamberg M.; Svensson J.; Wakabayashi T.; Samuelsson B. Isolation and Structure of Two Prostaglandin Endoperoxides That Cause Platelet Aggregation. Proc. Natl. Acad. Sci. U. S. A. 1974, 71 (2), 345–349. 10.1073/pnas.71.2.345. [DOI] [PMC free article] [PubMed] [Google Scholar]
Porter N. A.; Funk M. O. Peroxy Radical Cyclization as a Model for Prostaglandin Biosynthesis. J. Org. Chem. 1975, 40 (24), 3614–3615. 10.1021/jo00912a037. [DOI] [PubMed] [Google Scholar]
Porter N. A.; Funk M. O.; Gilmore D.; Isaac R.; Nixon J. The Formation of Cyclic Peroxides from Unsaturated Hydroperoxides: Models for Prostaglandin Biosynthesis. J. Am. Chem. Soc. 1976, 98, 6000–6005. 10.1021/ja00435a037. [DOI] [PubMed] [Google Scholar]
Pryor W. A.; Porter N. A. Suggested Mechanisms for the Production of 4-Hydroxy-2-Nonenal from the Autoxidation of Polyunsaturated Fatty Acids. Free Radic. Biol. Med. 1990, 8 (6), 541–543. 10.1016/0891-5849(90)90153-A. [DOI] [PubMed] [Google Scholar]
Roza M.; Francke A. Cyclic Peroxides from a Soya Lipoxygenase-Catalysed Oxygenation of Methyl Linolenate. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1978, 528 (1), 119–126. 10.1016/0005-2760(78)90057-7. [DOI] [PubMed] [Google Scholar]
Mihelich E. D. Structure and Stereochemistry of Novel Endoperoxides Isolated from the Sensitized Photooxidation of Methyl Linoleate. Implications for Prostaglandin Biosynthesis. J. Am. Chem. Soc. 1980, 102 (23), 7141–7143. 10.1021/ja00543a061. [DOI] [Google Scholar]
Chan H. W.-S.; Matthew J. A.; Coxon D. T. A Hydroperoxy-Epidioxide from the Autoxidation of a Hydroperoxide of Methyl Linolenate. J. Chem. Soc. Chem. Commun. 1980, (5), 235–236. 10.1039/c39800000235. [DOI] [Google Scholar]
Neff W. E.; Frankel E. N.; Weisleder D. High Pressure Liquid Chromatography of Autoxidized Lipids: II. Hydroperoxy-Cyclic Peroxides and Other Secondary Products from Methyl Linolenate. Lipids 1981, 16 (6), 439–448. 10.1007/BF02535012. [DOI] [Google Scholar]
Coxon D. T.; Price K. R.; Chan H. W.-S. Formation, Isolation and Structure Determination of Methyl Linolenate Diperoxides. Chem. Phys. Lipids 1981, 28 (4), 365–378. 10.1016/0009-3084(81)90022-0. [DOI] [Google Scholar]
Gunstone F. D.; Wijesundera R. C. Fatty Acids, Part 54. Some Reactions of Long-Chain Oxygenated Acids with Special Reference to Those Furnishing Furanoid Acids. Chem. Phys. Lipids 1979, 24 (2), 193–208. 10.1016/0009-3084(79)90088-4. [DOI] [Google Scholar]
Barden A. E.; Croft K. D.; Durand T.; Guy A.; Mueller M. J.; Mori T. A. Flaxseed Oil Supplementation Increases Plasma F₁-Phytoprostanes in Healthy Men. J. Nutr. 2009, 139 (10), 1890–1895. 10.3945/jn.109.108316. [DOI] [PubMed] [Google Scholar]
Dahle L. K.; Hill E. G.; Holman R. T. The Thiobarbituric Acid Reaction and the Autoxidations of Polyunsaturated Fatty Acid Methyl Esters. Arch. Biochem. Biophys. 1962, 98 (2), 253–261. 10.1016/0003-9861(62)90181-9. [DOI] [PubMed] [Google Scholar]
Porter N. A. Mechanisms for the Autoxidation of Polyunsaturated Lipids. Acc. Chem. Res. 1986, 19 (9), 262–268. 10.1021/ar00129a001. [DOI] [Google Scholar]
Corey E. J.; Shimoji K.; Shih C. Synthesis of Prostaglandins via a 1,2-Dioxabicyclo[2.2.1]Heptane (Endoperoxide) Intermediate. Stereochemical Divergence of Enzymic and Biomimetic Chemical Cyclization Reactions. J. Am. Chem. Soc. 1984, 106 (21), 6425–6427. 10.1021/ja00333a057. [DOI] [Google Scholar]
O’Connor D. E.; Mihelich E. D.; Coleman M. C. Isolation and Characterization of Bicycloendoperoxides Derived from Methyl Linolenate. J. Am. Chem. Soc. 1981, 103 (1), 223–224. 10.1021/ja00391a056. [DOI] [Google Scholar]
O’Connor D. E.; Mihelich E. D.; Coleman M. C. Stereochemical Course of the Autooxidative Cyclization of Lipid Hydroperoxides to Prostaglandin-like Bicyclic Endoperoxides. J. Am. Chem. Soc. 1984, 106 (12), 3577–3584. 10.1021/ja00324a028. [DOI] [Google Scholar]
Montine T. J.; Montine K. S.; Reich E. E.; Terry E. S.; Porter N. A.; Morrow J. D. Antioxidants Significantly Affect the Formation of Different Classes of Isoprostanes and Neuroprostanes in Rat Cerebral Synaptosomes. Biochem. Pharmacol. 2003, 65 (4), 611–617. 10.1016/S0006-2952(02)01607-6. [DOI] [PubMed] [Google Scholar]
Nugteren D. H.; Hazelhof E. Isolation and Properties of Intermediates in Prostaglandin Biosynthesis. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1973, 326 (3), 448–461. 10.1016/0005-2760(73)90145-8. [DOI] [PubMed] [Google Scholar]
Porter N. A.; Byers J. D.; Mebane R. C.; Gilmore D. W.; Nixon J. R. Prostaglandin H₂ Methyl Ester. J. Org. Chem. 1978, 43 (10), 2088–2090. 10.1021/jo00404a067. [DOI] [Google Scholar]
Chen Y.; Morrow J. D.; Roberts L. J. Formation of Reactive Cyclopentenone Compounds in Vivo as Products of the Isoprostane Pathway. J. Biol. Chem. 1999, 274 (16), 10863–10868. 10.1074/jbc.274.16.10863. [DOI] [PubMed] [Google Scholar]
Thoma I.; Loeffler C.; Sinha A. K.; Gupta M.; Krischke M.; Steffan B.; Roitsch T.; Mueller M. J. Cyclopentenone Isoprostanes Induced by Reactive Oxygen Species Trigger Defense Gene Activation and Phytoalexin Accumulation in Plants. Plant J. 2003, 34 (3), 363–375. 10.1046/j.1365-313X.2003.01730.x. [DOI] [PubMed] [Google Scholar]
Fessel J. P.; Porter N. A.; Moore K. P.; Sheller J. R.; Roberts L. J. Discovery of Lipid Peroxidation Products Formed in Vivo with a Substituted Tetrahydrofuran Ring (Isofurans) That Are Favored by Increased Oxygen Tension. Proc. Natl. Acad. Sci. U. S. A. 2002, 99 (26), 16713–16718. 10.1073/pnas.252649099. [DOI] [PMC free article] [PubMed] [Google Scholar]
Roberts L. J.; Fessel J. P. The Biochemistry of the Isoprostane, Neuroprostane, and Isofuran Pathways of Lipid Peroxidation. Chem. Phys. Lipids 2004, 128 (1), 173–186. 10.1016/j.chemphyslip.2003.09.016. [DOI] [PubMed] [Google Scholar]
Shearer G. C.; Newman J. W. Lipoprotein Lipase Releases Esterified Oxylipins from Very Low-Density Lipoproteins. Prostaglandins Leukot. Essent. Fatty Acids 2008, 79 (6), 215–222. 10.1016/j.plefa.2008.09.023. [DOI] [PMC free article] [PubMed] [Google Scholar]
Liang N.; Harsch B. A.; Zhou S.; Borkowska A.; Shearer G. C.; Kaddurah-Daouk R.; Newman J. W.; Borkowski K. Oxylipin Transport by Lipoprotein Particles and Its Functional Implications for Cardiometabolic and Neurological Disorders. Prog. Lipid Res. 2024, 93, 101265. 10.1016/j.plipres.2023.101265. [DOI] [PubMed] [Google Scholar]
O’Donnell V. B.; Murphy R. C. New Families of Bioactive Oxidized Phospholipids Generated by Immune Cells: Identification and Signaling Actions. Blood 2012, 120 (10), 1985–1992. 10.1182/blood-2012-04-402826. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fazzari M.; Vitturi D. A.; Woodcock S. R.; Salvatore S. R.; Freeman B. A.; Schopfer F. J. Electrophilic Fatty Acid Nitroalkenes Are Systemically Transported and Distributed upon Esterification to Complex Lipids. J. Lipid Res. 2019, 60 (2), 388–399. 10.1194/jlr.M088815. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fazzari M.; Khoo N. H.; Woodcock S. R.; Jorkasky D. K.; Li L.; Schopfer F. J.; Freeman B. A. Nitro-Fatty Acid Pharmacokinetics in the Adipose Tissue Compartment. J. Lipid Res. 2017, 58 (2), 375–385. 10.1194/jlr.M072058. [DOI] [PMC free article] [PubMed] [Google Scholar]
Annevelink C. E.; Walker R. E.; Shearer G. C. Esterified Oxylipins: Do They Matter?. Metabolites 2022, 12 (11), 1007. 10.3390/metabo12111007. [DOI] [PMC free article] [PubMed] [Google Scholar]
Carpanedo L.; Rund K. M.; Wende L. M.; Kampschulte N.; Schebb N. H. LC-HRMS Analysis of Phospholipids Bearing Oxylipins. Analytica Chimica Acta 2024, 1326, 343139. 10.1016/j.aca.2024.343139. [DOI] [PubMed] [Google Scholar]
Genva M.; Obounou Akong F.; Andersson M. X.; Deleu M.; Lins L.; Fauconnier M.-L. New Insights into the Biosynthesis of Esterified Oxylipins and Their Involvement in Plant Defense and Developmental Mechanisms. Phytochem. Rev. 2019, 18 (1), 343–358. 10.1007/s11101-018-9595-8. [DOI] [Google Scholar]
Astudillo A. M.; Balgoma D.; Balboa M. A.; Balsinde J. Dynamics of Arachidonic Acid Mobilization by Inflammatory Cells. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2012, 1821 (2), 249–256. 10.1016/j.bbalip.2011.11.006. [DOI] [PubMed] [Google Scholar]
Samuelsson B. Role of Basic Science in the Development of New Medicines: Examples from the Eicosanoid Field. J. Biol. Chem. 2012, 287 (13), 10070–10080. 10.1074/jbc.X112.351437. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kobe M. J.; Neau D. B.; Mitchell C. E.; Bartlett S. G.; Newcomer M. E. The Structure of Human 15-Lipoxygenase-2 with a Substrate Mimic. J. Biol. Chem. 2014, 289 (12), 8562–8569. 10.1074/jbc.M113.543777. [DOI] [PMC free article] [PubMed] [Google Scholar]
Belkner J.; Stender H.; Kühn H. The Rabbit 15-Lipoxygenase Preferentially Oxygenates LDL Cholesterol Esters, and This Reaction Does Not Require Vitamin E. J. Biol. Chem. 1998, 273 (36), 23225–23232. 10.1074/jbc.273.36.23225. [DOI] [PubMed] [Google Scholar]
O’Donnell V. B. New Appreciation for an Old Pathway: The Lands Cycle Moves into New Arenas in Health and Disease. Biochem. Soc. Trans. 2022, 50 (1), 1–11. 10.1042/BST20210579. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yuan H.; Li X.; Zhang X.; Kang R.; Tang D. Identification of ACSL4 as a Biomarker and Contributor of Ferroptosis. Biochem. Biophys. Res. Commun. 2016, 478 (3), 1338–1343. 10.1016/j.bbrc.2016.08.124. [DOI] [PubMed] [Google Scholar]
Klett E. L.; Chen S.; Yechoor A.; Lih F. B.; Coleman R. A. Long-Chain Acyl-CoA Synthetase Isoforms Differ in Preferences for Eicosanoid Species and Long-Chain Fatty Acids. J. Lipid Res. 2017, 58 (5), 884–894. 10.1194/jlr.M072512. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kanter J. E.; Tang C.; Oram J. F.; Bornfeldt K. E. Acyl-CoA Synthetase 1 Is Required for Oleate and Linoleate Mediated Inhibition of Cholesterol Efflux through ATP-Binding Cassette Transporter A1 in Macrophages. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2012, 1821 (3), 358–364. 10.1016/j.bbalip.2011.10.008. [DOI] [PMC free article] [PubMed] [Google Scholar]
Golej D. L.; Askari B.; Kramer F.; Barnhart S.; Vivekanandan-Giri A.; Pennathur S.; Bornfeldt K. E. Long-Chain Acyl-CoA Synthetase 4 Modulates Prostaglandin E₂ Release from Human Arterial Smooth Muscle Cells. J. Lipid Res. 2011, 52 (4), 782–793. 10.1194/jlr.M013292. [DOI] [PMC free article] [PubMed] [Google Scholar]
Walker R. E.; Richter C. K.; Skulas-Ray A. C.; Flock M. R.; Harsch B. A.; Annevelink C. E.; Kris-Etherton P. M.; Jensen G. L.; Shearer G. C. Effect of Omega-3 Ethyl Esters on the Triglyceride-Rich Lipoprotein Response to Endotoxin Challenge in Healthy Young Men. J. Lipid Res. 2023, 64 (5), 100353. 10.1016/j.jlr.2023.100353. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wopereis S.; Wolvers D.; van Erk M.; Gribnau M.; Kremer B.; van Dorsten F. A.; Boelsma E.; Garczarek U.; Cnubben N.; Frenken L.; van der Logt P.; Hendriks H. F. J.; Albers R.; van Duynhoven J.; van Ommen B.; Jacobs D. M. Assessment of Inflammatory Resilience in Healthy Subjects Using Dietary Lipid and Glucose Challenges. BMC Med. Genomics 2013, 6, 44. 10.1186/1755-8794-6-44. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wright D. N.; Katundu K. G. H.; Viscarra J. A.; Crocker D. E.; Newman J. W.; La Frano M. R.; Ortiz R. M. Oxylipin Responses to Fasting and Insulin Infusion in a Large Mammalian Model of Fasting-Induced Insulin Resistance, the Northern Elephant Seal. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 2021, 321 (4), R537–R546. 10.1152/ajpregu.00016.2021. [DOI] [PMC free article] [PubMed] [Google Scholar]
Shearer G. C.; Borkowski K.; Puumala S. L.; Harris W. S.; Pedersen T. L.; Newman J. W. Abnormal Lipoprotein Oxylipins in Metabolic Syndrome and Partial Correction by Omega-3 Fatty Acids. Prostaglandins Leukot. Essent. Fatty Acids 2018, 128, 1–10. 10.1016/j.plefa.2017.10.006. [DOI] [PubMed] [Google Scholar]
Schebb N. H.; Ostermann A. I.; Yang J.; Hammock B. D.; Hahn A.; Schuchardt J. P. Comparison of the Effects of Long-Chain Omega-3 Fatty Acid Supplementation on Plasma Levels of Free and Esterified Oxylipins. Prostaglandins Other Lipid Mediat. 2014, 113–115, 21–29. 10.1016/j.prostaglandins.2014.05.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Slatter D. A.; Aldrovandi M.; O’Connor A.; Allen S. M.; Brasher C. J.; Murphy R. C.; Mecklemann S.; Ravi S.; Darley-Usmar V.; O’Donnell V. B. Mapping the Human Platelet Lipidome Reveals Cytosolic Phospholipase A2 as a Regulator of Mitochondrial Bioenergetics during Activation. Cell Metab. 2016, 23 (5), 930–944. 10.1016/j.cmet.2016.04.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kolmert J.; Fauland A.; Fuchs D.; Säfholm J.; Gómez C.; Adner M.; Dahlén S.-E.; Wheelock C. E. Lipid Mediator Quantification in Isolated Human and Guinea Pig Airways: An Expanded Approach for Respiratory Research. Anal. Chem. 2018, 90 (17), 10239–10248. 10.1021/acs.analchem.8b01651. [DOI] [PubMed] [Google Scholar]
Pedersen T. L.; Gray I. J.; Newman J. W. Plasma and Serum Oxylipin, Endocannabinoid, Bile Acid, Steroid, Fatty Acid and Nonsteroidal Anti-Inflammatory Drug Quantification in a 96-Well Plate Format. Anal. Chim. Acta 2021, 1143, 189–200. 10.1016/j.aca.2020.11.019. [DOI] [PubMed] [Google Scholar]
Rajan M. R.; Sotak M.; Barrenäs F.; Shen T.; Borkowski K.; Ashton N. J.; Biörserud C.; Lindahl T. L.; Ramström S.; Schöll M.; Lindahl P.; Fiehn O.; Newman J. W.; Perkins R.; Wallenius V.; Lange S.; Börgeson E. Comparative Analysis of Obesity-Related Cardiometabolic and Renal Biomarkers in Human Plasma and Serum. Sci. Rep. 2019, 9 (1), 15385. 10.1038/s41598-019-51673-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
Matthew J. A.; Chan H. W. S.; Galliard T. A Simple Method for the Preparation of Pure 9-D-Hydroperoxide of Linoleic Acid and Methyl Linoleate Based on the Positional Specificity of Lipoxygenase in Tomato Fruit. Lipids 1977, 12 (3), 324–326. 10.1007/BF02533358. [DOI] [PubMed] [Google Scholar]
Gardner H. W.Analysis of Plant Lipoxygenase Metabolites in Advances in Lipid Methodology - Four; W.W. Christie, Oily Press, Dundee, 1997. [Google Scholar]
Iacazio G.; Langrand G.; Baratti J.; Buono G.; Triantaphylides C. Preparative, Enzymic Synthesis of Linoleic Acid (13S)-Hydroperoxide Using Soybean Lipoxygenase-1. J. Org. Chem. 1990, 55 (5), 1690–1691. 10.1021/jo00292a056. [DOI] [Google Scholar]
Corey E. J.; Lansbury P. T. Jr Stereochemical Course of 5-Lipoxygenation of Arachidonate by Rat Basophil Leukemic Cell (RBL-1) and Potato Enzymes. J. Am. Chem. Soc. 1983, 105 (12), 4093–4094. 10.1021/ja00350a059. [DOI] [Google Scholar]
Boeglin W. E.; Kim R. B.; Brash A. R. A 12R-Lipoxygenase in Human Skin: Mechanistic Evidence, Molecular Cloning, and Expression. Proc. Natl. Acad. Sci. U. S. A. 1998, 95 (12), 6744–6749. 10.1073/pnas.95.12.6744. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hamberg M. Fatty Acid Hydroperoxide Isomerase in Saprolegnia parasitica: Structural Studies of Epoxy Alcohols Formed from Isomeric Hydroperoxyoctadecadienoates. Lipids 1989, 24 (4), 249–255. 10.1007/BF02535158. [DOI] [Google Scholar]
Peers K. E.; Coxon D. T. Controlled Synthesis of Monohydroperoxides by α-Tocopherol Inhibited Autoxidation of Polyunsaturated Lipids. Chem. Phys. Lipids 1983, 32 (1), 49–56. 10.1016/0009-3084(83)90069-5. [DOI] [Google Scholar]
Hamberg M. Stereochemistry of Hydrogen Removal During Oxygenation of Linoleic Acid by Singlet Oxygen and Synthesis of 11(S)-Deuterium-Labeled Linoleic Acid. Lipids 2011, 46 (2), 201–206. 10.1007/s11745-010-3510-4. [DOI] [PubMed] [Google Scholar]
Przybyla D.; Göbel C.; Imboden A.; Hamberg M.; Feussner I.; Apel K. Enzymatic, but Not Non-Enzymatic, ¹O₂-Mediated Peroxidation of Polyunsaturated Fatty Acids Forms Part of the EXECUTER1-Dependent Stress Response Program in the Flu Mutant of Arabidopsis thaliana. Plant J. 2008, 54 (2), 236–248. 10.1111/j.1365-313X.2008.03409.x. [DOI] [PubMed] [Google Scholar]
Hamberg M. Steric Analysis of Hydroperoxides Formed by Lipoxygenase Oxygenation of Linoleic Acid. Anal. Biochem. 1971, 43 (2), 515–526. 10.1016/0003-2697(71)90282-X. [DOI] [PubMed] [Google Scholar]
Kai K.; Takeuchi J.; Kataoka T.; Yokoyama M.; Watanabe N. Structure-Activity Relationship Study of Flowering-Inducer FN against Lemna paucicostata. Tetrahedron 2008, 64 (28), 6760–6769. 10.1016/j.tet.2008.04.115. [DOI] [Google Scholar]
Bergström S.; Aulin-Erdtman G.; Rolander B.; Stenhagen E.; Östling S. The Monoketo- and Monohydroxyoctadecanoic Acids. Preparation and Characterization by Thermal and X-Ray Methods. Acta Chem. Scand. 1952, 6, 1157–1174. 10.3891/acta.chem.scand.06-1157. [DOI] [Google Scholar]
Iacazio G. Easy Access to Various Natural Keto Polyunsaturated Fatty Acids and Their Corresponding Racemic Alcohols. Chem. Phys. Lipids 2003, 125 (2), 115–121. 10.1016/S0009-3084(03)00087-2. [DOI] [PubMed] [Google Scholar]
Armstrong M. M.; Diaz G.; Kenyon V.; Holman T. R. Inhibitory and Mechanistic Investigations of Oxo-Lipids with Human Lipoxygenase Isozymes. Bioorg. Med. Chem. 2014, 22 (15), 4293–4297. 10.1016/j.bmc.2014.05.025. [DOI] [PMC free article] [PubMed] [Google Scholar]
Maerker G.; Haeberer E. T.; Ault W. C. Epoxidation of Methyl Linoleate. I. The Question of Positional Selectivity in Monoepoxidation. J. Am. Oil Chem. Soc. 1966, 43 (2), 100–104. 10.1007/BF02641026. [DOI] [PubMed] [Google Scholar]
Ferrari M.; Ghisalberti E. L.; Pagnoni U. M.; Pelizzoni F. Monoepoxidation of Methyl Linoleate: (±)-Methyl Vernolate and (±)-Methyl Coronarate. J. Am. Oil Chem. Soc. 1968, 45 (10), 649–651. 10.1007/BF02541248. [DOI] [Google Scholar]
Newman J. W.; Watanabe T.; Hammock B. D. The Simultaneous Quantification of Cytochrome P450 Dependent Linoleate and Arachidonate Metabolites in Urine by HPLC-MS/MS. J. Lipid Res. 2002, 43 (9), 1563–1578. 10.1194/jlr.D200018-JLR200. [DOI] [PubMed] [Google Scholar]
Teder T.; Boeglin W. E.; Brash A. R. Oxidation of C18 Hydroxy-Polyunsaturated Fatty Acids to Epoxide or Ketone by Catalase-Related Hemoproteins Activated with Iodosylbenzene. Lipids 2017, 52 (7), 587–597. 10.1007/s11745-017-4271-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
Municoy M.; González-Benjumea A.; Carro J.; Aranda C.; Linde D.; Renau-Mínguez C.; Ullrich R.; Hofrichter M.; Guallar V.; Gutiérrez A.; Martínez A. T. Fatty-Acid Oxygenation by Fungal Peroxygenases: From Computational Simulations to Preparative Regio- and Stereoselective Epoxidation. ACS Catal. 2020, 10 (22), 13584–13595. 10.1021/acscatal.0c03165. [DOI] [Google Scholar]
Gao B.; Boeglin W. E.; Zheng Y.; Schneider C.; Brash A. R. Evidence for an Ionic Intermediate in the Transformation of Fatty Acid Hydroperoxide by a Catalase-Related Allene Oxide Synthase from the Cyanobacterium Acaryochloris marina. J. Biol. Chem. 2009, 284 (33), 22087–22098. 10.1074/jbc.M109.013151. [DOI] [PMC free article] [PubMed] [Google Scholar]
SMITH C. R. Jr; BAGBY M. O.; LOHMAR R. L.; GLASS C. A.; WOLFF I. A. The Epoxy Acids of Chrysanthemum coronarium and Clarkia elegans Seed Oils. J. Org. Chem. 1960, 25 (2), 218–222. 10.1021/jo01072a019. [DOI] [Google Scholar]
Gunstone F. D. Fatty Acids. Part II. The Nature of the Oxygenated Acid Present in Vernonia anthelmintica(Willd.) Seed Oil. J. Chem. Soc. Resumed 1954, (0), 1611–1616. 10.1039/jr9540001611. [DOI] [Google Scholar]
Osbond J. M. 1043. Essential Fatty Acids. Part II. Synthesis of (±)-Vernolic, Linoleic, and γ-Linolenic Acid. J. Chem. Soc. Resumed 1961, 0 (0), 5270–5275. 10.1039/JR9610005270. [DOI] [Google Scholar]
Morris L. J.; Wharry D. M. Naturally Occurring Epoxy Acids. IV. The Absolute Optical Configuration of Vernolic Acid. Lipids 1966, 1 (1), 41–46. 10.1007/BF02668123. [DOI] [PubMed] [Google Scholar]
VanRheenen V.; Cha D. Y.; Hartley W. M.. Catalytic Osmium Tetroxide Oxidation of Olefins: cis-1,2-Cyclohexanediol. In Organic Syntheses; John Wiley & Sons, 2003; pp 43–43. 10.1002/0471264180.os058.08. [DOI] [Google Scholar]
Corey E. J.; Su W.; Mehrotra M. M. An Efficient and Simple Method for the Conversion of 15-HPETE to 14,15-EPETE (Lipotriene A) and 5-HPETE to Leukotriene a as the Methyl Esters. Tetrahedron Lett. 1984, 25 (45), 5123–5126. 10.1016/S0040-4039(01)81540-9. [DOI] [Google Scholar]
Hamberg M. Vanadium-Catalyzed Transformation of 13(S)-Hydroperoxy-9(Z), 11(E)-Octadecadienoic Acid: Structural Studies on Epoxy Alcohols and Trihydroxy Acids. Chem. Phys. Lipids 1987, 43 (1), 55–67. 10.1016/0009-3084(87)90017-X. [DOI] [Google Scholar]
Hamberg M.; Herman C. A.; Herman R. P. Novel Biological Transformations of 15-Ls-Hydroperoxy-5,8,11,13-Eicosatetraenoic Acid. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1986, 877 (3), 447–457. 10.1016/0005-2760(86)90211-0. [DOI] [PubMed] [Google Scholar]
Hamberg M.; Olsson U. Efficient and Specific Conversion of 9-Lipoxygenase Hydroperoxides in the Beetroot. Formation of Pinellic Acid. Lipids 2011, 46 (9), 873–878. 10.1007/s11745-011-3592-7. [DOI] [PubMed] [Google Scholar]
Thomas C. P.; Boeglin W. E.; Garcia-Diaz Y.; O’Donnell V. B.; Brash A. R. Steric Analysis of Epoxyalcohol and Trihydroxy Derivatives of 9-Hydroperoxy-Linoleic Acid from Hematin and Enzymatic Synthesis. Chem. Phys. Lipids 2013, 167–168, 21–32. 10.1016/j.chemphyslip.2013.01.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hamberg M. Regio- and Stereochemical Analysis of Trihydroxyoctadecenoic Acids Derived from Linoleic Acid 9- and 13-Hydroperoxides. Lipids 1991, 26 (6), 407–415. 10.1007/BF02536065. [DOI] [PubMed] [Google Scholar]
Fuchs D.; Hamberg M.; Sköld C. M.; Wheelock Å. M.; Wheelock C. E. An LC-MS/MS Workflow to Characterize 16 Regio- and Stereoisomeric Trihydroxyoctadecenoic Acids. J. Lipid Res. 2018, 59 (10), 2025–2033. 10.1194/jlr.D087429. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hamberg M.; Hamberg G. Peroxygenase-Catalyzed Fatty Acid Epoxidation in Cereal Seeds (Sequential Oxidation of Linoleic Acid into 9(S),12(S),13(S)-Trihydroxy-10(E)-Octadecenoic Acid). Plant Physiol. 1996, 110 (3), 807–815. 10.1104/pp.110.3.807. [DOI] [PMC free article] [PubMed] [Google Scholar]
Davis R. W.; Allweil A.; Tian J.; Brash A. R.; Sulikowski G. A. Stereocontrolled Synthesis of Four Isomeric Linoleate Triols of Relevance to Skin Barrier Formation and Function. Tetrahedron Lett. 2018, 59 (52), 4571–4573. 10.1016/j.tetlet.2018.11.033. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sabitha G.; Bhikshapathi M.; Reddy E. V.; Yadav J. S. Synthesis of (−)-Pinellic Acid and Its (9R,12S,13S)-Diastereoisomer. Helv. Chim. Acta 2009, 92 (10), 2052–2057. 10.1002/hlca.200900095. [DOI] [Google Scholar]
Woodcock S. R.; Bonacci G.; Gelhaus S. L.; Schopfer F. J. Nitrated Fatty Acids: Synthesis and Measurement. Free Radic. Biol. Med. 2013, 59, 14–26. 10.1016/j.freeradbiomed.2012.11.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kaya K.; Miura T. Effects of Nitrogen Dioxide on Fatty Acid Compositions of Red Cell Membranes, Sera, and Livers in Rats. Environ. Res. 1982, 27 (1), 24–35. 10.1016/0013-9351(82)90054-8. [DOI] [PubMed] [Google Scholar]
Kunimoto M.; Mochitate K.; Kaya K.; Miura T.; Kubota K. Effects of Nitrogen Dioxide on Red Blood Cells of Rats: Alterations of Cell Membrane Components and Populational Changes of Red Blood Cells during in Vivo Exposure to NO2. Environ. Res. 1984, 33 (2), 361–369. 10.1016/0013-9351(84)90034-3. [DOI] [PubMed] [Google Scholar]
Kaya K.; Miura T.; Kubota K. Effects of Nitrogen Dioxide on Red Blood Cells of Rats: Changes in Components of Red Cell Membranes during in Vivo Exposure to NO2. Environ. Res. 1980, 23 (2), 397–409. 10.1016/0013-9351(80)90074-2. [DOI] [PubMed] [Google Scholar]
O’Donnell V. B.; Eiserich J. P.; Chumley P. H.; Jablonsky M. J.; Krishna N. R.; Kirk M.; Barnes S.; Darley-Usmar V. M.; Freeman B. A. Nitration of Unsaturated Fatty Acids by Nitric Oxide-Derived Reactive Nitrogen Species Peroxynitrite, Nitrous Acid, Nitrogen Dioxide, and Nitronium Ion. Chem. Res. Toxicol. 1999, 12 (1), 83–92. 10.1021/tx980207u. [DOI] [PubMed] [Google Scholar]
Gorczynski M. J.; Huang J.; King S. B. Regio- and Stereospecific Syntheses and Nitric Oxide Donor Properties of (E)-9- and (E)-10-Nitrooctadec-9-Enoic Acids. Org. Lett. 2006, 8 (11), 2305–2308. 10.1021/ol060548w. [DOI] [PMC free article] [PubMed] [Google Scholar]
Woodcock S. R.; Marwitz A. J. V.; Bruno P.; Branchaud B. P. Synthesis of Nitrolipids. All Four Possible Diastereomers of Nitrooleic Acids: (E)- and (Z)-, 9- and 10-Nitro-Octadec-9-Enoic Acids. Org. Lett. 2006, 8 (18), 3931–3934. 10.1021/ol0613463. [DOI] [PubMed] [Google Scholar]
Dunny E.; Evans P. Stereocontrolled Synthesis of the PPAR-γ Agonist 10-Nitrolinoleic Acid. J. Org. Chem. 2010, 75 (15), 5334–5336. 10.1021/jo1007493. [DOI] [PubMed] [Google Scholar]
Zanoni G.; Valli M.; Bendjeddou L.; Porta A.; Bruno P.; Vidari G. Improved Synthesis of (E)-12-Nitrooctadec-12-Enoic Acid, a Potent PPARγ Activator. Development of a “Buffer-Free” Enzymatic Method for Hydrolysis of Methyl Esters. J. Org. Chem. 2010, 75 (23), 8311–8314. 10.1021/jo101806m. [DOI] [PubMed] [Google Scholar]
Hassan M.; Krieg S.-C.; Ndefo Nde C.; Roos J.; Maier T. J.; El Rady E. A.; Raslan M. A.; Sadek K. U.; Manolikakes G. Streamlined One-Pot Synthesis of Nitro Fatty Acids. Eur. J. Org. Chem. 2021, 2021 (15), 2239–2252. 10.1002/ejoc.202100247. [DOI] [Google Scholar]
Elix J. A.; Sargent M. V. Studies in Furan Chemistry. Part VI. The Synthesis of 8-(5-Hexyl-2-Furyl)Octanoic Acid, a Fatty Acid Found in Exocarpus Seed Oil. J. Chem. Soc. C Org. 1968, (0), 595–596. 10.1039/j39680000595. [DOI] [Google Scholar]
Lie Ken Jie M. S. F.; Wong K. P. Synthesis of Dimethyl-Substituted C₁₈ Furanoid Fatty Ester. J. Am. Oil Chem. Soc. 1992, 69 (5), 485–487. 10.1007/BF02540955. [DOI] [Google Scholar]
Lie Ken Jie M. S. F.; Lam C. H. Fatty Acids, Part XVI: The Synthesis of All Isomeric C18 Furan-Containing Fatty Acids. Chem. Phys. Lipids 1978, 21 (3), 275–287. 10.1016/0009-3084(78)90073-7. [DOI] [Google Scholar]
Alaiz M.; Hidalgo F. J.; Zamora R.; Millán F.; Maza M. P.; Vioque E. Synthesis of 9,12-Epoxyoctadeca-9,11-Dienoic Acid. Chem. Phys. Lipids 1988, 48 (3), 289–292. 10.1016/0009-3084(88)90099-0. [DOI] [Google Scholar]
El Fangour S.; Guy A.; Vidal J.-P.; Rossi J.-C.; Durand T. Total Synthesis of Phytoprostane F₁ and Its 16 Epimer. Tetrahedron Lett. 2003, 44 (10), 2105–2108. 10.1016/S0040-4039(03)00171-0. [DOI] [Google Scholar]
El Fangour S.; Guy A.; Despres V.; Vidal J.-P.; Rossi J.-C.; Durand T. Total Synthesis of the Eight Diastereomers of the Syn-Anti-Syn Phytoprostanes F₁ Types I and II. J. Org. Chem. 2004, 69 (7), 2498–2503. 10.1021/jo035638i. [DOI] [PubMed] [Google Scholar]
Roland A.; Durand T.; Egron D.; Vidal J.-P.; Rossi J.-C. Radical Cyclization of Highly Functionalized Precursors: Stereocontrol of Ring Closure of Acyclic 1-Substituted-2,4-Dihydroxylated Hex-5-Enyl Radicals. J. Chem. Soc. Perkin 1 2000, (2), 245–251. 10.1039/a904206g. [DOI] [Google Scholar]
Durand T.; Guy A.; Henry O.; Vidal J.-P.; Rossi J.-C.; Rivalta C.; Valagussa A.; Chiabrando C. Total Syntheses of Four Metabolites of 15-F_2t-Isoprostane. Eur. J. Org. Chem. 2001, 2001 (4), 809–819. . [DOI] [Google Scholar]
Smrček J.; Hájek M.; Hodek O.; Čížek K.; Pohl R.; Jahn E.; Galano J.; Oger C.; Durand T.; Cvačka J.; Jahn U. First Total Synthesis of Phytoprostanes with Prostaglandin-Like Configuration, Evidence for Their Formation in Edible Vegetable Oils and Orienting Study of Their Biological Activity. Chem. - Eur. J. 2021, 27 (37), 9556–9562. 10.1002/chem.202100872. [DOI] [PubMed] [Google Scholar]
Rodrıguez A. R.; Spur B. W. First Total Synthesis of the E Type I Phytoprostanes. Tetrahedron Lett. 2003, 44 (40), 7411–7415. 10.1016/j.tetlet.2003.08.042. [DOI] [Google Scholar]
Brinkmann Y.; Oger C.; Guy A.; Durand T.; Galano J.-M. Total Synthesis of 15-D2t- and 15-Epi-15-E2t-Isoprostanes. J. Org. Chem. 2010, 75 (7), 2411–2414. 10.1021/jo1000274. [DOI] [PubMed] [Google Scholar]
Oger C.; Brinkmann Y.; Bouazzaoui S.; Durand T.; Galano J.-M. Stereocontrolled Access to Isoprostanes via a Bicyclo[3.3.0]Octene Framework. Org. Lett. 2008, 10 (21), 5087–5090. 10.1021/ol802104z. [DOI] [PubMed] [Google Scholar]
Oger C.; Marton Z.; Brinkmann Y.; Bultel-Poncé V.; Durand T.; Graber M.; Galano J.-M. Lipase-Catalyzed Regioselective Monoacetylation of Unsymmetrical 1,5-Primary Diols. J. Org. Chem. 2010, 75 (6), 1892–1897. 10.1021/jo902541c. [DOI] [PubMed] [Google Scholar]
Porta A.; Chiesa F.; Quaroni M.; Persico M.; Moratti R.; Zanoni G.; Vidari G. A Divergent Enantioselective Synthesis of 9-J1-Phytoprostane and 9-A1-Phytoprostane Methyl Ester. Eur. J. Org. Chem. 2014, 2014 (10), 2111–2119. 10.1002/ejoc.201301703. [DOI] [Google Scholar]
Zanoni G.; Porta A.; Meriggi A.; Franzini M.; Vidari G. 1,2-Oxopalladation versus π-Allyl Palladium Route. A Regioconvergent Approach to a Key Intermediate for Cyclopentanoids Synthesis. New Insights into the Pd(II)-Catalyzed Lactonization Reaction. J. Org. Chem. 2002, 67 (17), 6064–6069. 10.1021/jo025722i. [DOI] [PubMed] [Google Scholar]
Zanoni G.; Agnelli F.; Meriggi A.; Vidari G. Enantioselective Syntheses of Isoprostane and Iridoid Lactones Intermediates by Enzymatic Transesterification. Tetrahedron Asymmetry 2001, 12 (12), 1779–1784. 10.1016/S0957-4166(01)00311-1. [DOI] [Google Scholar]
Schmidt A.; Boland W. General Strategy for the Synthesis of B1 Phytoprostanes, Dinor Isoprostanes, and Analogs. J. Org. Chem. 2007, 72 (5), 1699–1706. 10.1021/jo062359x. [DOI] [PubMed] [Google Scholar]
Perlikowska W.; Mikołajczyk M. A Short Synthesis of Enantiomeric Phytoprostanes B₁ Type I. Synthesis 2009, 2009 (16), 2715–2718. 10.1055/s-0029-1216883. [DOI] [Google Scholar]
Perlikowska W.; Mikołajczyk M. A Concise Approach to Both Enantiomers of Phytoprostane B₁ Type II. Tetrahedron Asymmetry 2011, 22 (18), 1767–1771. 10.1016/j.tetasy.2011.10.005. [DOI] [Google Scholar]
Vázquez-Romero A.; Verdaguer X.; Riera A. General Approach to Prostanes B₁ by Intermolecular Pauson-Khand Reaction: Syntheses of Methyl Esters of Prostaglandin B₁ and Phytoprostanes 16-B₁-PhytoP and 9-L₁-PhytoP. Eur. J. Org. Chem. 2013, 2013 (9), 1716–1725. 10.1002/ejoc.201201442. [DOI] [Google Scholar]
Vázquez-Romero A.; Cárdenas L.; Blasi E.; Verdaguer X.; Riera A. Synthesis of Prostaglandin and Phytoprostane B₁ Via Regioselective Intermolecular Pauson–Khand Reactions. Org. Lett. 2009, 11 (14), 3104–3107. 10.1021/ol901213d. [DOI] [PubMed] [Google Scholar]
Guy A.; Flanagan S.; Durand T.; Oger C.; Galano J.-M. Facile Synthesis of Cyclopentenone B₁- and L₁-Type Phytoprostanes. Front. Chem. 2015, 3, 1–10. 10.3389/fchem.2015.00041. [DOI] [PMC free article] [PubMed] [Google Scholar]
Beretta R.; Giambelli Gallotti M.; Pennè U.; Porta A.; Gil Romero J. F.; Zanoni G.; Vidari G. General Strategy for the Synthesis of B₁ and L₁ Prostanoids: Synthesis of Phytoprostanes (RS)-9-L₁-PhytoP, (R)-9-L₁-PhytoP, (RS)-16-B₁-PhytoP, and (RS)-16-L₁-PhytoP. J. Org. Chem. 2015, 80 (3), 1601–1609. 10.1021/jo502538b. [DOI] [PubMed] [Google Scholar]
Cuyamendous C.; Leung K. S.; Durand T.; Lee J. C.-Y.; Oger C.; Galano J.-M. Synthesis and Discovery of Phytofurans: Metabolites of α-Linolenic Acid Peroxidation. Chem. Commun. 2015, 51 (86), 15696–15699. 10.1039/C5CC05736A. [DOI] [PubMed] [Google Scholar]
Cuyamendous C.; Leung K. S.; Bultel-Poncé V.; Guy A.; Durand T.; Galano J.-M.; Lee J. C.-Y.; Oger C. Total Synthesis and in Vivo Quantitation of Phytofurans Derived from α-Linolenic Acid. Eur. J. Org. Chem. 2017, 2017 (17), 2486–2490. 10.1002/ejoc.201700270. [DOI] [Google Scholar]
Kelly R. W. Physical Methods of Measurement of Prostaglandins. Clin. Endocrinol. Metab. 1973, 2 (3), 375–392. 10.1016/S0300-595X(73)80005-2. [DOI] [PubMed] [Google Scholar]
Borgeat P.; Samuelsson B. Metabolism of Arachidonic Acid in Polymorphonuclear Leukocytes. Structural Analysis of Novel Hydroxylated Compounds. J. Biol. Chem. 1979, 254 (16), 7865–7869. 10.1016/S0021-9258(18)36026-5. [DOI] [PubMed] [Google Scholar]
Ingram C. D.; Brash A. R. Characterization of HETEs and Related Conjugated Dienes by UV Spectroscopy. Lipids 1988, 23 (4), 340–344. 10.1007/BF02537345. [DOI] [PubMed] [Google Scholar]
Jin J.; Boeglin W. E.; Brash A. R. Analysis of 12/15-Lipoxygenase Metabolism of EPA and DHA with Special Attention to Authentication of Docosatrienes. J. Lipid Res. 2021, 62, 100088. 10.1016/j.jlr.2021.100088. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ryhage R. The Mass Spectrometry Laboratory at the Karolinska Institute 1944–1987. Mass Spectrom. Rev. 1993, 12 (1), 1–49. 10.1002/mas.1280120102. [DOI] [Google Scholar]
Tsikas D.; Zoerner A. A. Analysis of Eicosanoids by LC-MS/MS and GC-MS/MS: A Historical Retrospect and a Discussion. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2014, 964, 79–88. 10.1016/j.jchromb.2014.03.017. [DOI] [PubMed] [Google Scholar]
Tsikas D. Application of Gas Chromatography-Mass Spectrometry and Gas Chromatography-Tandem Mass Spectrometry to Assess in Vivo Synthesis of Prostaglandins, Thromboxane, Leukotrienes, Isoprostanes and Related Compounds in Humans. J. Chromatogr. B. Biomed. Sci. Appl. 1998, 717 (1–2), 201–245. 10.1016/S0378-4347(98)00210-2. [DOI] [PubMed] [Google Scholar]
Karara A.; Dishman E.; Blair I.; Falck J. R.; Capdevila J. H. Endogenous Epoxyeicosatrienoic Acids. Cytochrome P-450 Controlled Stereoselectivity of the Hepatic Arachidonic Acid Epoxygenase. J. Biol. Chem. 1989, 264 (33), 19822–19827. 10.1016/S0021-9258(19)47185-8. [DOI] [PubMed] [Google Scholar]
Schulze B.; Lauchli R.; Sonwa M. M.; Schmidt A.; Boland W. Profiling of Structurally Labile Oxylipins in Plants by in Situ Derivatization with Pentafluorobenzyl Hydroxylamine. Anal. Biochem. 2006, 348 (2), 269–283. 10.1016/j.ab.2005.10.021. [DOI] [PubMed] [Google Scholar]
Newman J. W.; Hammock B. D. Optimized Thiol Derivatizing Reagent for the Mass Spectral Analysis of Disubstituted Epoxy Fatty Acids. J. Chromatogr. A 2001, 925 (1–2), 223–240. 10.1016/S0021-9673(01)00998-0. [DOI] [PubMed] [Google Scholar]
Murphy R. C.; Barkley R. M.; Zemski Berry K.; Hankin J.; Harrison K.; Johnson C.; Krank J.; McAnoy A.; Uhlson C.; Zarini S. Electrospray Ionization and Tandem Mass Spectrometry of Eicosanoids. Anal. Biochem. 2005, 346 (1), 1–42. 10.1016/j.ab.2005.04.042. [DOI] [PubMed] [Google Scholar]
Mesaros C.; Lee S. H.; Blair I. A. Targeted Quantitative Analysis of Eicosanoid Lipids in Biological Samples Using Liquid Chromatography-Tandem Mass Spectrometry. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2009, 877 (26), 2736–2745. 10.1016/j.jchromb.2009.03.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
Astarita G.; Kendall A. C.; Dennis E. A.; Nicolaou A. Targeted Lipidomic Strategies for Oxygenated Metabolites of Polyunsaturated Fatty Acids. Biochim. Biophys. Acta 2015, 1851 (4), 456–468. 10.1016/j.bbalip.2014.11.012. [DOI] [PMC free article] [PubMed] [Google Scholar]
Chhonker Y. S.; Bala V.; Murry D. J. Quantification of Eicosanoids and Their Metabolites in Biological Matrices: A Review. Bioanalysis 2018, 10 (24), 2027–2046. 10.4155/bio-2018-0173. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kita Y.; Tokuoka S. M.; Shimizu T. Mediator Lipidomics by Liquid Chromatography-Tandem Mass Spectrometry. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2017, 1862 (8), 777–781. 10.1016/j.bbalip.2017.03.008. [DOI] [PubMed] [Google Scholar]
Norris P. C.; Serhan C. N. Metabololipidomic Profiling of Functional Immunoresolvent Clusters and Eicosanoids in Mammalian Tissues. Biochem. Biophys. Res. Commun. 2018, 504 (3), 553–561. 10.1016/j.bbrc.2018.03.037. [DOI] [PMC free article] [PubMed] [Google Scholar]
Gladine C.; Ostermann A. I.; Newman J. W.; Schebb N. H. MS-Based Targeted Metabolomics of Eicosanoids and Other Oxylipins: Analytical and Inter-Individual Variabilities. Free Radic. Biol. Med. 2019, 144, 72–89. 10.1016/j.freeradbiomed.2019.05.012. [DOI] [PubMed] [Google Scholar]
O’Donnell V. B.; Milne G. L.; Nogueira M. S.; Giera M.; Helge Schebb N.. Quantitation of Oxylipins in Biological Samples, Focusing on Plasma, and Urine. In Mass Spectrometry for Lipidomics; John Wiley & Sons, 2023; pp 317–350. 10.1002/9783527836512.ch12. [DOI] [Google Scholar]
Willenberg I.; Ostermann A. I.; Schebb N. H. Targeted Metabolomics of the Arachidonic Acid Cascade: Current State and Challenges of LC-MS Analysis of Oxylipins. Anal. Bioanal. Chem. 2015, 407 (10), 2675–2683. 10.1007/s00216-014-8369-4. [DOI] [PubMed] [Google Scholar]
Giera M.; Galano J. M.. Eicosanoids. In Encyclopedia of Analytical Science, 3rd ed.; Worsfold P., Poole C., Townshend A., Miró M., Eds.; Academic Press: Oxford, 2019; pp 259–263. 10.1016/B978-0-12-409547-2.13984-8. [DOI] [Google Scholar]
Ostermann A. I.; Willenberg I.; Schebb N. H. Comparison of Sample Preparation Methods for the Quantitative Analysis of Eicosanoids and Other Oxylipins in Plasma by Means of LC-MS/MS. Anal. Bioanal. Chem. 2015, 407 (5), 1403–1414. 10.1007/s00216-014-8377-4. [DOI] [PubMed] [Google Scholar]
Yang W.; Schoeman J. C.; Di X.; Lamont L.; Harms A. C.; Hankemeier T. A Comprehensive UHPLC-MS/MS Method for Metabolomics Profiling of Signaling Lipids: Markers of Oxidative Stress, Immunity and Inflammation. Anal. Chim. Acta 2024, 1297, 342348. 10.1016/j.aca.2024.342348. [DOI] [PubMed] [Google Scholar]
O’Donnell V. B.; Murphy R. C.; Watson S. P. Platelet Lipidomics: Modern Day Perspective on Lipid Discovery and Characterization in Platelets. Circ. Res. 2014, 114 (7), 1185–1203. 10.1161/CIRCRESAHA.114.301597. [DOI] [PMC free article] [PubMed] [Google Scholar]
Aukema H. M.; Ravandi A. Factors Affecting Variability in Free Oxylipins in Mammalian Tissues. Curr. Opin. Clin. Nutr. Metab. Care 2022, 26 (2), 91–98. 10.1097/MCO.0000000000000892. [DOI] [PubMed] [Google Scholar]
Deems R.; Buczynski M. W.; Bowers-Gentry R.; Harkewicz R.; Dennis E. A. Detection and Quantitation of Eicosanoids via High Performance Liquid Chromatography-Electrospray Ionization-Mass Spectrometry. Methods Enzymol. 2007, 432, 59–82. 10.1016/S0076-6879(07)32003-X. [DOI] [PubMed] [Google Scholar]
Yang J.; Schmelzer K.; Georgi K.; Hammock B. D. Quantitative Profiling Method for Oxylipin Metabolome by Liquid Chromatography Electrospray Ionization Tandem Mass Spectrometry. Anal. Chem. 2009, 81 (19), 8085–8093. 10.1021/ac901282n. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yuan Z.-X.; Majchrzak-Hong S.; Keyes G. S.; Iadarola M. J.; Mannes A. J.; Ramsden C. E. Lipidomic Profiling of Targeted Oxylipins with Ultra-Performance Liquid Chromatography-Tandem Mass Spectrometry. Anal. Bioanal. Chem. 2018, 410 (23), 6009–6029. 10.1007/s00216-018-1222-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
Masoodi M.; Eiden M.; Koulman A.; Spaner D.; Volmer D. A. Comprehensive Lipidomics Analysis of Bioactive Lipids in Complex Regulatory Networks. Anal. Chem. 2010, 82 (19), 8176–8185. 10.1021/ac1015563. [DOI] [PubMed] [Google Scholar]
Hinz C.; Liggi S.; Mocciaro G.; Jung S.; Induruwa I.; Pereira M.; Bryant C. E.; Meckelmann S. W.; O’Donnell V. B.; Farndale R. W.; Fjeldsted J.; Griffin J. L. A Comprehensive UHPLC Ion Mobility Quadrupole Time-of-Flight Method for Profiling and Quantification of Eicosanoids, Other Oxylipins, and Fatty Acids. Anal. Chem. 2019, 91 (13), 8025–8035. 10.1021/acs.analchem.8b04615. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ostermann A. I.; Koch E.; Rund K. M.; Kutzner L.; Mainka M.; Schebb N. H. Targeting Esterified Oxylipins by LC-MS - Effect of Sample Preparation on Oxylipin Pattern. Prostaglandins Other Lipid Mediat. 2020, 146, 106384. 10.1016/j.prostaglandins.2019.106384. [DOI] [PubMed] [Google Scholar]
Koch E.; Mainka M.; Dalle C.; Ostermann A. I.; Rund K. M.; Kutzner L.; Froehlich L.-F.; Bertrand-Michel J.; Gladine C.; Schebb N. H. Stability of Oxylipins during Plasma Generation and Long-Term Storage. Talanta 2020, 217, 121074. 10.1016/j.talanta.2020.121074. [DOI] [PubMed] [Google Scholar]
Tsuji K.; Shimada W.; Kishino S.; Ogawa J.; Arita M. Comprehensive Analysis of Fatty Acid Metabolites Produced by Gut Microbiota Using LC-MS/MS-Based Lipidomics. Med. Mass Spectrom. 2022, 6, 112–125. 10.24508/mms.2022.11.003. [DOI] [Google Scholar]
Cebo M.; Fu X.; Gawaz M.; Chatterjee M.; Lämmerhofer M. Micro-UHPLC-MS/MS Method for Analysis of Oxylipins in Plasma and Platelets. J. Pharm. Biomed. Anal. 2020, 189, 113426. 10.1016/j.jpba.2020.113426. [DOI] [PubMed] [Google Scholar]
Yamada M.; Kita Y.; Kohira T.; Yoshida K.; Hamano F.; Tokuoka S. M.; Shimizu T. A Comprehensive Quantification Method for Eicosanoids and Related Compounds by Using Liquid Chromatography/Mass Spectrometry with High Speed Continuous Ionization Polarity Switching. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2015, 995–996, 74–84. 10.1016/j.jchromb.2015.05.015. [DOI] [PubMed] [Google Scholar]
Kampschulte N.; Loewen A.; Kirchhoff R.; Schebb N. Deducing Formation Routes of Oxylipins by Quantitative Multiple Heart-Cutting Achiral-Chiral 2D-LC-MS. J. Lipid Res. 2024, 100694. 10.1016/j.jlr.2024.100694. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wender M.; Adamczewska-Goncerzewicz Z.; Sroczyński E.; Zórawski A. Effect of Cyanide Intoxication on Free Fatty Acid Pattern in the Brain of Rats Developing on a Diet with Various Amounts of Lipids. Neuropathol. Polym. 1986, 24, 341–350. [PubMed] [Google Scholar]
Elloumi A.; Mas-Normand L.; Bride J.; Reversat G.; Bultel-Poncé V.; Guy A.; Oger C.; Demion M.; Le Guennec J.-Y.; Durand T.; Vigor C.; Sánchez-Illana Á.; Galano J.-M. From MS/MS Library Implementation to Molecular Networks: Exploring Oxylipin Diversity with NEO-MSMS. Sci. Data 2024, 11 (1), 193. 10.1038/s41597-024-03034-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
Mantzourani C.; Kokotou M. G. Liquid Chromatography-Mass Spectrometry (LC-MS) Derivatization-Based Methods for the Determination of Fatty Acids in Biological Samples. Mol. Basel Switz. 2022, 27 (17), 5717. 10.3390/molecules27175717. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bollinger J. G.; Thompson W.; Lai Y.; Oslund R. C.; Hallstrand T. S.; Sadilek M.; Turecek F.; Gelb M. H. Improved Sensitivity Mass Spectrometric Detection of Eicosanoids by Charge Reversal Derivatization. Anal. Chem. 2010, 82 (16), 6790–6796. 10.1021/ac100720p. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bollinger J. G.; Rohan G.; Sadilek M.; Gelb M. H. LC/ESI-MS/MS Detection of FAs by Charge Reversal Derivatization with More than Four Orders of Magnitude Improvement in Sensitivity. J. Lipid Res. 2013, 54, 3523–3530. 10.1194/jlr.D040782. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ianni F.; Saluti G.; Galarini R.; Fiorito S.; Sardella R.; Natalini B. Enantioselective High-Performance Liquid Chromatography Analysis of Oxygenated Polyunsaturated Fatty Acids. Free Radic. Biol. Med. 2019, 144, 35–54. 10.1016/j.freeradbiomed.2019.04.038. [DOI] [PubMed] [Google Scholar]
Blum M.; Dogan I.; Karber M.; Rothe M.; Schunck W.-H. Chiral Lipidomics of Monoepoxy and Monohydroxy Metabolites Derived from Long-Chain Polyunsaturated Fatty Acids. J. Lipid Res. 2019, 60 (1), 135–148. 10.1194/jlr.M089755. [DOI] [PMC free article] [PubMed] [Google Scholar]
Cebo M.; Fu X.; Gawaz M.; Chatterjee M.; Lämmerhofer M. Enantioselective Ultra-High Performance Liquid Chromatography-Tandem Mass Spectrometry Method Based on Sub-2μm Particle Polysaccharide Column for Chiral Separation of Oxylipins and Its Application for the Analysis of Autoxidized Fatty Acids and Platelet Releasates. J. Chromatogr. A 2020, 1624, 461206. 10.1016/j.chroma.2020.461206. [DOI] [PubMed] [Google Scholar]
Quaranta A.; Zöhrer B.; Revol-Cavalier J.; Benkestock K.; Balas L.; Oger C.; Keyes G. S.; Wheelock Å. M.; Durand T.; Galano J.-M.; Ramsden C. E.; Hamberg M.; Wheelock C. E. Development of a Chiral Supercritical Fluid Chromatography-Tandem Mass Spectrometry and Reversed-Phase Liquid Chromatography-Tandem Mass Spectrometry Platform for the Quantitative Metabolic Profiling of Octadecanoid Oxylipins. Anal. Chem. 2022, 94 (42), 14618–14626. 10.1021/acs.analchem.2c02601. [DOI] [PMC free article] [PubMed] [Google Scholar]
Salamin O.; Wheelock C. E. Quantification of Octadecanoids in Human Plasma Using Chiral Supercritical Fluid Chromatography-Tandem Mass Spectrometry. Methods Mol. Biol. Clifton NJ. 2025, 2855, 315–339. 10.1007/978-1-0716-4116-3_19. [DOI] [PubMed] [Google Scholar]
de Oliveira M. A. L.; Porto B. L. S.; Faria I. D. L.; de Oliveira P. L.; de Castro Barra P. M.; de Jesus Coelho Castro R.; Sato R. T. 20 Years of Fatty Acid Analysis by Capillary Electrophoresis. Mol. Basel Switz. 2014, 19, 14094–14113. 10.3390/molecules190914094. [DOI] [PMC free article] [PubMed] [Google Scholar]
VanderNoot V. A.; VanRollins M. Capillary Electrophoresis of Cytochrome P-450 Epoxygenase Metabolites of Arachidonic Acid. 1. Resolution of Regioisomers. Anal. Chem. 2002, 74 (22), 5859–5865. 10.1021/ac025909+. [DOI] [PubMed] [Google Scholar]
Vandernoot V. A.; VanRollins M. Capillary Electrophoresis of Cytochrome P-450 Epoxygenase Metabolites of Arachidonic Acid. 2. Resolution of Stereoisomers. Anal. Chem. 2002, 74 (22), 5866–5870. 10.1021/ac0259109. [DOI] [PubMed] [Google Scholar]
Dahl S. R.; Kleiveland C. R.; Kassem M.; Lea T.; Lundanes E.; Greibrokk T. Detecting PM Concentrations of Prostaglandins in Cell Culture Supernatants by Capillary SCX-LC-MS/MS. J. Sep. Sci. 2008, 31, 2627–2633. 10.1002/jssc.200800184. [DOI] [PubMed] [Google Scholar]
Wan H.; Blomberg L. G.; Hamberg M. Highly Efficient Separation of Isomeric Epoxy Fatty Acids by Micellar Electrokinetic Chromatography. Electrophoresis 1999, 20 (1), 132–137. . [DOI] [PubMed] [Google Scholar]
Jussila M.; Sundberg S.; Hopia A.; Mäkinen M.; Riekkola M. L. Separation of Linoleic Acid Oxidation Products by Micellar Electrokinetic Capillary Chromatography and Nonaqueous Capillary Electrophoresis. Electrophoresis 1999, 20 (1), 111–117. . [DOI] [PubMed] [Google Scholar]
Tsikas D.; Suchy M.-T.; Tödter K.; Heeren J.; Scheja L. Utilizing Immunoaffinity Chromatography (IAC) Cross-Reactivity in GC-MS/MS Exemplified at the Measurement of Prostaglandin E₁ in Human Plasma Using Prostaglandin E₂-Specific IAC Columns. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2016, 1021, 101–107. 10.1016/j.jchromb.2015.04.026. [DOI] [PubMed] [Google Scholar]
Kyle J. E.; Aly N.; Zheng X.; Burnum-Johnson K. E.; Smith R. D.; Baker E. S. Evaluating Lipid Mediator Structural Complexity Using Ion Mobility Spectrometry Combined with Mass Spectrometry. Bioanalysis 2018, 10 (5), 279–289. 10.4155/bio-2017-0245. [DOI] [PMC free article] [PubMed] [Google Scholar]
Jónasdóttir H. S.; Papan C.; Fabritz S.; Balas L.; Durand T.; Hardardottir I.; Freysdottir J.; Giera M. Differential Mobility Separation of Leukotrienes and Protectins. Anal. Chem. 2015, 87 (10), 5036–5040. 10.1021/acs.analchem.5b00786. [DOI] [PubMed] [Google Scholar]
da Silva K. M.; Wölk M.; Nepachalovich P.; Iturrospe E.; Covaci A.; van Nuijs A. L. N.; Fedorova M. Investigating the Potential of Drift Tube Ion Mobility for the Analysis of Oxidized Lipids. Anal. Chem. 2023, 95 (36), 13566–13574. 10.1021/acs.analchem.3c02213. [DOI] [PubMed] [Google Scholar]
Duncan K. D.; Sun X.; Baker E. S.; Dey S. K.; Lanekoff I. In Situ Imaging Reveals Disparity between Prostaglandin Localization and Abundance of Prostaglandin Synthases. Commun. Biol. 2021, 4 (1), 966. 10.1038/s42003-021-02488-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
Duncan K. D.; Fang R.; Yuan J.; Chu R. K.; Dey S. K.; Burnum-Johnson K. E.; Lanekoff I. Quantitative Mass Spectrometry Imaging of Prostaglandins as Silver Ion Adducts with Nanospray Desorption Electrospray Ionization. Anal. Chem. 2018, 90 (12), 7246–7252. 10.1021/acs.analchem.8b00350. [DOI] [PMC free article] [PubMed] [Google Scholar]
Mavroudakis L.; Lanekoff I. Identification and Imaging of Prostaglandin Isomers Utilizing MS³ Product Ions and Silver Cationization. J. Am. Soc. Mass Spectrom. 2023, 34 (10), 2341–2349. 10.1021/jasms.3c00233. [DOI] [PMC free article] [PubMed] [Google Scholar]
Weigand M. R.; Unsihuay Vila D. M.; Yang M.; Hu H.; Hernly E.; Muhoberac M.; Tichy S.; Laskin J. Lipid Isobar and Isomer Imaging Using Nanospray Desorption Electrospray Ionization Combined with Triple Quadrupole Mass Spectrometry. Anal. Chem. 2024, 96, 2975. 10.1021/acs.analchem.3c04705. [DOI] [PubMed] [Google Scholar]
Smith M. J.; Nie M.; Adner M.; Säfholm J.; Wheelock C. E. Development of a Desorption Electrospray Ionization-Multiple-Reaction-Monitoring Mass Spectrometry (DESI-MRM) Workflow for Spatially Mapping Oxylipins in Pulmonary Tissue. Anal. Chem. 2024, 96, 17950. 10.1021/acs.analchem.4c02350. [DOI] [PMC free article] [PubMed] [Google Scholar]
Piesche M.; Roos J.; Kühn B.; Fettel J.; Hellmuth N.; Brat C.; Maucher I. V.; Awad O.; Matrone C.; Comerma Steffensen S. G.; Manolikakes G.; Heinicke U.; Zacharowski K. D.; Steinhilber D.; Maier T. J. The Emerging Therapeutic Potential of Nitro Fatty Acids and Other Michael Acceptor-Containing Drugs for the Treatment of Inflammation and Cancer. Front. Pharmacol. 2020, 11, 1297. 10.3389/fphar.2020.01297. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schopfer F. J.; Khoo N. K. H. Nitro-Fatty Acid Logistics: Formation, Biodistribution, Signaling, and Pharmacology. Trends Endocrinol. Metab. 2019, 30 (8), 505–519. 10.1016/j.tem.2019.04.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fang M.-Y.; Huang K.-H.; Tu W.-J.; Chen Y.-T.; Pan P.-Y.; Hsiao W.-C.; Ke Y.-Y.; Tsou L. K.; Zhang M. M. Chemoproteomic Profiling Reveals Cellular Targets of Nitro-Fatty Acids. Redox Biol. 2021, 46, 102126. 10.1016/j.redox.2021.102126. [DOI] [PMC free article] [PubMed] [Google Scholar]
Vangaveti V.; Baune B. T.; Kennedy R. L. Review: Hydroxyoctadecadienoic Acids: Novel Regulators of Macrophage Differentiation and Atherogenesis. Ther. Adv. Endocrinol. Metab. 2010, 1 (2), 51–60. 10.1177/2042018810375656. [DOI] [PMC free article] [PubMed] [Google Scholar]
Singh N. K.; Rao G. N. Emerging Role of 12/15-Lipoxygenase (ALOX15) in Human Pathologies. Prog. Lipid Res. 2019, 73, 28–45. 10.1016/j.plipres.2018.11.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
Collin F. Chemical Basis of Reactive Oxygen Species Reactivity and Involvement in Neurodegenerative Diseases. Int. J. Mol. Sci. 2019, 20 (10), 2407. 10.3390/ijms20102407. [DOI] [PMC free article] [PubMed] [Google Scholar]
Oliw E. H.; Brodowsky I. D.; Hornsten L.; Hamberg M. Bis-Allylic Hydroxylation of Polyunsaturated Fatty Acids by Hepatic Monooxygenases and Its Relation to the Enzymatic and Nonenzymatic Formation of Conjugated Hydroxy Fatty Acids. Arch. Biochem. Biophys. 1993, 300 (1), 434–439. 10.1006/abbi.1993.1059. [DOI] [PubMed] [Google Scholar]
Corteselli E. M.; Gibbs-Flournoy E.; Simmons S. O.; Bromberg P.; Gold A.; Samet J. M. Long Chain Lipid Hydroperoxides Increase the Glutathione Redox Potential through Glutathione Peroxidase 4. Biochim. Biophys. Acta BBA - Gen. Subj. 2019, 1863 (5), 950–959. 10.1016/j.bbagen.2019.03.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Delerive P.; Furman C.; Teissier E.; Fruchart J.-C.; Duriez P.; Staels B. Oxidized Phospholipids Activate PPARα in a Phospholipase A₂-Dependent Manner. FEBS Lett. 2000, 471 (1), 34–38. 10.1016/S0014-5793(00)01364-8. [DOI] [PubMed] [Google Scholar]
Lecka-Czernik B.; Moerman E. J.; Grant D. F.; Lehmann J. M.; Manolagas S. C.; Jilka R. L. Divergent Effects of Selective Peroxisome Proliferator-Activated Receptor-Γ2 Ligands on Adipocyte Versus Osteoblast Differentiation. Endocrinology 2002, 143 (6), 2376–2384. 10.1210/endo.143.6.8834. [DOI] [PubMed] [Google Scholar]
Benítez-Angeles M.; Morales-Lázaro S. L.; Juárez-González E.; Rosenbaum T. TRPV1: Structure, Endogenous Agonists, and Mechanisms. Int. J. Mol. Sci. 2020, 21 (10), 3421. 10.3390/ijms21103421. [DOI] [PMC free article] [PubMed] [Google Scholar]
Obinata H.; Hattori T.; Nakane S.; Tatei K.; Izumi T. Identification of 9-Hydroxyoctadecadienoic Acid and Other Oxidized Free Fatty Acids as Ligands of the G Protein-Coupled Receptor G2A. J. Biol. Chem. 2005, 280 (49), 40676–40683. 10.1074/jbc.M507787200. [DOI] [PubMed] [Google Scholar]
Wheeler J. J.; Domenichiello A. F.; Jensen J. R.; Keyes G. S.; Maiden K. M.; Davis J. M.; Ramsden C. E.; Mishra S. K. Endogenous Derivatives of Linoleic Acid and Their Stable Analogs Are Potential Pain Mediators. JID Innov. Skin Sci. Mol. Popul. Health 2023, 3 (2), 100177. 10.1016/j.xjidi.2022.100177. [DOI] [PMC free article] [PubMed] [Google Scholar]
Buchanan M.; Bertomeu M.; Haas T.; Orr F.; Eltringham-Smith L. Localization of 13-Hydroxyoctadecadienoic Acid and the Vitronectin Receptor in Human Endothelial Cells and Endothelial Cell/Platelet Interactions in Vitro. Blood 1993, 81 (12), 3303–3312. 10.1182/blood.V81.12.3303.3303. [DOI] [PubMed] [Google Scholar]
Frömel T.; Jungblut B.; Hu J.; Trouvain C.; Barbosa-Sicard E.; Popp R.; Liebner S.; Dimmeler S.; Hammock B. D.; Fleming I. Soluble Epoxide Hydrolase Regulates Hematopoietic Progenitor Cell Function via Generation of Fatty Acid Diols. Proc. Natl. Acad. Sci. U. S. A. 2012, 109 (25), 9995–10000. 10.1073/pnas.1206493109. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hohmann S. W.; Angioni C.; Tunaru S.; Lee S.; Woolf C. J.; Offermanns S.; Geisslinger G.; Scholich K.; Sisignano M. The G2A Receptor (GPR132) Contributes to Oxaliplatin-Induced Mechanical Pain Hypersensitivity. Sci. Rep. 2017, 7 (1), 446. 10.1038/s41598-017-00591-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lecarpentier Y.; Claes V.; Vallée A.; Hébert J.-L. Interactions between PPAR Gamma and the Canonical Wnt/Beta-Catenin Pathway in Type 2 Diabetes and Colon Cancer. PPAR Res. 2017, 2017, 5879090. 10.1155/2017/5879090. [DOI] [PMC free article] [PubMed] [Google Scholar]
Cheng W. Y.; Huynh H.; Chen P.; Peña-Llopis S.; Wan Y. Macrophage PPARγ Inhibits Gpr132 to Mediate the Anti-Tumor Effects of Rosiglitazone. eLife 2016, 5, e18501 10.7554/eLife.18501. [DOI] [PMC free article] [PubMed] [Google Scholar]
Vangaveti V.; Shashidhar V.; Collier F.; Hodge J.; Rush C.; Malabu U.; Baune B.; Kennedy R. L. 9- and 13-HODE Regulate Fatty Acid Binding Protein-4 in Human Macrophages, but Does Not Involve HODE/GPR132 Axis in PPAR-γ Regulation of FABP4. Ther. Adv. Endocrinol. Metab. 2018, 9 (5), 137–150. 10.1177/2042018818759894. [DOI] [PMC free article] [PubMed] [Google Scholar]
Xu X.; Otsuki M.; Saito H.; Sumitani S.; Yamamoto H.; Asanuma N.; Kouhara H.; Kasayama S. PPARalpha and GR Differentially Down-Regulate the Expression of Nuclear Factor-KappaB-Responsive Genes in Vascular Endothelial Cells. Endocrinology 2001, 142 (8), 3332–3339. 10.1210/endo.142.8.8340. [DOI] [PubMed] [Google Scholar]
Parnova R. G. GPR40/FFA1 Free Fatty Acid Receptors and Their Functional Role. Neurosci. Behav. Physiol. 2021, 51 (2), 256–264. 10.1007/s11055-021-01064-8. [DOI] [Google Scholar]
Huang J. T.; Welch J. S.; Ricote M.; Binder C. J.; Willson T. M.; Kelly C.; Witztum J. L.; Funk C. D.; Conrad D.; Glass C. K. Interleukin-4-Dependent Production of PPAR-γ Ligands in Macrophages by 12/15-Lipoxygenase. Nature 1999, 400 (6742), 378–382. 10.1038/22572. [DOI] [PubMed] [Google Scholar]
Shureiqi I.; Jiang W.; Zuo X.; Wu Y.; Stimmel J. B.; Leesnitzer L. M.; Morris J. S.; Fan H.-Z.; Fischer S. M.; Lippman S. M. The 15-Lipoxygenase-1 Product 13-S-Hydroxyoctadecadienoic Acid down-Regulates PPAR- to Induce Apoptosis in Colorectal Cancer Cells. Proc. Natl. Acad. Sci. U. S. A. 2003, 100 (17), 9968–9973. 10.1073/pnas.1631086100. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zuo X.; Wu Y.; Morris J. S.; Stimmel J. B.; Leesnitzer L. M.; Fischer S. M.; Lippman S. M.; Shureiqi I. Oxidative Metabolism of Linoleic Acid Modulates PPAR-Beta/Delta Suppression of PPAR-Gamma Activity. Oncogene 2006, 25 (8), 1225–1241. 10.1038/sj.onc.1209160. [DOI] [PMC free article] [PubMed] [Google Scholar]
Enayetallah A.; Cao L.; Grant D. F. Novel Role of Soluble Epoxide Hydrolase in Regulating Cholesterol in Mammalian Cells. Open Drug Metab. J. 2007, 1, 1–6. 10.2174/1874073100701010001. [DOI] [PMC free article] [PubMed] [Google Scholar]
Schopfer F. J.; Lin Y.; Baker P. R. S.; Cui T.; Garcia-Barrio M.; Zhang J.; Chen K.; Chen Y. E.; Freeman B. A. Nitrolinoleic Acid: An Endogenous Peroxisome Proliferator-Activated Receptor γ Ligand. Proc. Natl. Acad. Sci. U. S. A. 2005, 102 (7), 2340–2345. 10.1073/pnas.0408384102. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zhao L.; Funk C. D. Lipoxygenase Pathways in Atherogenesis. Trends Cardiovasc. Med. 2004, 14 (5), 191–195. 10.1016/j.tcm.2004.04.003. [DOI] [PubMed] [Google Scholar]
Umeno A.; Sakashita M.; Sugino S.; Murotomi K.; Okuzawa T.; Morita N.; Tomii K.; Tsuchiya Y.; Yamasaki K.; Horie M.; Takahara K.; Yoshida Y. Comprehensive Analysis of PPARγ Agonist Activities of Stereo-, Regio-, and Enantio-Isomers of Hydroxyoctadecadienoic Acids. Biosci. Rep. 2020, 40 (4), BSR20193767. 10.1042/BSR20193767. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lin P.; Ye R. D. The Lysophospholipid Receptor G2A Activates a Specific Combination of G Proteins and Promotes Apoptosis. J. Biol. Chem. 2003, 278 (16), 14379–14386. 10.1074/jbc.M209101200. [DOI] [PubMed] [Google Scholar]
Obinata H.; Izumi T. G2A as a Receptor for Oxidized Free Fatty Acids. Prostaglandins Other Lipid Mediat. 2009, 89 (3–4), 66–72. 10.1016/j.prostaglandins.2008.11.002. [DOI] [PubMed] [Google Scholar]
Yin H.; Chu A.; Li W.; Wang B.; Shelton F.; Otero F.; Nguyen D. G.; Caldwell J. S.; Chen Y. A. Lipid G Protein-Coupled Receptor Ligand Identification Using β-Arrestin PathHunter^TM Assay. J. Biol. Chem. 2009, 284 (18), 12328–12338. 10.1074/jbc.M806516200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hattori T.; Obinata H.; Ogawa A.; Kishi M.; Tatei K.; Ishikawa O.; Izumi T. G2A Plays Proinflammatory Roles in Human Keratinocytes under Oxidative Stress as a Receptor for 9-Hydroxyoctadecadienoic Acid. J. Invest. Dermatol. 2008, 128 (5), 1123–1133. 10.1038/sj.jid.5701172. [DOI] [PubMed] [Google Scholar]
Kämmerer I.; Ringseis R.; Biemann R.; Wen G.; Eder K. 13-Hydroxy Linoleic Acid Increases Expression of the Cholesterol Transporters ABCA1, ABCG1 and SR-BI and Stimulates ApoA-I-Dependent Cholesterol Efflux in RAW264.7 Macrophages. Lipids Health Dis. 2011, 10 (1), 222. 10.1186/1476-511X-10-222. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hohmann S. W.; Angioni C.; Tunaru S.; Lee S.; Woolf C. J.; Offermanns S.; Geisslinger G.; Scholich K.; Sisignano M. The G2A Receptor (GPR132) Contributes to Oxaliplatin-Induced Mechanical Pain Hypersensitivity. Sci. Rep. 2017, 7 (1), 446. 10.1038/s41598-017-00591-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
De Petrocellis L.; Schiano Moriello A.; Imperatore R.; Cristino L.; Starowicz K.; Di Marzo V. A Re-Evaluation of 9-HODE Activity at TRPV1 Channels in Comparison with Anandamide: Enantioselectivity and Effects at Other TRP Channels and in Sensory Neurons: HODEs versus Anandamide as Endovanilloids. Br. J. Pharmacol. 2012, 167 (8), 1643–1651. 10.1111/j.1476-5381.2012.02122.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
Alsalem M.; Wong A.; Millns P.; Arya P. H.; Chan M. S. L.; Bennett A.; Barrett D. A.; Chapman V.; Kendall D. A. The Contribution of the Endogenous TRPV1 Ligands 9-HODE and 13-HODE to Nociceptive Processing and Their Role in Peripheral Inflammatory Pain Mechanisms: Linoleic Acid Metabolites and Inflammatory Pain. Br. J. Pharmacol. 2013, 168 (8), 1961–1974. 10.1111/bph.12092. [DOI] [PMC free article] [PubMed] [Google Scholar]
Green D. P.; Ruparel S.; Gao X.; Ruparel N.; Patil M.; Akopian A.; Hargreaves K. M. Central Activation of TRPV1 and TRPA1 by Novel Endogenous Agonists Contributes to Mechanical and Thermal Allodynia after Burn Injury. Mol. Pain 2016, 12, 1744806916661725. 10.1177/1744806916661725. [DOI] [PMC free article] [PubMed] [Google Scholar]
Vitturi D. A.; Chen C.-S.; Woodcock S. R.; Salvatore S. R.; Bonacci G.; Koenitzer J. R.; Stewart N. A.; Wakabayashi N.; Kensler T. W.; Freeman B. A.; Schopfer F. J. Modulation of Nitro-Fatty Acid Signaling: Prostaglandin Reductase-1 Is a Nitroalkene Reductase. J. Biol. Chem. 2013, 288 (35), 25626–25637. 10.1074/jbc.M113.486282. [DOI] [PMC free article] [PubMed] [Google Scholar]
Aranda-Caño L.; Sánchez-Calvo B.; Begara-Morales J. C.; Chaki M.; Mata-Pérez C.; Padilla M. N.; Valderrama R.; Barroso J. B. Post-Translational Modification of Proteins Mediated by Nitro-Fatty Acids in Plants: Nitroalkylation. Plants 2019, 8 (4), 82. 10.3390/plants8040082. [DOI] [PMC free article] [PubMed] [Google Scholar]
Natarajan R.; Reddy M. A.; Malik K. U.; Fatima S.; Khan B. V. Signaling Mechanisms of Nuclear Factor-ΚB-Mediated Activation of Inflammatory Genes by 13-Hydroperoxyoctadecadienoic Acid in Cultured Vascular Smooth Muscle Cells. Arterioscler. Thromb. Vasc. Biol. 2001, 21 (9), 1408–1413. 10.1161/hq0901.095278. [DOI] [PubMed] [Google Scholar]
Nagy L.; Tontonoz P.; Alvarez J. G. A.; Chen H.; Evans R. M. Oxidized LDL Regulates Macrophage Gene Expression through Ligand Activation of PPARγ. Cell 1998, 93 (2), 229–240. 10.1016/S0092-8674(00)81574-3. [DOI] [PubMed] [Google Scholar]
Iversen L.; Fogh K.; Bojesen G.; Kragballe K. Linoleic Acid and Dihomogammalinolenic Acid Inhibit Leukotriene B4 Formation and Stimulate the Formation of Their 15-Lipoxygenase Products by Human Neutrophilsin Vitro. Evidence of Formation of Antiinflammatory Compounds. Agents Actions 1991, 33 (3), 286–291. 10.1007/BF01986575. [DOI] [PubMed] [Google Scholar]
van de Velde M. J.; Engels F.; Henricks P. A. J.; Bloemen P. G. M.; Nijkamp F. P. The Linoleic Acid Metabolite 13-HODE Modulates Degranulation of Human Polymorphonuclear Leukocytes. FEBS Lett. 1995, 369 (2–3), 301–304. 10.1016/0014-5793(95)00773-3. [DOI] [PubMed] [Google Scholar]
Frasch S. C.; Fernandez-Boyanapalli R. F.; Berry K. Z.; Leslie C. C.; Bonventre J. V.; Murphy R. C.; Henson P. M.; Bratton D. L. Signaling via Macrophage G2A Enhances Efferocytosis of Dying Neutrophils by Augmentation of Rac Activity. J. Biol. Chem. 2011, 286 (14), 12108–12122. 10.1074/jbc.M110.181800. [DOI] [PMC free article] [PubMed] [Google Scholar]
Qin X.; Qiu C.; Zhao L. Lysophosphatidylcholine Perpetuates Macrophage Polarization toward Classically Activated Phenotype in Inflammation. Cell. Immunol. 2014, 289 (1–2), 185–190. 10.1016/j.cellimm.2014.04.010. [DOI] [PubMed] [Google Scholar]
Cambiaggi L.; Chakravarty A.; Noureddine N.; Hersberger M. The Role of α-Linolenic Acid and Its Oxylipins in Human Cardiovascular Diseases. Int. J. Mol. Sci. 2023, 24 (7), 6110. 10.3390/ijms24076110. [DOI] [PMC free article] [PubMed] [Google Scholar]
Pauls S. D.; Rodway L. A.; Winter T.; Taylor C. G.; Zahradka P.; Aukema H. M. Anti-Inflammatory Effects of α-Linolenic Acid in M1-like Macrophages Are Associated with Enhanced Production of Oxylipins from α-Linolenic and Linoleic Acid. J. Nutr. Biochem. 2018, 57, 121–129. 10.1016/j.jnutbio.2018.03.020. [DOI] [PubMed] [Google Scholar]
Totani Y.; Saito Y.; Ishizaki T.; Sasaki F.; Ameshima S.; Miyamori I. Leukotoxin and Its Diol Induce Neutrophil Chemotaxis through Signal Transduction Different from That of FMLP. Eur. Respir. J. 2000, 15 (1), 75–79. 10.1183/09031936.00.15107500. [DOI] [PubMed] [Google Scholar]
Thompson D. A.; Hammock B. D. Dihydroxyoctadecamonoenoate Esters Inhibit the Neutrophil Respiratory Burst. J. Biosci. 2007, 32 (2), 279–291. 10.1007/s12038-007-0028-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
Edwards L. M.; Lawler N. G.; Nikolic S. B.; Peters J. M.; Horne J.; Wilson R.; Davies N. W.; Sharman J. E. Metabolomics Reveals Increased Isoleukotoxin Diol (12,13-DHOME) in Human Plasma after Acute Intralipid Infusion. J. Lipid Res. 2012, 53 (9), 1979–1986. 10.1194/jlr.P027706. [DOI] [PMC free article] [PubMed] [Google Scholar]
Valencia R.; Kranrod J. W.; Fang L.; Soliman A. M.; Azer B.; Clemente-Casares X.; Seubert J. M. Linoleic Acid-Derived Diol 12,13-DiHOME Enhances NLRP3 Inflammasome Activation in Macrophages. FASEB J. Off. Publ. Fed. Am. Soc. Exp. Biol. 2024, 38 (13), e23748 10.1096/fj.202301640RR. [DOI] [PubMed] [Google Scholar]
Ohue-Kitano R.; Yasuoka Y.; Goto T.; Kitamura N.; Park S.; Kishino S.; Kimura I.; Kasubuchi M.; Takahashi H.; Li Y.; Yeh Y.; Jheng H.; Iwase M.; Tanaka M.; Masuda S.; Inoue T.; Yamakage H.; Kusakabe T.; Tani F.; Shimatsu A.; Takahashi N.; Ogawa J.; Satoh-Asahara N.; Kawada T. Α-Linolenic Acid-derived Metabolites from Gut Lactic Acid Bacteria Induce Differentiation of Anti-inflammatory M2Macrophages through G Protein-coupled Receptor 40. FASEB J. 2018, 32 (1), 304–318. 10.1096/fj.201700273R. [DOI] [PubMed] [Google Scholar]
Goto T.; Kim Y.-I.; Furuzono T.; Takahashi N.; Yamakuni K.; Yang H.-E.; Li Y.; Ohue R.; Nomura W.; Sugawara T.; Yu R.; Kitamura N.; Park S.-B.; Kishino S.; Ogawa J.; Kawada T. 10-Oxo-12(Z)-Octadecenoic Acid, a Linoleic Acid Metabolite Produced by Gut Lactic Acid Bacteria, Potently Activates PPARγ and Stimulates Adipogenesis. Biochem. Biophys. Res. Commun. 2015, 459 (4), 597–603. 10.1016/j.bbrc.2015.02.154. [DOI] [PubMed] [Google Scholar]
Miyamoto J.; Mizukure T.; Park S.-B.; Kishino S.; Kimura I.; Hirano K.; Bergamo P.; Rossi M.; Suzuki T.; Arita M.; Ogawa J.; Tanabe S. A Gut Microbial Metabolite of Linoleic Acid, 10-Hydroxy-cis-12-Octadecenoic Acid, Ameliorates Intestinal Epithelial Barrier Impairment Partially via GPR40-MEK-ERK Pathway. J. Biol. Chem. 2015, 290 (5), 2902–2918. 10.1074/jbc.M114.610733. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bergamo P.; Luongo D.; Miyamoto J.; Cocca E.; Kishino S.; Ogawa J.; Tanabe S.; Rossi M. Immunomodulatory Activity of a Gut Microbial Metabolite of Dietary Linoleic Acid, 10-Hydroxy-Cis-12-Octadecenoic Acid, Associated with Improved Antioxidant/Detoxifying Defences. J. Funct. Foods 2014, 11, 192–202. 10.1016/j.jff.2014.10.007. [DOI] [Google Scholar]
Cui T.; Schopfer F. J.; Zhang J.; Chen K.; Ichikawa T.; Baker Paul R. S.; Batthyany C.; Chacko B. K.; Feng X.; Patel R. P.; Agarwal A.; Freeman B. A.; Chen Y. E. Nitrated Fatty Acids: Endogenous Anti-Inflammatory Signaling Mediators. J. Biol. Chem. 2006, 281 (47), 35686–35698. 10.1074/jbc.M603357200. [DOI] [PMC free article] [PubMed] [Google Scholar]
Freeman B. A.; O’Donnell V. B.; Schopfer F. J. The Discovery of Nitro-Fatty Acids as Products of Metabolic and Inflammatory Reactions and Mediators of Adaptive Cell Signaling. Nitric Oxide Biol. Chem. 2018, 77, 106–111. 10.1016/j.niox.2018.05.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hansen A. L.; Rahbek L. S. J.; Sørensen A. S.; Hundahl M. P.; Lomholt S.; Holm C. K.; Kragstrup T. W. Nitro-Fatty Acids Decrease Type I Interferons and Monocyte Chemoattractant Protein 1 in Ex Vivo Models of Inflammatory Arthritis. BMC Immunol. 2021, 22 (1), 77. 10.1186/s12865-021-00471-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
Villacorta L.; Minarrieta L.; Salvatore S. R.; Khoo N. K.; Rom O.; Gao Z.; Berman R. C.; Jobbagy S.; Li L.; Woodcock S. R.; Chen Y. E.; Freeman B. A.; Ferreira A. M.; Schopfer F. J.; Vitturi D. A. In Situ Generation, Metabolism and Immunomodulatory Signaling Actions of Nitro-Conjugated Linoleic Acid in a Murine Model of Inflammation. Redox Biol. 2018, 15, 522–531. 10.1016/j.redox.2018.01.005. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wang W.; Li C.; Yang T. Protection of Nitro-Fatty Acid against Kidney Diseases. Am. J. Physiol.-Ren. Physiol. 2016, 310 (8), F697–F704. 10.1152/ajprenal.00321.2015. [DOI] [PMC free article] [PubMed] [Google Scholar]
Khoo N. K. H.; Li L.; Salvatore S. R.; Schopfer F. J.; Freeman B. A. Electrophilic Fatty Acid Nitroalkenes Regulate Nrf2 and NF-ΚB Signaling: A Medicinal Chemistry Investigation of Structure-Function Relationships. Sci. Rep. 2018, 8 (1), 2295. 10.1038/s41598-018-20460-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
Cole M. P.; Rudolph T. K.; Khoo N. K. H.; Motanya U. N.; Golin-Bisello F.; Wertz J. W.; Schopfer F. J.; Rudolph V.; Woodcock S. R.; Bolisetty S.; Ali M. S.; Zhang J.; Chen Y. E.; Agarwal A.; Freeman B. A.; Bauer P. M. Nitro-Fatty Acid Inhibition of Neointima Formation After Endoluminal Vessel Injury. Circ. Res. 2009, 105 (10), 965–972. 10.1161/CIRCRESAHA.109.199075. [DOI] [PMC free article] [PubMed] [Google Scholar]
Barber G. N. STING: Infection, Inflammation and Cancer. Nat. Rev. Immunol. 2015, 15 (12), 760–770. 10.1038/nri3921. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fritsch L. E.; Kelly C.; Pickrell A. M. The Role of STING Signaling in Central Nervous System Infection and Neuroinflammatory Disease. WIREs Mech. Dis. 2023, 15 (3), e1597 10.1002/wsbm.1597. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hansen A. L.; Buchan G. J.; Rühl M.; Mukai K.; Salvatore S. R.; Ogawa E.; Andersen S. D.; Iversen M. B.; Thielke A. L.; Gunderstofte C.; Motwani M.; Møller C. T.; Jakobsen A. S.; Fitzgerald K. A.; Roos J.; Lin R.; Maier T. J.; Goldbach-Mansky R.; Miner C. A.; Qian W.; Miner J. J.; Rigby R. E.; Rehwinkel J.; Jakobsen M. R.; Arai H.; Taguchi T.; Schopfer F. J.; Olagnier D.; Holm C. K. Nitro-Fatty Acids Are Formed in Response to Virus Infection and Are Potent Inhibitors of STING Palmitoylation and Signaling. Proc. Natl. Acad. Sci. U. S. A. 2018, 115 (33), E7768-E7775 10.1073/pnas.1806239115. [DOI] [PMC free article] [PubMed] [Google Scholar]
Awwad K.; Steinbrink S. D.; Frömel T.; Lill N.; Isaak J.; Häfner A.-K.; Roos J.; Hofmann B.; Heide H.; Geisslinger G.; Steinhilber D.; Freeman B. A.; Maier T. J.; Fleming I. Electrophilic Fatty Acid Species Inhibit 5-Lipoxygenase and Attenuate Sepsis-Induced Pulmonary Inflammation. Antioxid. Redox Signal. 2014, 20 (17), 2667–2680. 10.1089/ars.2013.5473. [DOI] [PMC free article] [PubMed] [Google Scholar]
Das Mahapatra A.; Choubey R.; Datta B. Small Molecule Soluble Epoxide Hydrolase Inhibitors in Multitarget and Combination Therapies for Inflammation and Cancer. Molecules 2020, 25 (23), 5488. 10.3390/molecules25235488. [DOI] [PMC free article] [PubMed] [Google Scholar]
Dai N.; Yang C.; Fan Q.; Wang M.; Liu X.; Zhao H.; Zhao C. The Anti-Inflammatory Effect of Soluble Epoxide Hydrolase Inhibitor and 14,15-EET in Kawasaki Disease Through PPARγ/STAT1 Signaling Pathway. Front. Pediatr. 2020, 8, 451. 10.3389/fped.2020.00451. [DOI] [PMC free article] [PubMed] [Google Scholar]
Burr G. O.; Burr M. M. A New Deficiency Disease Produced By The Rigid Exclusion Of Fat From The Diet. J. Biol. Chem. 1929, 82 (2), 345–367. 10.1016/S0021-9258(20)78281-5. [DOI] [PubMed] [Google Scholar]
Hansen H. S.; Jensen B.; von Wettstein-Knowles P. Apparent in Vivo Retroconversion of Dietary Arachidonic to Linoleic Acid in Essential Fatty Acid-Deficient Rats. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1986, 878 (2), 284–287. 10.1016/0005-2760(86)90158-X. [DOI] [PubMed] [Google Scholar]
Fischer J.; Bourrat E. Genetics of Inherited Ichthyoses and Related Diseases. Acta Derm. Venereol. 2020, 100 (7), 186–196. 10.2340/00015555-3432. [DOI] [PMC free article] [PubMed] [Google Scholar]
Jobard F.; Lefèvre C.; Karaduman A.; Blanchet-Bardon C.; Emre S.; Weissenbach J.; Özgüc M.; Lathrop M.; Prud’homme J.-F.; Fischer J. Lipoxygenase-3 (ALOXE3) and 12(R)-Lipoxygenase (ALOX12B) Are Mutated in Non-Bullous Congenital Ichthyosiform Erythroderma (NCIE) Linked to Chromosome 17p13.1. Hum. Mol. Genet. 2002, 11 (1), 107–113. 10.1093/hmg/11.1.107. [DOI] [PubMed] [Google Scholar]
Takeichi T.; Hirabayashi T.; Miyasaka Y.; Kawamoto A.; Okuno Y.; Taguchi S.; Tanahashi K.; Murase C.; Takama H.; Tanaka K.; Boeglin W. E.; Calcutt M. W.; Watanabe D.; Kono M.; Muro Y.; Ishikawa J.; Ohno T.; Brash A. R.; Akiyama M. SDR9C7 Catalyzes Critical Dehydrogenation of Acylceramides for Skin Barrier Formation. J. Clin. Invest. 2019, 130 (2), 890–903. 10.1172/JCI130675. [DOI] [PMC free article] [PubMed] [Google Scholar]
Epp N.; Fürstenberger G.; Müller K.; de Juanes S.; Leitges M.; Hausser I.; Thieme F.; Liebisch G.; Schmitz G.; Krieg P. 12R-Lipoxygenase Deficiency Disrupts Epidermal Barrier Function. J. Cell Biol. 2007, 177 (1), 173–182. 10.1083/jcb.200612116. [DOI] [PMC free article] [PubMed] [Google Scholar]
Moran J. L.; Qiu H.; Turbe-Doan A.; Yun Y.; Boeglin W. E.; Brash A. R.; Beier D. R. A Mouse Mutation in the 12R-Lipoxygenase, Alox12b, Disrupts Formation of the Epidermal Permeability Barrier. J. Invest. Dermatol. 2007, 127 (8), 1893–1897. 10.1038/sj.jid.5700825. [DOI] [PubMed] [Google Scholar]
Krieg P.; Rosenberger S.; de Juanes S.; Latzko S.; Hou J.; Dick A.; Kloz U.; van der Hoeven F.; Hausser I.; Esposito I.; Rauh M.; Schneider H. Aloxe3 Knockout Mice Reveal a Function of Epidermal Lipoxygenase-3 as Hepoxilin Synthase and Its Pivotal Role in Barrier Formation. J. Invest. Dermatol. 2013, 133 (1), 172–180. 10.1038/jid.2012.250. [DOI] [PubMed] [Google Scholar]
Ohno Y.; Nakamura T.; Iwasaki T.; Katsuyama A.; Ichikawa S.; Kihara A. Determining the Structure of Protein-Bound Ceramides, Essential Lipids for Skin Barrier Function. iScience 2023, 26 (11), 108248. 10.1016/j.isci.2023.108248. [DOI] [PMC free article] [PubMed] [Google Scholar]
Elias P. M.; Gruber R.; Crumrine D.; Menon G.; Williams M. L.; Wakefield J. S.; Holleran W. M.; Uchida Y. Formation and Functions of the Corneocyte Lipid Envelope (CLE). Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2014, 1841 (3), 314–318. 10.1016/j.bbalip.2013.09.011. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hidalgo F. J.; Zamora R. In Vitro Production of Long Chain Pyrrole Fatty Esters from Carbonyl-Amine Reactions. J. Lipid Res. 1995, 36 (4), 725–735. 10.1016/S0022-2275(20)40058-6. [DOI] [PubMed] [Google Scholar]
Hidalgo F. J.; Zamora R. Epoxyoxoene Fatty Esters: Key Intermediates for the Synthesis of Long-Chain Pyrrole and Furan Fatty Esters. Chem. Phys. Lipids 1995, 77, 1–11. 10.1016/0009-3084(95)02449-S. [DOI] [Google Scholar]
Marekov L. N.; Steinert P. M. Ceramides Are Bound to Structural Proteins of the Human Foreskin Epidermal Cornified Cell Envelope. J. Biol. Chem. 1998, 273 (28), 17763–17770. 10.1074/jbc.273.28.17763. [DOI] [PubMed] [Google Scholar]
Mathers A. R.; Carey C. D.; Killeen M. E.; Salvatore S. R.; Ferris L. K.; Freeman B. A.; Schopfer F. J.; Falo L. D. Topical Electrophilic Nitro-Fatty Acids Potentiate Cutaneous Inflammation. Free Radic. Biol. Med. 2018, 115, 31–42. 10.1016/j.freeradbiomed.2017.11.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
Stewart M. E.; Grahek M. O.; Cambier L. S.; Wertz P. W.; Downing D. T. Dilutional Effect of Increased Sebaceous Gland Activity on the Proportion of Linoleic Acid in Sebaceous Wax Esters and in Epidermal Acylceramides. J. Invest. Dermatol. 1986, 87 (6), 733–736. 10.1111/1523-1747.ep12456856. [DOI] [PubMed] [Google Scholar]
Osthues T.; Sisignano M. Oxidized Lipids in Persistent Pain States. Front. Pharmacol. 2019, 10, 1147. 10.3389/fphar.2019.01147. [DOI] [PMC free article] [PubMed] [Google Scholar]
Patwardhan A. M.; Scotland P. E.; Akopian A. N.; Hargreaves K. M. Activation of TRPV1 in the Spinal Cord by Oxidized Linoleic Acid Metabolites Contributes to Inflammatory Hyperalgesia. Proc. Natl. Acad. Sci. U. S. A. 2009, 106 (44), 18820–18824. 10.1073/pnas.0905415106. [DOI] [PMC free article] [PubMed] [Google Scholar]
Shapiro H.; Singer P.; Ariel A. Beyond the Classic Eicosanoids: Peripherally-Acting Oxygenated Metabolites of Polyunsaturated Fatty Acids Mediate Pain Associated with Tissue Injury and Inflammation. Prostaglandins Leukot. Essent. Fatty Acids 2016, 111, 45–61. 10.1016/j.plefa.2016.03.001. [DOI] [PubMed] [Google Scholar]
Patwardhan A. M.; Akopian A. N.; Ruparel N. B.; Diogenes A.; Weintraub S. T.; Uhlson C.; Murphy R. C.; Hargreaves K. M. Heat Generates Oxidized Linoleic Acid Metabolites That Activate TRPV1 and Produce Pain in Rodents. J. Clin. Invest. 2010, 120 (5), 1617–1626. 10.1172/JCI41678. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wedel S.; Osthues T.; Zimmer B.; Angioni C.; Geisslinger G.; Sisignano M. Oxidized Linoleic Acid Metabolites Maintain Mechanical and Thermal Hypersensitivity during Sub-Chronic Inflammatory Pain. Biochem. Pharmacol. 2022, 198, 114953. 10.1016/j.bcp.2022.114953. [DOI] [PubMed] [Google Scholar]
Ruparel S.; Green D.; Chen P.; Hargreaves K. M. The Cytochrome P450 Inhibitor, Ketoconazole, Inhibits Oxidized Linoleic Acid Metabolite-Mediated Peripheral Inflammatory Pain. Mol. Pain 2012, 8, 1744–8069–8-73. 10.1186/1744-8069-8-73. [DOI] [PMC free article] [PubMed] [Google Scholar]
Green D. P.; Ruparel S.; Roman L.; Henry M. A.; Hargreaves K. M. Role of Endogenous TRPV1 Agonists in a Postburn Pain Model of Partial-Thickness Injury. Pain 2013, 154 (11), 2512–2520. 10.1016/j.pain.2013.07.040. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hargreaves K. M.; Ruparel S. Role of Oxidized Lipids and TRP Channels in Orofacial Pain and Inflammation. J. Dent. Res. 2016, 95 (10), 1117–1123. 10.1177/0022034516653751. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sisignano M.; Angioni C.; Park C.-K.; Meyer Dos Santos S.; Jordan H.; Kuzikov M.; Liu D.; Zinn S.; Hohman S. W.; Schreiber Y.; Zimmer B.; Schmidt M.; Lu R.; Suo J.; Zhang D.-D.; Schäfer S. M. G.; Hofmann M.; Yekkirala A. S.; de Bruin N.; Parnham M. J.; Woolf C. J.; Ji R.-R.; Scholich K.; Geisslinger G. Targeting CYP2J to Reduce Paclitaxel-Induced Peripheral Neuropathic Pain. Proc. Natl. Acad. Sci. U. S. A. 2016, 113 (44), 12544–12549. 10.1073/pnas.1613246113. [DOI] [PMC free article] [PubMed] [Google Scholar]
Osthues T.; Zimmer B.; Rimola V.; Klann K.; Schilling K.; Mathoor P.; Angioni C.; Weigert A.; Geisslinger G.; Münch C.; Scholich K.; Sisignano M. The Lipid Receptor G2A (GPR132) Mediates Macrophage Migration in Nerve Injury-Induced Neuropathic Pain. Cells 2020, 9 (7), 1740. 10.3390/cells9071740. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hellström F.; Gouveia-Figueira S.; Nording M. L.; Björklund M.; Fowler C. J. Association between Plasma Concentrations of Linoleic Acid-Derived Oxylipins and the Perceived Pain Scores in an Exploratory Study in Women with Chronic Neck Pain. BMC Musculoskelet. Disord. 2016, 17 (1), 103. 10.1186/s12891-016-0951-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sisignano M.; Angioni C.; Ferreiros N.; Schuh C.-D.; Suo J.; Schreiber Y.; Dawes J. M.; Antunes-Martins A.; Bennett D. L. H.; McMahon S. B.; Geisslinger G.; Scholich K. Synthesis of Lipid Mediators during UVB-Induced Inflammatory Hyperalgesia in Rats and Mice. PLoS One 2013, 8 (12), e81228 10.1371/journal.pone.0081228. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sisignano M.; Steinhilber D.; Parnham M. J.; Geisslinger G. Exploring CYP2J2: Lipid Mediators, Inhibitors and Therapeutic Implications. Drug Discovery Today 2020, 25 (9), 1744–1753. 10.1016/j.drudis.2020.07.002. [DOI] [PubMed] [Google Scholar]
Jensen J. R.; Pitcher M. H.; Yuan Z. X.; Ramsden C. E.; Domenichiello A. F. Concentrations of Oxidized Linoleic Acid Derived Lipid Mediators in the Amygdala and Periaqueductal Grey Are Reduced in a Mouse Model of Chronic Inflammatory Pain. Prostaglandins Leukot. Essent. Fatty Acids 2018, 135, 128–136. 10.1016/j.plefa.2018.07.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
Doolen S.; Keyes G. S.; Ramsden C. E. Hydroxy-Epoxide and Keto-Epoxide Derivatives of Linoleic Acid Activate Trigeminal Neurons. Neurobiol. Pain 2020, 7, 100046. 10.1016/j.ynpai.2020.100046. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yu Z.; Schneider C.; Boeglin W. E.; Brash A. R. Human and Mouse ELOX3 Have Distinct Substrate Specificities: Implications for Their Linkage with Lipoxygenases in Skin. Arch. Biochem. Biophys. 2006, 455 (2), 188–196. 10.1016/j.abb.2006.09.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wheeler J. J.; Domenichiello A. F.; Jensen J. R.; Keyes G. S.; Maiden K. M.; Davis J. M.; Ramsden C. E.; Mishra S. K. Endogenous Derivatives of Linoleic Acid and Their Stable Analogues Are Potential Pain Mediators. JID Innov. 2023, 3, 100177. 10.1016/j.xjidi.2022.100177. [DOI] [PMC free article] [PubMed] [Google Scholar]
Boyd J. T.; LoCoco P. M.; Furr A. R.; Bendele M. R.; Tram M.; Li Q.; Chang F.-M.; Colley M. E.; Samenuk G. M.; Arris D. A.; Locke E. E.; Bach S. B. H.; Tobon A.; Ruparel S. B.; Hargreaves K. M. Elevated Dietary ω-6 Polyunsaturated Fatty Acids Induce Reversible Peripheral Nerve Dysfunction That Exacerbates Comorbid Pain Conditions. Nat. Metab. 2021, 3 (6), 762–773. 10.1038/s42255-021-00410-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
Birkic N.; Azar T.; Maddipati K. R.; Minic Z.; Reynolds C. A. Excessive Dietary Linoleic Acid Promotes Plasma Accumulation of Pronociceptive Fatty Acyl Lipid Mediators. Sci. Rep. 2022, 12 (1), 17832. 10.1038/s41598-022-21823-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
Taha A. Y.; Cheon Y.; Faurot K. F.; MacIntosh B.; Majchrzak-Hong S. F.; Mann J. D.; Hibbeln J. R.; Ringel A.; Ramsden C. E. Dietary Omega-6 Fatty Acid Lowering Increases Bioavailability of Omega-3 Polyunsaturated Fatty Acids in Human Plasma Lipid Pools. Prostaglandins Leukot. Essent. Fatty Acids 2014, 90 (5), 151–157. 10.1016/j.plefa.2014.02.003. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sculptoreanu A.; Kullmann F. A.; Artim D. E.; Bazley F. A.; Schopfer F.; Woodcock S.; Freeman B. A.; de Groat W. C. Nitro-Oleic Acid Inhibits Firing and Activates TRPV1- and TRPA1-Mediated Inward Currents in Dorsal Root Ganglion Neurons from Adult Male Rats. J. Pharmacol. Exp. Ther. 2010, 333, 883–895. 10.1124/jpet.109.163154. [DOI] [PMC free article] [PubMed] [Google Scholar]
Beckel J. M.; de Groat W. C. The Effect of the Electrophilic Fatty Acid Nitro-Oleic Acid on TRP Channel Function in Sensory Neurons. Nitric Oxide Biol. Chem. 2018, 78, 154 10.1016/j.niox.2018.03.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
Gaudet R. A Primer on Ankyrin Repeat Function in TRP Channels and Beyond. Mol. Biosyst. 2008, 4 (5), 372–379. 10.1039/b801481g. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hinman A.; Chuang H.; Bautista D. M.; Julius D. TRP Channel Activation by Reversible Covalent Modification. Proc. Natl. Acad. Sci. U. S. A. 2006, 103 (51), 19564–19568. 10.1073/pnas.0609598103. [DOI] [PMC free article] [PubMed] [Google Scholar]
Mori Y.; Takahashi N.; Kurokawa T.; Kiyonaka S. TRP Channels in Oxygen Physiology: Distinctive Functional Properties and Roles of TRPA1 in O₂ Sensing. Proc. Jpn. Acad. Ser. B Phys. Biol. Sci. 2017, 93 (7), 464–482. 10.2183/pjab.93.028. [DOI] [PMC free article] [PubMed] [Google Scholar]
Miller C. C.; Ziboh V. A. Induction of Epidermal Hyperproliferation by Topical n-3 Polyunsaturated Fatty Acids on Guinea Pig Skin Linked to Decreased Levels of 13-Hydroxyoctadecadienoic Acid (13-Hode). J. Invest. Dermatol. 1990, 94 (3), 353–358. 10.1111/1523-1747.ep12874482. [DOI] [PubMed] [Google Scholar]
Hampel J. K. A.; Brownrigg L. M.; Vignarajah D.; Croft K. D.; Dharmarajan A. M.; Bentel J. M.; Puddey I. B.; Yeap B. B. Differential Modulation of Cell Cycle, Apoptosis and PPARγ2 Gene Expression by PPARγ Agonists Ciglitazone and 9-Hydroxyoctadecadienoic Acid in Monocytic Cells. Prostaglandins Leukot. Essent. Fatty Acids 2006, 74 (5), 283–293. 10.1016/j.plefa.2006.03.002. [DOI] [PubMed] [Google Scholar]
Almeida M.; Ambrogini E.; Han L.; Manolagas S. C.; Jilka R. L. Increased Lipid Oxidation Causes Oxidative Stress, Increased Peroxisome Proliferator-Activated Receptor-γ Expression, and Diminished Pro-Osteogenic Wnt Signaling in the Skeleton. J. Biol. Chem. 2009, 284 (40), 27438–27448. 10.1074/jbc.M109.023572. [DOI] [PMC free article] [PubMed] [Google Scholar]
Glasgow W. C.; Eling T. E. Structure-Activity Relationship for Potentiation of EGF-Dependent Mitogenesis by Oxygenated Metabolites of Linoleic-Acid. Arch. Biochem. Biophys. 1994, 311 (2), 286–292. 10.1006/abbi.1994.1239. [DOI] [PubMed] [Google Scholar]
Honn K. V.; Nelson K. K.; Renaud C.; Bazaz R.; Diglio C. A.; Timar J. Fatty Acid Modulation of Tumor Cell Adhesion to Microvessel Endothelium and Experimental Metastasis. Prostaglandins 1992, 44 (5), 413–429. 10.1016/0090-6980(92)90137-I. [DOI] [PubMed] [Google Scholar]
Tavakoli Yaraki M.; Karami Tehrani F. Apoptosis Induced by 13-S-Hydroxyoctadecadienoic Acid in the Breast Cancer Cell Lines, MCF-7 and MDA-MB-231. Iran. J. Basic Med. Sci. 2013, 16 (4), 653–659. [PMC free article] [PubMed] [Google Scholar]
Felding-Habermann B.; Cheresh D. A. Vitronectin and Its Receptors. Curr. Opin. Cell Biol. 1993, 5 (5), 864–868. 10.1016/0955-0674(93)90036-P. [DOI] [PubMed] [Google Scholar]
Buchanan M. R.; Bertomeu M. C.; Brister S. J.; Haas T. A. 13-Hydroxyoctadecadienoic Acid (13-HODE) Metabolism and Endothelial Cell Adhesion Molecule Expression: Effect on Platelet Vessel Wall Adhesion. Wien. Klin. Wochenschr. 1991, 103, 416–421. [PubMed] [Google Scholar]
Buchanan M. R.; Horsewood P.; Brister S. J. Regulation of Endothelial Cell and Platelet Receptor-Ligand Binding by the 12- and 15-Lipoxygenase Monohydroxides, 12-, 15-HETE and 13-HODE. Prostaglandins Leukot. Essent. Fatty Acids 1998, 58 (5), 339–346. 10.1016/S0952-3278(98)90069-2. [DOI] [PubMed] [Google Scholar]
Frömel T.; Jungblut B.; Hu J.; Trouvain C.; Barbosa-Sicard E.; Popp R.; Liebner S.; Dimmeler S.; Hammock B. D.; Fleming I. Soluble Epoxide Hydrolase Regulates Hematopoietic Progenitor Cell Function via Generation of Fatty Acid Diols. Proc. Natl. Acad. Sci. U. S. A. 2012, 109 (25), 9995–10000. 10.1073/pnas.1206493109. [DOI] [PMC free article] [PubMed] [Google Scholar]
Naeem Z.; Zukunft S.; Günther S.; Liebner S.; Weigert A.; Hammock B. D.; Frömel T.; Fleming I. Role of the Soluble Epoxide Hydrolase in the Hair Follicle Stem Cell Homeostasis and Hair Growth. Pflüg. Arch. - Eur. J. Physiol. 2022, 474 (9), 1021–1035. 10.1007/s00424-022-02709-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
Wang W.; Yang J.; Edin M. L.; Wang Y.; Luo Y.; Wan D.; Yang H.; Song C.-Q.; Xue W.; Sanidad K. Z.; Song M.; Bisbee H. A.; Bradbury J. A.; Nan G.; Zhang J.; Shih P. B.; Lee K. S. S.; Minter L. M.; Kim D.; Xiao H.; Liu J.-Y.; Hammock B. D.; Zeldin D. C.; Zhang G. Targeted Metabolomics Identifies the Cytochrome P450 Monooxygenase Eicosanoid Pathway as a Novel Therapeutic Target of Colon Tumorigenesis. Cancer Res. 2019, 79 (8), 1822–1830. 10.1158/0008-5472.CAN-18-3221. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kong C.; Yan X.; Zhu Y.; Zhu H.; Luo Y.; Liu P.; Ferrandon S.; Kalady M. F.; Gao R.; He J.; Yin F.; Qu X.; Zheng J.; Gao Y.; Wei Q.; Ma Y.; Liu J.-Y.; Qin H. Fusobacterium nucleatum Promotes the Development of Colorectal Cancer by Activating a Cytochrome P450/Epoxyoctadecenoic Acid Axis via TLR4/Keap1/NRF2 Signaling. Cancer Res. 2021, 81 (17), 4485–4498. 10.1158/0008-5472.CAN-21-0453. [DOI] [PubMed] [Google Scholar]
Baird L.; Yamamoto M. The Molecular Mechanisms Regulating the KEAP1-NRF2 Pathway. Mol. Cell. Biol. 2020, 40 (13), 00099–20 10.1128/MCB.00099-20. [DOI] [PMC free article] [PubMed] [Google Scholar]
Fan Z.; Wirth A.-K.; Chen D.; Wruck C. J.; Rauh M.; Buchfelder M.; Savaskan N. Nrf2-Keap1 Pathway Promotes Cell Proliferation and Diminishes Ferroptosis. Oncogenesis 2017, 6 (8), e371 10.1038/oncsis.2017.65. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yang C.-C.; Chang M.-T.; Chang C.-K.; Shyur L.-F. Phytogalactolipid DLGG Inhibits Mouse Melanoma Brain Metastasis through Regulating Oxylipin Activity and Re-Programming Macrophage Polarity in the Tumor Microenvironment. Cancers 2021, 13 (16), 4120. 10.3390/cancers13164120. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kansanen E.; Bonacci G.; Schopfer F. J.; Kuosmanen S. M.; Tong K. I.; Leinonen H.; Woodcock S. R.; Yamamoto M.; Carlberg C.; Ylä-Herttuala S.; Freeman B. A.; Levonen A.-L. Electrophilic Nitro-Fatty Acids Activate NRF2 by a KEAP1 Cysteine 151-Independent Mechanism. J. Biol. Chem. 2011, 286 (16), 14019–14027. 10.1074/jbc.M110.190710. [DOI] [PMC free article] [PubMed] [Google Scholar]
Asan A.; Skoko J. J.; Woodcock C.-S. C.; Wingert B. M.; Woodcock S. R.; Normolle D.; Huang Y.; Stark J. M.; Camacho C. J.; Freeman B. A.; Neumann C. A. Electrophilic Fatty Acids Impair RAD51 Function and Potentiate the Effects of DNA-Damaging Agents on Growth of Triple-Negative Breast Cells. J. Biol. Chem. 2019, 294 (2), 397–404. 10.1074/jbc.AC118.005899. [DOI] [PMC free article] [PubMed] [Google Scholar]
Noguchi E.; Gadaleta M.. Cell Cycle Control Mechanisms and Protocols; Methods in Molecular Biology; Springer Science+Business, Humana: New York, 2014. [Google Scholar]
Shu L.; Wang D.; Saba N. F.; Chen Z. G. A Historic Perspective and Overview of H-Ras Structure, Oncogenicity, and Targeting. Mol. Cancer Ther. 2020, 19 (4), 999–1007. 10.1158/1535-7163.MCT-19-0660. [DOI] [PubMed] [Google Scholar]
Pundlik S. S.; Sahoo S. S.; Barik A.; Jaysingh M. A.; Venkateshvaran A.; Math R. G. H.; Ramanathan A. C-Terminal Cysteines of HRas Control Erk Signaling and 15-Deoxy-Δ12,14-Prostaglandin J₂ (15d-PGJ₂) Mediated Inhibition of Myoblast Differentiation. BioRxiv 2023, 2023.05.20.541575. 10.1101/2023.05.20.541575. [DOI] [Google Scholar]
Renedo M.; Gayarre J.; García-Domínguez C. A.; Pérez-Rodríguez A.; Prieto A.; Cañada F. J.; Rojas J. M.; Pérez-Sala D. Modification and Activation of Ras Proteins by Electrophilic Prostanoids with Different Structure Are Site-Selective. Biochemistry 2007, 46 (22), 6607–6616. 10.1021/bi602389p. [DOI] [PubMed] [Google Scholar]
Kato T.; Yamaguchi Y.; Uyehara T.; Yokoyama T.; Namai T.; Yamanaka S. Self Defensive Substances in Rice Plant against Rice Blast Disease. Tetrahedron Lett. 1983, 24 (43), 4715–4718. 10.1016/S0040-4039(00)86236-X. [DOI] [Google Scholar]
Ozawa T.; Hayakawa M.; Takamura T.; Sugiyama S.; Suzuki K.; Iwata M.; Taki F.; Tomita T. Biosynthesis of Leukotoxin, 9,10-Epoxy-12 Octadecenoate, by Leukocytes in Lung Lavages of Rat after Exposure to Hyperoxia. Biochem. Biophys. Res. Commun. 1986, 134 (3), 1071–1078. 10.1016/0006-291X(86)90360-8. [DOI] [PubMed] [Google Scholar]
Hayakawa M.; Sugiyama S.; Takamura T.; Yokoo K.; Iwata M.; Suzuki K.; Taki F.; Takahashi S.; Ozawa T. Neutrophils Biosynthesize Leukotoxin, 9,10-Epoxy-12-Octadecenoate. Biochem. Biophys. Res. Commun. 1986, 137 (1), 424–430. 10.1016/0006-291X(86)91227-1. [DOI] [PubMed] [Google Scholar]
Jia-ning H.; Taki F.; Sugiyama S.; Asai J.; Izawa Y.; Satake T.; Ozawa T. Neutrophil-Derived Epoxide, 9,10-Epoxy-12-Octadecenoate, Induces Pulmonary Edema. Lung 1988, 166 (1), 327–337. 10.1007/BF02714065. [DOI] [PubMed] [Google Scholar]
Ozawa T.; Sugiyama S.; Hayakawa M.; Taki F.; Hanaki Y. Neutrophil Microsomes Biosynthesize Linoleate Epoxide (9,10-Epoxy-12-Octadecenoate), a Biological Active Substance. Biochem. Biophys. Res. Commun. 1988, 152 (3), 1310–1318. 10.1016/S0006-291X(88)80428-5. [DOI] [PubMed] [Google Scholar]
Ozawa T.; Sugiyama S.; Hayakawa M.; Satake T.; Taki F.; Iwata M.; Taki K. Existence of Leukotoxin 9,10-Epoxy-12-Octadecenoate in Lung Lavages from Rats Breathing Pure Oxygen and from Patients with the Adult Respiratory Distress Syndrome. Am. Rev. Respir. Dis. 1988, 137 (3), 535–540. 10.1164/ajrccm/137.3.535. [DOI] [PubMed] [Google Scholar]
Ozawa T.; Nishikimi M.; Sugiyama S.; Taki F.; Hayakawa M.; Shionoya H. Cytotoxic Activity of Leukotoxin, a Neutrophil-Derived Fatty Acid Epoxide, on Cultured Human Cells. Biochem. Int. 1988, 16, 369–373. [PubMed] [Google Scholar]
Sakai T.; Ishizaki T.; Ohnishi T.; Sasaki F.; Ameshima S.; Nakai T.; Miyabo S.; Matsukawa S.; Hayakawa M.; Ozawa T. Leukotoxin, 9,10-Epoxy-12-Octadecenoate Inhibits Mitochondrial Respiration of Isolated Perfused Rat Lung. Am. J. Physiol.-Lung Cell. Mol. Physiol. 1995, 269 (3), L326–L331. 10.1152/ajplung.1995.269.3.L326. [DOI] [PubMed] [Google Scholar]
Moghaddam M. F.; Grant D. F.; Cheek J. M.; Greene J. F.; Williamson K. C.; Hammock B. D. Bioactivation of Leukotoxins to Their Toxic Diols by Epoxide Hydrolase. Nat. Med. 1997, 3 (5), 562–566. 10.1038/nm0597-562. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zheng J.; Plopper C. G.; Lakritz J.; Storms D. H.; Hammock B. D. Leukotoxin-Diol. Am. J. Respir. Cell Mol. Biol. 2001, 25 (4), 434–438. 10.1165/ajrcmb.25.4.4104. [DOI] [PubMed] [Google Scholar]
Nummela A.; Laaksonen L.; Scheinin A.; Kaisti K.; Vahlberg T.; Neuvonen M.; Valli K.; Revonsuo A.; Perola M.; Niemi M.; Scheinin H.; Laitio T. Circulating Oxylipin and Bile Acid Profiles of Dexmedetomidine, Propofol, Sevoflurane, and S-Ketamine: A Randomised Controlled Trial Using Tandem Mass Spectrometry. BJA Open 2022, 4, 100114. 10.1016/j.bjao.2022.100114. [DOI] [PMC free article] [PubMed] [Google Scholar]
McReynolds C.; Hammock B.; Morisseau C. Regulatory Lipid Vicinal Diols Counteract the Biological Activity of Epoxy Fatty Acids and Can Act as Biomarkers and Mechanisms for Disease Progression. Pharmacol. Ther. 2023, 248, 108454. 10.1016/j.pharmthera.2023.108454. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sisemore M. F.; Zheng J.; Yang J. C.; Thompson D. A.; Plopper C. G.; Cortopassi G. A.; Hammock B. D. Cellular Characterization of Leukotoxin Diol-Induced Mitochondrial Dysfunction. Arch. Biochem. Biophys. 2001, 392 (1), 32–37. 10.1006/abbi.2001.2434. [DOI] [PubMed] [Google Scholar]
Samokhvalov V.; Jamieson K. L.; Darwesh A. M.; Keshavarz-Bahaghighat H.; Lee T. Y. T.; Edin M.; Lih F.; Zeldin D. C.; Seubert J. M. Deficiency of Soluble Epoxide Hydrolase Protects Cardiac Function Impaired by LPS-Induced Acute Inflammation. Front. Pharmacol. 2019, 9, 1572. 10.3389/fphar.2018.01572. [DOI] [PMC free article] [PubMed] [Google Scholar]
Moran J. H.; Weise R.; Schnellmann R. G.; Freeman J. P.; Grant D. F. Cytotoxicity of Linoleic Acid Diols to Renal Proximal Tubular Cells. Toxicol. Appl. Pharmacol. 1997, 146 (1), 53–59. 10.1006/taap.1997.8197. [DOI] [PubMed] [Google Scholar]
Warner D. R.; Liu H.; Miller M. E.; Ramsden C. E.; Gao B.; Feldstein A. E.; Schuster S.; McClain C. J.; Kirpich I. A. Dietary Linoleic Acid and Its Oxidized Metabolites Exacerbate Liver Injury Caused by Ethanol via Induction of Hepatic Proinflammatory Response in Mice. Am. J. Pathol. 2017, 187 (10), 2232–2245. 10.1016/j.ajpath.2017.06.008. [DOI] [PMC free article] [PubMed] [Google Scholar]
Moran J. H.; Nowak G.; Grant D. F. Analysis of the Toxic Effects of Linoleic Acid, 12,13-cis-Epoxyoctadecenoic Acid, and 12,13-Dihydroxyoctadecenoic Acid in Rabbit Renal Cortical Mitochondria. Toxicol. Appl. Pharmacol. 2001, 172 (2), 150–161. 10.1006/taap.2001.9149. [DOI] [PubMed] [Google Scholar]
Nowak G.; Grant D. F.; Moran J. H. Linoleic Acid Epoxide Promotes the Maintenance of Mitochondrial Function and Active Na+ Transport Following Hypoxia. Toxicol. Lett. 2004, 147 (2), 161–175. 10.1016/j.toxlet.2003.11.002. [DOI] [PubMed] [Google Scholar]
Sosnowski D. K.; Jamieson K. L.; Gruzdev A.; Li Y.; Valencia R.; Yousef A.; Kassiri Z.; Zeldin D. C.; Seubert J. M. Cardiomyocyte-Specific Disruption of Soluble Epoxide Hydrolase Limits Inflammation to Preserve Cardiac Function. Am. J. Physiol.-Heart Circ. Physiol. 2022, 323 (4), H670–H687. 10.1152/ajpheart.00217.2022. [DOI] [PMC free article] [PubMed] [Google Scholar]
Xue H.-M.; Sun W.-T.; Chen H.-X.; He G.-W.; Yang Q. Targeting IRE1α-JNK-c-Jun/AP-1-SEH Signaling Pathway Improves Myocardial and Coronary Endothelial Function Following Global Myocardial Ischemia/Reperfusion. Int. J. Med. Sci. 2022, 19 (9), 1460–1472. 10.7150/ijms.74533. [DOI] [PMC free article] [PubMed] [Google Scholar]
Koenitzer J. R.; Bonacci G.; Woodcock S. R.; Chen C.-S.; Cantu-Medellin N.; Kelley E. E.; Schopfer F. J. Fatty Acid Nitroalkenes Induce Resistance to Ischemic Cardiac Injury by Modulating Mitochondrial Respiration at Complex II. Redox Biol. 2016, 8, 1–10. 10.1016/j.redox.2015.11.002. [DOI] [PMC free article] [PubMed] [Google Scholar]
Nadtochiy S. M.; Zhu Q.; Urciuoli W.; Rafikov R.; Black S. M.; Brookes P. S. Nitroalkenes Confer Acute Cardioprotection via Adenine Nucleotide Translocase 1. J. Biol. Chem. 2012, 287 (5), 3573–3580. 10.1074/jbc.M111.298406. [DOI] [PMC free article] [PubMed] [Google Scholar]
Brand M. D.; Pakay J. L.; Ocloo A.; Kokoszka J.; Wallace D. C.; Brookes P. S.; Cornwall E. J. The Basal Proton Conductance of Mitochondria Depends on Adenine Nucleotide Translocase Content. Biochem. J. 2005, 392, 353–362. 10.1042/BJ20050890. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bauer M. K. A.; Schubert A.; Rocks O.; Grimm S. Adenine Nucleotide Translocase-1, a Component of the Permeability Transition Pore, Can Dominantly Induce Apoptosis. J. Cell Biol. 1999, 147 (7), 1493–1502. 10.1083/jcb.147.7.1493. [DOI] [PMC free article] [PubMed] [Google Scholar]
Liu Y.; Chen X. J. Adenine Nucleotide Translocase, Mitochondrial Stress, and Degenerative Cell Death. Oxid. Med. Cell. Longev. 2013, 2013, 146860. 10.1155/2013/146860. [DOI] [PMC free article] [PubMed] [Google Scholar]
Pan S.; Wang N.; Bisetto S.; Yi B.; Sheu S.-S. Downregulation of Adenine Nucleotide Translocator 1 Exacerbates Tumor Necrosis Factor-α-Mediated Cardiac Inflammatory Responses. Am. J. Physiol.-Heart Circ. Physiol. 2015, 308 (1), H39–H48. 10.1152/ajpheart.00330.2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
Berger A.; Gershwin M. E.; German J. B. Effects of Various Dietary Fats on Cardiolipin Acyl Composition during Ontogeny of Mice. Lipids 1992, 27 (8), 605–612. 10.1007/BF02536118. [DOI] [PubMed] [Google Scholar]
Angelotti A.; Snoke D. B.; Ormiston K.; Cole R. M.; Borkowski K.; Newman J. W.; Orchard T. S.; Belury M. A. Potential Cardioprotective Effects and Lipid Mediator Differences in Long-Chain Omega-3 Polyunsaturated Fatty Acid Supplemented Mice Given Chemotherapy. Metabolites 2022, 12 (9), 782. 10.3390/metabo12090782. [DOI] [PMC free article] [PubMed] [Google Scholar]
Mabalirajan U.; Rehman R.; Ahmad T.; Kumar S.; Singh S.; Leishangthem G. D.; Aich J.; Kumar M.; Khanna K.; Singh V. P.; Dinda A. K.; Biswal S.; Agrawal A.; Ghosh B. Linoleic Acid Metabolite Drives Severe Asthma by Causing Airway Epithelial Injury. Sci. Rep. 2013, 3 (1), 1349. 10.1038/srep01349. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kim Y.-I.; Hirai S.; Takahashi H.; Goto T.; Ohyane C.; Tsugane T.; Konishi C.; Fujii T.; Inai S.; Iijima Y.; Aoki K.; Shibata D.; Takahashi N.; Kawada T. 9-Oxo-10(E),12(E)-Octadecadienoic Acid Derived from Tomato Is a Potent PPAR α Agonist to Decrease Triglyceride Accumulation in Mouse Primary Hepatocytes. Mol. Nutr. Food Res. 2011, 55 (4), 585–593. 10.1002/mnfr.201000264. [DOI] [PubMed] [Google Scholar]
Kim M.; Furuzono T.; Yamakuni K.; Li Y.; Kim Y.; Takahashi H.; Ohue-Kitano R.; Jheng H.; Takahashi N.; Kano Y.; Yu R.; Kishino S.; Ogawa J.; Uchida K.; Yamazaki J.; Tominaga M.; Kawada T.; Goto T. 10-oxo-12(Z)-octadecenoic Acid, a Linoleic Acid Metabolite Produced by Gut Lactic Acid Bacteria, Enhances Energy Metabolism by Activation of TRPV1. FASEB J. 2017, 31 (11), 5036–5048. 10.1096/fj.201700151R. [DOI] [PubMed] [Google Scholar]
Murthy S.; Born E.; Mathur S.; Field F. J. 13-Hydroxy Octadecadienoic Acid (13-HODE) Inhibits Triacylglycerol-Rich Lipoprotein Secretion by CaCo-2 Cells. J. Lipid Res. 1998, 39 (6), 1254–1262. 10.1016/S0022-2275(20)32550-5. [DOI] [PubMed] [Google Scholar]
Kwon S. Y.; Massey K.; Watson M. A.; Hussain T.; Volpe G.; Buckley C. D.; Nicolaou A.; Badenhorst P. Oxidised Metabolites of the Omega-6 Fatty Acid Linoleic Acid Activate DFOXO. Life Sci. Alliance 2020, 3 (2), e201900356 10.26508/lsa.201900356. [DOI] [PMC free article] [PubMed] [Google Scholar]
Furuhashi M.; Saitoh S.; Shimamoto K.; Miura T. Fatty Acid-Binding Protein 4 (FABP4): Pathophysiological Insights and Potent Clinical Biomarker of Metabolic and Cardiovascular Diseases. Clin. Med. Insights Cardiol. 2014, 8s3, CMC.S17067. 10.4137/CMC.S17067. [DOI] [PMC free article] [PubMed] [Google Scholar]
Pinckard K. M.; Stanford K. I. The Heartwarming Effect of Brown Adipose Tissue. Mol. Pharmacol. 2022, 102 (1), 39–50. 10.1124/molpharm.121.000328. [DOI] [PMC free article] [PubMed] [Google Scholar]
Peres Valgas da Silva C.; Hernández-Saavedra D.; White J. D.; Stanford K. I. Cold and Exercise: Therapeutic Tools to Activate Brown Adipose Tissue and Combat Obesity. Biology 2019, 8 (1), 9. 10.3390/biology8010009. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hernández-Saavedra D.; Stanford K. I. The Regulation of Lipokines by Environmental Factors. Nutrients 2019, 11 (10), 2422. 10.3390/nu11102422. [DOI] [PMC free article] [PubMed] [Google Scholar]
Huang L.; Tepaamorndech S.; Kirschke C. P.; Newman J. W.; Keyes W. R.; Pedersen T. L.; Dumnil J. Aberrant Fatty Acid Metabolism in Skeletal Muscle Contributes to Insulin Resistance in Zinc Transporter 7 (Znt7)-Knockout Mice. J. Biol. Chem. 2018, 293 (20), 7549–7563. 10.1074/jbc.M117.817692. [DOI] [PMC free article] [PubMed] [Google Scholar]
Dreyfuss J. M.; Djordjilović V.; Pan H.; Bussberg V.; MacDonald A. M.; Narain N. R.; Kiebish M. A.; Blüher M.; Tseng Y.-H.; Lynes M. D. ScreenDMT Reveals DiHOMEs Are Replicably Inversely Associated with BMI and Stimulate Adipocyte Calcium Influx. Commun. Biol. 2024, 7 (1), 996. 10.1038/s42003-024-06646-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kim Y.; Hirai S.; Goto T.; Ohyane C.; Takahashi H.; Tsugane T.; Konishi C.; Fujii T.; Inai S.; Iijima Y.; Aoki K.; Shibata D.; Takahashi N.; Kawada T. Potent PPARα Activator Derived from Tomato Juice, 13-Oxo-9,11-Octadecadienoic Acid, Decreases Plasma and Hepatic Triglyceride in Obese Diabetic Mice. PLoS One 2012, 7 (2), e31317 10.1371/journal.pone.0031317. [DOI] [PMC free article] [PubMed] [Google Scholar]
Takahashi H.; Hara H.; Goto T.; Kamakari K.; Wataru N.; Mohri S.; Takahashi N.; Suzuki H.; Shibata D.; Kawada T. 13-Oxo-9(Z),11(E),15(Z)-Octadecatrienoic Acid Activates Peroxisome Proliferator-Activated Receptor γ in Adipocytes. Lipids 2015, 50 (1), 3–12. 10.1007/s11745-014-3972-x. [DOI] [PubMed] [Google Scholar]
Nieman D. C.; Shanely R. A.; Luo B.; Meaney M. P.; Dew D. A.; Pappan K. L. Metabolomics Approach to Assessing Plasma 13- and 9-Hydroxy-Octadecadienoic Acid and Linoleic Acid Metabolite Responses to 75-Km Cycling. Am. J. Physiol.-Regul. Integr. Comp. Physiol. 2014, 307 (1), R68–R74. 10.1152/ajpregu.00092.2014. [DOI] [PubMed] [Google Scholar]
Nieman D. C.; Meaney M. P.; John C. S.; Knagge K. J.; Chen H. 9- and 13-Hydroxy-Octadecadienoic Acids (9 + 13 HODE) Are Inversely Related to Granulocyte Colony Stimulating Factor and IL-6 in Runners after 2h Running. Brain. Behav. Immun. 2016, 56, 246–252. 10.1016/j.bbi.2016.03.020. [DOI] [PubMed] [Google Scholar]
Mani S. K.; Reyna A. M.; Alejandro M. A.; Crowley J.; Markaverich B. M. Disruption of Male Sexual Behavior in Rats by Tetrahydrofurandiols (THF-Diols). Steroids 2005, 70 (11), 750–754. 10.1016/j.steroids.2005.04.004. [DOI] [PubMed] [Google Scholar]
Markaverich B. M.; Crowley J. R.; Alejandro M. A.; Shoulars K.; Casajuna N.; Mani S.; Reyna A.; Sharp J. Leukotoxin Diols from Ground Corncob Bedding Disrupt Estrous Cyclicity in Rats and Stimulate MCF-7 Breast Cancer Cell Proliferation. Environ. Health Perspect. 2005, 113 (12), 1698–1704. 10.1289/ehp.8231. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ishizaki T.; Shigemori K.; Yamamura Y.; Matsukawa S.; Nakai T.; Miyabo S.; Hayakawa M.; Ozawa T.; Voelkel N. F. Increased Nitric Oxide Biosynthesis in Leukotoxin, 9,10-Epoxy-12-Octadecenoate Injured Lung. Biochem. Biophys. Res. Commun. 1995, 210 (1), 133–137. 10.1006/bbrc.1995.1637. [DOI] [PubMed] [Google Scholar]
Hull E. M.; Lumley L. A.; Matuszewich L.; Dominguez J.; Moses J.; Lorrain D. S. The Roles of Nitric Oxide in Sexual Function of Male Rats. Neuropharmacology 1994, 33 (11), 1499–1504. 10.1016/0028-3908(94)90054-X. [DOI] [PubMed] [Google Scholar]
Okamura S.; Ameshima S.; Demura Y.; Ishizaki T.; Matsukawa S.; Miyamori I. Leukotoxin-Activated Human Pulmonary Artery Endothelial Cell Produces Nitric Oxide and Superoxide Anion. Pulm. Pharmacol. Ther. 2002, 15 (1), 25–33. 10.1006/pupt.2001.0322. [DOI] [PubMed] [Google Scholar]
Markaverich B. M.; Alejandro M.; Thompson T.; Mani S.; Reyna A.; Portillo W.; Sharp J.; Turk J.; Crowley J. R. Tetrahydrofurandiols (THF-Diols), Leukotoxindiols (LTX-Diols), and Endocrine Disruption in Rats. Environ. Health Perspect. 2007, 115 (5), 702–708. 10.1289/ehp.9311. [DOI] [PMC free article] [PubMed] [Google Scholar]
Siegfried M. R.; Aoki N.; Lefer A. M.; Elisseou E. M.; Zipkin R. E. Direct Cardiovascular Actions of Two Metabolites of Linoleic Acid. Life Sci. 1990, 46 (6), 427–433. 10.1016/0024-3205(90)90086-7. [DOI] [PubMed] [Google Scholar]
Fukushima A.; Hayakawa M.; Sugiyama S.; Ajioka M.; Ito T.; Satake T.; Ozawa T. Cardiovascular Effects of Leukotoxin (9,10-Epoxy-12-Octadecenoate) and Free Fatty Acids in Dogs. Cardiovasc. Res. 1988, 22, 213–218. 10.1093/cvr/22.3.213. [DOI] [PubMed] [Google Scholar]
Matsuda H.; Miyatake K.; Dahlén S. E. Pharmacodynamics of 15(S)-Hydroperoxyeicosatetraenoic (15-HPETE) and 15(S)-Hydroxyeicosatetraenoic Acid (15-HETE) in Isolated Arteries from Guinea Pig, Rabbit, Rat and Human. J. Pharmacol. Exp. Ther. 1995, 273 (3), 1182–1189. [PubMed] [Google Scholar]
De Meyer G. R. Y.; Bult H.; Verbeuren T. J.; Herman A. G. The Role of Endothelial Cells in the Relaxations Induced by 13-Hydroxy- and 13-Hydroperoxylinoleic Acid in Canine Arteries. Br. J. Pharmacol. 1992, 107 (2), 597–603. 10.1111/j.1476-5381.1992.tb12789.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
Edin M. L.; Wang Z.; Bradbury J. A.; Graves J. P.; Lih F. B.; DeGraff L. M.; Foley J. F.; Torphy R.; Ronnekleiv O. K.; Tomer K. B.; Lee C. R.; Zeldin D. C. Endothelial Expression of Human Cytochrome P450 Epoxygenase CYP2C8 Increases Susceptibility to Ischemia-Reperfusion Injury in Isolated Mouse Heart. FASEB J. 2011, 25 (10), 3436–3447. 10.1096/fj.11-188300. [DOI] [PMC free article] [PubMed] [Google Scholar]
Tloti M. A.; Moon D. G.; Weston L. K.; Kaplan J. E. Effect of 13-Hydroxyoctadeca-9,11-Dienoic Acid (13-HODE) on Thrombin Induced Platelet Adherence to Endothelial Cells in Vitro. Thromb. Res. 1991, 62 (4), 305–317. 10.1016/0049-3848(91)90151-L. [DOI] [PubMed] [Google Scholar]
Coene M.-C.; Bult H.; Claeys M.; Laekeman G. M.; Herman A. G. Inhibition of Rabbit Platelet Activation by Lipoxygenase Products of Arachidonic and Linoleic Acid. Thromb. Res. 1986, 42 (2), 205–214. 10.1016/0049-3848(86)90296-3. [DOI] [PubMed] [Google Scholar]
Lê Q. H.; El Alaoui M.; Véricel E.; Ségrestin B.; Soulère L.; Guichardant M.; Lagarde M.; Moulin P.; Calzada C. Glycoxidized HDL, HDL Enriched With Oxidized Phospholipids and HDL From Diabetic Patients Inhibit Platelet Function. J. Clin. Endocrinol. Metab. 2015, 100 (5), 2006–2014. 10.1210/jc.2014-4214. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zhang J.; Villacorta L.; Chang L.; Fan Z.; Hamblin M.; Zhu T.; Chen C. S.; Cole M. P.; Schopfer F. J.; Deng C. X.; Garcia-Barrio M. T.; Feng Y.-H.; Freeman B. A.; Chen Y. E. Nitro-Oleic Acid Inhibits Angiotensin II-Induced Hypertension. Circ. Res. 2010, 107 (4), 540–548. 10.1161/CIRCRESAHA.110.218404. [DOI] [PMC free article] [PubMed] [Google Scholar]
Khoo N. K. H.; Rudolph V.; Cole M. P.; Golin-Bisello F.; Schopfer F. J.; Woodcock S. R.; Batthyany C.; Freeman B. A. Activation Of Vascular Endothelial Nitric Oxide Synthase And Heme Oxygenase-1 Expression By Electrophilic Nitro-Fatty Acids. Free Radic. Biol. Med. 2010, 48 (2), 230. 10.1016/j.freeradbiomed.2009.10.046. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yoshida Y.; Yoshikawa A.; Kinumi T.; Ogawa Y.; Saito Y.; Ohara K.; Yamamoto H.; Imai Y.; Niki E. Hydroxyoctadecadienoic Acid and Oxidatively Modified Peroxiredoxins in the Blood of Alzheimer’s Disease Patients and Their Potential as Biomarkers. Neurobiol. Aging 2009, 30 (2), 174–185. 10.1016/j.neurobiolaging.2007.06.012. [DOI] [PubMed] [Google Scholar]
Kurano M.; Saito Y.; Uranbileg B.; Saigusa D.; Kano K.; Aoki J.; Yatomi Y. Modulations of Bioactive Lipids and Their Receptors in Postmortem Alzheimer’s Disease Brains. Front. Aging Neurosci. 2022, 14, 1066578. 10.3389/fnagi.2022.1066578. [DOI] [PMC free article] [PubMed] [Google Scholar]
Borkowski K.; Pedersen T. L.; Seyfried N. T.; Lah J. J.; Levey A. I.; Hales C. M.; Dammer E. B.; Blach C.; Louie G.; Kaddurah-Daouk R.; Newman J. W. Alzheimer’s Disease Metabolomics Consortium. Association of Plasma and CSF Cytochrome P450, Soluble Epoxide Hydrolase, and Ethanolamide Metabolism with Alzheimer’s Disease. Alzheimers Res. Ther. 2021, 13 (1), 149. 10.1186/s13195-021-00893-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
Morris J. K.; Piccolo B. D.; John C. S.; Green Z. D.; Thyfault J. P.; Adams S. H. Oxylipin Profiling of Alzheimer’s Disease in Nondiabetic and Type 2 Diabetic Elderly. Metabolites 2019, 9 (9), 177. 10.3390/metabo9090177. [DOI] [PMC free article] [PubMed] [Google Scholar]
Shen Q.; Patten K. T.; Valenzuela A.; Lein P. J.; Taha A. Y. Probing Changes in Brain Esterified Oxylipin Concentrations during the Early Stages of Pathogenesis in Alzheimer’s Disease Transgenic Rats. Neurosci. Lett. 2022, 791, 136921. 10.1016/j.neulet.2022.136921. [DOI] [PMC free article] [PubMed] [Google Scholar]
Shinto L. H.; Raber J.; Mishra A.; Roese N.; Silbert L. C. A Review of Oxylipins in Alzheimer’s Disease and Related Dementias (ADRD): Potential Therapeutic Targets for the Modulation of Vascular Tone and Inflammation. Metabolites 2022, 12 (9), 826. 10.3390/metabo12090826. [DOI] [PMC free article] [PubMed] [Google Scholar]
Devassy J. G.; Leng S.; Gabbs M.; Monirujjaman M.; Aukema H. M. Omega-3 Polyunsaturated Fatty Acids and Oxylipins in Neuroinflammation and Management of Alzheimer Disease12. Adv. Nutr. 2016, 7 (5), 905–916. 10.3945/an.116.012187. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hennebelle M.; Zhang Z.; Metherel A. H.; Kitson A. P.; Otoki Y.; Richardson C. E.; Yang J.; Lee K. S. S.; Hammock B. D.; Zhang L.; Bazinet R. P.; Taha A. Y. Linoleic Acid Participates in the Response to Ischemic Brain Injury through Oxidized Metabolites That Regulate Neurotransmission. Sci. Rep. 2017, 7 (1), 4342. 10.1038/s41598-017-02914-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hennebelle M.; Metherel A. H.; Kitson A. P.; Otoki Y.; Yang J.; Lee K. S. S.; Hammock B. D.; Bazinet R. P.; Taha A. Y. Brain Oxylipin Concentrations Following Hypercapnia/Ischemia: Effects of Brain Dissection and Dissection Time. J. Lipid Res. 2019, 60 (3), 671–682. 10.1194/jlr.D084228. [DOI] [PMC free article] [PubMed] [Google Scholar]
Szczuko M.; Kotlęga D.; Palma J.; Zembroń-Łacny A.; Tylutka A.; Gołąb-Janowska M.; Drozd A. Lipoxins, RevD₁ and 9, 13 HODE as the Most Important Derivatives after an Early Incident of Ischemic Stroke. Sci. Rep. 2020, 10 (1), 12849. 10.1038/s41598-020-69831-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
Yu D.; Hennebelle M.; Sahlas D. J.; Ramirez J.; Gao F.; Masellis M.; Cogo-Moreira H.; Swartz R. H.; Herrmann N.; Chan P. C.; Pettersen J. A.; Stuss D. T.; Black S. E.; Taha A. Y.; Swardfager W. Soluble Epoxide Hydrolase-Derived Linoleic Acid Oxylipins in Serum Are Associated with Periventricular White Matter Hyperintensities and Vascular Cognitive Impairment. Transl. Stroke Res. 2019, 10 (5), 522–533. 10.1007/s12975-018-0672-5. [DOI] [PubMed] [Google Scholar]
Shinto L.; Lahna D.; Murchison C. F.; Dodge H.; Hagen K.; David J.; Kaye J.; Quinn J. F.; Wall R.; Silbert L. C. Oxidized Products of Omega-6 and Omega-3 Long Chain Fatty Acids Are Associated with Increased White Matter Hyperintensity and Poorer Executive Function Performance in a Cohort of Cognitively Normal Hypertensive Older Adults. J. Alzheimers Dis. 2020, 74 (1), 65–77. 10.3233/JAD-191197. [DOI] [PMC free article] [PubMed] [Google Scholar]
Borkowski K.; Taha A. Y.; Pedersen T. L.; De Jager P. L.; Bennett D. A.; Arnold M.; Kaddurah-Daouk R.; Newman J. W. Serum Metabolomic Biomarkers of Perceptual Speed in Cognitively Normal and Mildly Impaired Subjects with Fasting State Stratification. Sci. Rep. 2021, 11 (1), 18964. 10.1038/s41598-021-98640-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
Anita N. Z.; Kwan F.; Ryoo S. W.; Major-Orfao C.; Lin W. Z.; Noor S.; Lanctôt K. L.; Herrmann N.; Oh P. I.; Shah B. R.; Gilbert J.; Assal A.; Halperin I. J.; Taha A. Y.; Swardfager W. Cytochrome P450-Soluble Epoxide Hydrolase Derived Linoleic Acid Oxylipins and Cognitive Performance in Type 2 Diabetes. J. Lipid Res. 2023, 64, 100395. 10.1016/j.jlr.2023.100395. [DOI] [PMC free article] [PubMed] [Google Scholar]
Filippi M.; Bar-Or A.; Piehl F.; Preziosa P.; Solari A.; Vukusic S.; Rocca M. A. Multiple Sclerosis. Nat. Rev. Dis. Primer 2018, 4 (1), 1–27. 10.1038/s41572-018-0041-4. [DOI] [PubMed] [Google Scholar]
Villoslada P.; Alonso C.; Agirrezabal I.; Kotelnikova E.; Zubizarreta I.; Pulido-Valdeolivas I.; Saiz A.; Comabella M.; Montalban X.; Villar L.; Alvarez-Cermeño J. C.; Fernández O.; Alvarez-Lafuente R.; Arroyo R.; Castro A. Metabolomic Signatures Associated with Disease Severity in Multiple Sclerosis. Neurol. - Neuroimmunol. Neuroinflammation 2017, 4 (2), e321 10.1212/NXI.0000000000000321. [DOI] [PMC free article] [PubMed] [Google Scholar]
Håkansson I.; Gouveia-Figueira S.; Ernerudh J.; Vrethem M.; Ghafouri N.; Ghafouri B.; Nording M. Oxylipins in Cerebrospinal Fluid in Clinically Isolated Syndrome and Relapsing Remitting Multiple Sclerosis. Prostaglandins Other Lipid Mediat. 2018, 138, 41–47. 10.1016/j.prostaglandins.2018.08.003. [DOI] [PubMed] [Google Scholar]
Shen Q.; Otoki Y.; Sobel R. A.; Nagra R. M.; Taha A. Y. Evidence of Increased Sequestration of Pro-Resolving Lipid Mediators within Brain Esterified Lipid Pools of Multiple Sclerosis Patients. Mult. Scler. Relat. Disord. 2022, 68, 104236. 10.1016/j.msard.2022.104236. [DOI] [PubMed] [Google Scholar]
Shan J.; Hashimoto K. Soluble Epoxide Hydrolase as a Therapeutic Target for Neuropsychiatric Disorders. Int. J. Mol. Sci. 2022, 23 (9), 4951. 10.3390/ijms23094951. [DOI] [PMC free article] [PubMed] [Google Scholar]
Brooks C. J. W.; Harland W. A.; Steel G.; Gilbert J. D. Lipids of Human Atheroma: Isolation of Hydroxyoctadecadienoic Acids from Advanced Aortal Lesions. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1970, 202 (3), 563–566. 10.1016/0005-2760(70)90131-1. [DOI] [PubMed] [Google Scholar]
Harland W. A.; Gilbert J. D.; Steel G.; Brooks C. J. W. Lipids of Human Atheroma Part 5. The Occurrence of a New Group of Polar Sterol Esters in Various Stages of Human Atherosclerosis. Atherosclerosis 1971, 13 (2), 239–246. 10.1016/0021-9150(71)90026-8. [DOI] [PubMed] [Google Scholar]
Harland W. A.; Gilbert J. D.; Brooks C. J. W. Lipids of Human Atheroma: VIII. Oxidised Derivatives of Cholesteryl Linoleate. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1973, 316 (3), 378–385. 10.1016/0005-2760(73)90076-3. [DOI] [PubMed] [Google Scholar]
Kühn H.; Heydeck D.; Hugou I.; Gniwotta C. In Vivo Action of 15-Lipoxygenase in Early Stages of Human Atherogenesis. J. Clin. Invest. 1997, 99 (5), 888–893. 10.1172/JCI119253. [DOI] [PMC free article] [PubMed] [Google Scholar]
Parthasarathy S.; Steinberg D.; Witztum J. L. The Role of Oxidized Low-Density Lipoproteins in the Pathogenesis of Atherosclerosis. Annu. Rev. Med. 1992, 43 (1), 219–225. 10.1146/annurev.me.43.020192.001251. [DOI] [PubMed] [Google Scholar]
Steinberg D. Low Density Lipoprotein Oxidation and Its Pathobiological Significance. J. Biol. Chem. 1997, 272 (34), 20963–20966. 10.1074/jbc.272.34.20963. [DOI] [PubMed] [Google Scholar]
Kühn H.; Belkner J.; Wiesner R.; Schewe T.; Lankin V. Z.; Tikhae A. K. Structure Elucidation of Oxygenated Lipids in Human Atherosclerotic Lesions. Eicosanoids 1992, 5, 17–22. [PubMed] [Google Scholar]
Waddington E.; Sienuarine K.; Puddey I.; Croft K. Identification and Quantitation of Unique Fatty Acid Oxidation Products in Human Atherosclerotic Plaque Using High-Performance Liquid Chromatography. Anal. Biochem. 2001, 292 (2), 234–244. 10.1006/abio.2001.5075. [DOI] [PubMed] [Google Scholar]
Kühn H.; Belkner G.; Zaiss S.; Fährenklemper T.; Wohlfeil S. Involvement of 15-Lipoxygenase in Early Stages of Atherogenesis. J. Exp. Med. 1994, 179 (6), 1903–1911. 10.1084/jem.179.6.1903. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kühn H.; Belkner J.; Suzuki H.; Yamamoto S. Oxidative Modification of Human Lipoproteins by Lipoxygenases of Different Positional Specificities. J. Lipid Res. 1994, 35 (10), 1749–1759. 10.1016/S0022-2275(20)39770-4. [DOI] [PubMed] [Google Scholar]
Parthasarathy S.; Wieland E.; Steinberg D. A Role for Endothelial Cell Lipoxygenase in the Oxidative Modification of Low Density Lipoprotein. Proc. Natl. Acad. Sci. U. S. A. 1989, 86 (3), 1046–1050. 10.1073/pnas.86.3.1046. [DOI] [PMC free article] [PubMed] [Google Scholar]
McNally A. K.; Chisolm G. M. 3rd; Morel D. W.; Cathcart M. K. Activated Human Monocytes Oxidize Low-Density Lipoprotein by a Lipoxygenase-Dependent Pathway. J. Immunol. 1990, 145 (1), 254–259. 10.4049/jimmunol.145.1.254. [DOI] [PubMed] [Google Scholar]
Cyrus T.; Witztum J. L.; Rader D. J.; Tangirala R.; Fazio S.; Linton M. F.; Funk C. D. Disruption of the 12/15-Lipoxygenase Gene Diminishes Atherosclerosis in Apo E-Deficient Mice. J. Clin. Invest. 1999, 103 (11), 1597–1604. 10.1172/JCI5897. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bojic L. A.; McLaren D. G.; Harms A. C.; Hankemeier T.; Dane A.; Wang S.-P.; Rosa R.; Previs S. F.; Johns D. G.; Castro-Perez J. M. Quantitative Profiling of Oxylipins in Plasma and Atherosclerotic Plaques of Hypercholesterolemic Rabbits. Anal. Bioanal. Chem. 2016, 408 (1), 97–105. 10.1007/s00216-015-9105-4. [DOI] [PubMed] [Google Scholar]
Zhu S.; Zhao Y.; Liu L.; Xu Y.; Zhu J.; Li W.; Liu Y.; Xia M. High Plant Protein Diet Ameliorated Hepatic Lipid Accumulation Through the Modulation of Gut Microbiota. Mol. Nutr. Food Res. 2023, 67 (24), e2300515 10.1002/mnfr.202300515. [DOI] [PubMed] [Google Scholar]
Ishizaki T.; Ozawa T.; Voelkel N. F. Leukotoxins and the Lung. Pulm. Pharmacol. Ther. 1999, 12 (3), 145–155. 10.1006/pupt.1999.0179. [DOI] [PubMed] [Google Scholar]
Balgoma D.; Yang M.; Sjödin M.; Snowden S.; Karimi R.; Levänen B.; Merikallio H.; Kaarteenaho R.; Palmberg L.; Larsson K.; Erle D. J.; Dahlén S.-E.; Dahlén B.; Sköld C. M.; Wheelock Å. M.; Wheelock C. E. Linoleic Acid-Derived Lipid Mediators Increase in a Female-Dominated Subphenotype of COPD. Eur. Respir. J. 2016, 47 (6), 1645–1656. 10.1183/13993003.01080-2015. [DOI] [PubMed] [Google Scholar]
Schmelzer K. R.; Wheelock Å. M.; Dettmer K.; Morin D.; Hammock B. D. The Role of Inflammatory Mediators in the Synergistic Toxicity of Ozone and 1-Nitronaphthalene in Rat Airways. Environ. Health Perspect. 2006, 114 (9), 1354–1360. 10.1289/ehp.8373. [DOI] [PMC free article] [PubMed] [Google Scholar]
Gouveia-Figueira S.; Karimpour M.; Bosson J. A.; Blomberg A.; Unosson J.; Sehlstedt M.; Pourazar J.; Sandström T.; Behndig A. F.; Nording M. L. Mass Spectrometry Profiling Reveals Altered Plasma Levels of Monohydroxy Fatty Acids and Related Lipids in Healthy Humans after Controlled Exposure to Biodiesel Exhaust. Anal. Chim. Acta 2018, 1018, 62–69. 10.1016/j.aca.2018.02.032. [DOI] [PubMed] [Google Scholar]
Gouveia-Figueira S.; Karimpour M.; Bosson J. A.; Blomberg A.; Unosson J.; Pourazar J.; Sandström T.; Behndig A. F.; Nording M. L. Mass Spectrometry Profiling of Oxylipins, Endocannabinoids, and N-Acylethanolamines in Human Lung Lavage Fluids Reveals Responsiveness of Prostaglandin E₂ and Associated Lipid Metabolites to Biodiesel Exhaust Exposure. Anal. Bioanal. Chem. 2017, 409 (11), 2967–2980. 10.1007/s00216-017-0243-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
Johnson R. K.; Manke J.; Campbell M.; Armstrong M.; Boorgula M. P.; Pinheiro G.; Santana C. V. N.; Mathias R. A.; Barnes K. C.; Cruz A.; Reisdorph N.; Figueiredo C. A. Lipid Mediators Are Detectable in the Nasal Epithelium and Differ by Asthma Status in Female Subjects. J. Allergy Clin. Immunol. 2022, 150 (4), 965–971. 10.1016/j.jaci.2022.02.026. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kelly R. S.; Mendez K. M.; Huang M.; Hobbs B. D.; Clish C. B.; Gerszten R.; Cho M. H.; Wheelock C. E.; McGeachie M. J.; Chu S. H.; Celedón J. C.; Weiss S. T.; Lasky-Su J. Metabo-Endotypes of Asthma Reveal Differences in Lung Function: Discovery and Validation in Two TOPMed Cohorts. Am. J. Respir. Crit. Care Med. 2022, 205 (3), 288–299. 10.1164/rccm.202105-1268OC. [DOI] [PMC free article] [PubMed] [Google Scholar]
Liu Y.; Zhang X.; Zhang L.; Oliver B. G.; Wang H. G.; Liu Z. P.; Chen Z. H.; Wood L.; Hsu A. C.-Y.; Xie M.; McDonald V.; Wan H. J.; Luo F. M.; Liu D.; Li W. M.; Wang G. Sputum Metabolomic Profiling Reveals Metabolic Pathways and Signatures Associated With Inflammatory Phenotypes in Patients With Asthma. Allergy Asthma Immunol. Res. 2022, 14 (4), 393–411. 10.4168/aair.2022.14.4.393. [DOI] [PMC free article] [PubMed] [Google Scholar]
Panda L.; Gheware A.; Rehman R.; Yadav M. K.; Jayaraj B. S.; Madhunapantula S. V.; Mahesh P. A.; Ghosh B.; Agrawal A.; Mabalirajan U. Linoleic Acid Metabolite Leads to Steroid Resistant Asthma Features Partially through NF-ΚB. Sci. Rep. 2017, 7, 9565. 10.1038/s41598-017-09869-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
Henricks P.; Engels F.; Vanderlinde H.; Garssen J.; Nijkamp F. 13-Hydroxy-Linoleic Acid Induces Airway Hyperresponsiveness to Histamine and Methacholine in Guinea Pigs in Vivo. J. Allergy Clin. Immunol. 1995, 96 (1), 36–43. 10.1016/S0091-6749(95)70030-7. [DOI] [PubMed] [Google Scholar]
McReynolds C. B.; Cortes-Puch I.; Ravindran R.; Khan I. H.; Hammock B. G.; Shih P. B.; Hammock B. D.; Yang J. Plasma Linoleate Diols Are Potential Biomarkers for Severe COVID-19 Infections. Front. Physiol. 2021, 12, 663869. 10.3389/fphys.2021.663869. [DOI] [PMC free article] [PubMed] [Google Scholar]
Karu N.; Kindt A.; Lamont L.; van Gammeren A. J.; Ermens A. A. M.; Harms A. C.; Portengen L.; Vermeulen R. C. H.; Dik W. A.; Langerak A. W.; van der Velden V. H. J.; Hankemeier T. Plasma Oxylipins and Their Precursors Are Strongly Associated with COVID-19 Severity and with Immune Response Markers. Metabolites 2022, 12 (7), 619. 10.3390/metabo12070619. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kovarik J. J.; Bileck A.; Hagn G.; Meier-Menches S. M.; Frey T.; Kaempf A.; Hollenstein M.; Shoumariyeh T.; Skos L.; Reiter B.; Gerner M. C.; Spannbauer A.; Hasimbegovic E.; Schmidl D.; Garhöfer G.; Gyöngyösi M.; Schmetterer K. G.; Gerner C. A Multi-Omics Based Anti-Inflammatory Immune Signature Characterizes Long COVID-19 Syndrome. iScience 2023, 26 (1), 105717. 10.1016/j.isci.2022.105717. [DOI] [PMC free article] [PubMed] [Google Scholar]
Arnardottir H.; Pawelzik S.; Sarajlic P.; Quaranta A.; Kolmert J.; Religa D.; Wheelock C. E.; Bäck M. Immunomodulation by Intravenous Omega-3 Fatty Acid Treatment in Older Subjects Hospitalized for COVID-19: A Single-blind Randomized Controlled Trial. Clin. Transl. Med. 2022, 12 (9), e895 10.1002/ctm2.895. [DOI] [PMC free article] [PubMed] [Google Scholar]
Edin M. L.; Gruzdev A.; Graves J. P.; Lih F. B.; Morisseau C.; Ward J. M.; Hammock B. D.; Bosio C. M.; Zeldin D. C. Effects of SEH Inhibition on the Eicosanoid and Cytokine Storms in SARS-CoV-2-Infected Mice. FASEB J. Off. Publ. Fed. Am. Soc. Exp. Biol. 2024, 38 (10), e23692 10.1096/fj.202302202RR. [DOI] [PMC free article] [PubMed] [Google Scholar]
Druzak S.; Iffrig E.; Roberts B. R.; Zhang T.; Fibben K. S.; Sakurai Y.; Verkerke H. P.; Rostad C. A.; Chahroudi A.; Schneider F.; Wong A. K. H.; Roberts A. M.; Chandler J. D.; Kim S. O.; Mosunjac M.; Mosunjac M.; Geller R.; Albizua I.; Stowell S. R.; Arthur C. M.; Anderson E. J.; Ivanova A. A.; Ahn J.; Liu X.; Maner-Smith K.; Bowen T.; Paiardini M.; Bosinger S. E.; Roback J. D.; Kulpa D. A.; Silvestri G.; Lam W. A.; Ortlund E. A.; Maier C. L. Multiplatform Analyses Reveal Distinct Drivers of Systemic Pathogenesis in Adult versus Pediatric Severe Acute COVID-19. Nat. Commun. 2023, 14 (1), 1638. 10.1038/s41467-023-37269-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
Tam V. C.; Quehenberger O.; Oshansky C. M.; Suen R.; Armando A. M.; Treuting P. M.; Thomas P. G.; Dennis E. A.; Aderem A. Lipidomic Profiling of Influenza Infection Identifies Mediators That Induce and Resolve Inflammation. Cell 2013, 154 (1), 213–227. 10.1016/j.cell.2013.05.052. [DOI] [PMC free article] [PubMed] [Google Scholar]
Maciejewska D.; Ossowski P.; Drozd A.; Ryterska K.; Jamioł-Milc D.; Banaszczak M.; Kaczorowska M.; Sabinicz A.; Raszeja-Wyszomirska J.; Stachowska E. Metabolites of Arachidonic Acid and Linoleic Acid in Early Stages of Non-Alcoholic Fatty Liver Disease—A Pilot Study. Prostaglandins Other Lipid Mediat. 2015, 121, 184–189. 10.1016/j.prostaglandins.2015.09.003. [DOI] [PubMed] [Google Scholar]
Mazi T. A.; Borkowski K.; Fiehn O.; Bowlus C. L.; Sarkar S.; Matsukuma K.; Ali M. R.; Kieffer D. A.; Wan Y.-J. Y.; Stanhope K. L.; Havel P. J.; Newman J. W.; Medici V. Plasma Oxylipin Profile Discriminates Ethnicities in Subjects with Non-Alcoholic Steatohepatitis: An Exploratory Analysis. Metabolites 2022, 12 (2), 192. 10.3390/metabo12020192. [DOI] [PMC free article] [PubMed] [Google Scholar]
Zirnheld K. H.; Warner D. R.; Warner J. B.; Hardesty J. E.; McClain C. J.; Kirpich I. A. Dietary Fatty Acids and Bioactive Fatty Acid Metabolites in Alcoholic Liver Disease. Liver Res. 2019, 3 (3), 206–217. 10.1016/j.livres.2019.10.001. [DOI] [Google Scholar]
Warner D. R.; Liu H.; Ghosh Dastidar S.; Warner J. B.; Prodhan M. A. I.; Yin X.; Zhang X.; Feldstein A. E.; Gao B.; Prough R. A.; McClain C. J.; Kirpich I. A. Ethanol and Unsaturated Dietary Fat Induce Unique Patterns of Hepatic ω-6 and ω-3 PUFA Oxylipins in a Mouse Model of Alcoholic Liver Disease. PLoS One 2018, 13, e0204119 10.1371/journal.pone.0204119. [DOI] [PMC free article] [PubMed] [Google Scholar]
Warner D.; Vatsalya V.; Zirnheld K. H.; Warner J. B.; Hardesty J. E.; Umhau J. C.; McClain C. J.; Maddipati K.; Kirpich I. A. Linoleic Acid-Derived Oxylipins Differentiate Early Stage Alcoholic Hepatitis From Mild Alcohol-Associated Liver Injury. Hepatol. Commun. 2021, 5 (6), 947–960. 10.1002/hep4.1686. [DOI] [PMC free article] [PubMed] [Google Scholar]
López-Vicario C.; Checa A.; Urdangarin A.; Aguilar F.; Alcaraz-Quiles J.; Caraceni P.; Amorós A.; Pavesi M.; Gómez-Cabrero D.; Trebicka J.; Oettl K.; Moreau R.; Planell N.; Arroyo V.; Wheelock C. E.; Clària J. Targeted Lipidomics Reveals Extensive Changes in Circulating Lipid Mediators in Patients with Acutely Decompensated Cirrhosis☆. J. Hepatol. 2020, 73 (4), 817–828. 10.1016/j.jhep.2020.03.046. [DOI] [PubMed] [Google Scholar]
Yoshida Y.; Kodai S.; Takemura S.; Minamiyama Y.; Niki E. Simultaneous Measurement of F2-Isoprostane, Hydroxyoctadecadienoic Acid, Hydroxyeicosatetraenoic Acid, and Hydroxycholesterols from Physiological Samples. Anal. Biochem. 2008, 379 (1), 105–115. 10.1016/j.ab.2008.04.028. [DOI] [PubMed] [Google Scholar]
Lu Y.; Fang J.; Zou L.; Cui L.; Liang X.; Lim S. G.; Dan Y.-Y.; Ong C. N. Omega-6-Derived Oxylipin Changes in Serum of Patients with Hepatitis B Virus-Related Liver Diseases. Metabolomics 2018, 14 (3), 26. 10.1007/s11306-018-1326-z. [DOI] [PubMed] [Google Scholar]
Warner J.; Hardesty J.; Zirnheld K.; McClain C.; Warner D.; Kirpich I. Soluble Epoxide Hydrolase Inhibition in Liver Diseases: A Review of Current Research and Knowledge Gaps. Biology 2020, 9 (6), 124. 10.3390/biology9060124. [DOI] [PMC free article] [PubMed] [Google Scholar]
Helmstädter M.; Schmidt J.; Kaiser A.; Weizel L.; Proschak E.; Merk D. Differential Therapeutic Effects of FXR Activation, SEH Inhibition, and Dual FXR/SEH Modulation in NASH in Diet-Induced Obese Mice. ACS Pharmacol. Transl. Sci. 2021, 4 (2), 966–979. 10.1021/acsptsci.1c00041. [DOI] [PMC free article] [PubMed] [Google Scholar]
Oliw E. H.; Su C.; Skogström T.; Benthin G. Analysis of Novel Hydroperoxides and Other Metabolites of Oleic, Linoleic, and Linolenic Acids by Liquid Chromatography-Mass Spectrometry with Ion Trap MSⁿ. Lipids 1998, 33 (9), 843–852. 10.1007/s11745-998-0280-0. [DOI] [PubMed] [Google Scholar]
Schenck E. J.; Plataki M.; Wheelock C. E. A Lipid Map for Community-Acquired Pneumonia with Sepsis: Observation Is the First Step in Scientific Progress. Am. J. Respir. Crit. Care Med. 2024, 209 (8), 903–904. 10.1164/rccm.202401-0213ED. [DOI] [PMC free article] [PubMed] [Google Scholar]
Chouchane O.; Schuurman A. R.; Reijnders T. D. Y.; Peters-Sengers H.; Butler J. M.; Uhel F.; Schultz M. J.; Bonten M. J.; Cremer O. L.; Calfee C. S.; Matthay M. A.; Langley R. J.; Alipanah-Lechner N.; Kingsmore S. F.; Rogers A.; van Weeghel M.; Vaz F. M.; van der Poll T. The Plasma Lipidomic Landscape in Patients with Sepsis Due to Community-Acquired Pneumonia. Am. J. Respir. Crit. Care Med. 2024, 209 (8), 973–986. 10.1164/rccm.202308-1321OC. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bruegel M.; Ludwig U.; Kleinhempel A.; Petros S.; Kortz L.; Ceglarek U.; Holdt L. M.; Thiery J.; Fiedler G. M. Sepsis-Associated Changes of the Arachidonic Acid Metabolism and Their Diagnostic Potential in Septic Patients. Crit. Care Med. 2012, 40 (5), 1478–1486. 10.1097/CCM.0b013e3182416f05. [DOI] [PubMed] [Google Scholar]
Amunugama K.; Pike D. P.; Ford D. A. The Lipid Biology of Sepsis. J. Lipid Res. 2021, 62, 100090. 10.1016/j.jlr.2021.100090. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bulger E. M.; Maier R. V. Lipid Mediators in the Pathophysiology of Critical Illness. Crit. Care Med. 2000, 28, N27–N36. 10.1097/00003246-200004001-00004. [DOI] [PubMed] [Google Scholar]
Tunctan B.; Senol S. P.; Temiz-Resitoglu M.; Guden D. S.; Sahan-Firat S.; Falck J. R.; Malik K. U. Eicosanoids Derived from Cytochrome P450 Pathway of Arachidonic Acid and Inflammatory Shock. Prostaglandins Other Lipid Mediat. 2019, 145, 106377. 10.1016/j.prostaglandins.2019.106377. [DOI] [PubMed] [Google Scholar]
Padovan M. G.; Norling L. V. Pro-Resolving Lipid Mediators in Sepsis and Critical Illness. Curr. Opin. Clin. Nutr. Metab. Care 2020, 23 (2), 76–81. 10.1097/MCO.0000000000000633. [DOI] [PubMed] [Google Scholar]
Dalli J.; Colas R. A.; Quintana C.; Barragan-Bradford D.; Hurwitz S.; Levy B. D.; Choi A. M.; Serhan C. N.; Baron R. M. Human Sepsis Eicosanoid and Proresolving Lipid Mediator Temporal Profiles: Correlations With Survival and Clinical Outcomes. Crit. Care Med. 2017, 45 (1), 58–68. 10.1097/CCM.0000000000002014. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hamaguchi M.; Wu H. N.; Tanaka M.; Tsuda N.; Tantengco O. A. G.; Matsushima T.; Nakao T.; Ishibe T.; Sakata I.; Yanagihara I. A Case Series of the Dynamics of Lipid Mediators in Patients with Sepsis. Acute Med. Surg. 2019, 6 (4), 413–418. 10.1002/ams2.443. [DOI] [PMC free article] [PubMed] [Google Scholar]
Montague B.; Summers A.; Bhawal R.; Anderson E. T.; Kraus-Malett S.; Zhang S.; Goggs R. Identifying Potential Biomarkers and Therapeutic Targets for Dogs with Sepsis Using Metabolomics and Lipidomics Analyses. PloS One 2022, 17 (7), e0271137 10.1371/journal.pone.0271137. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sulaiman D.; Wu D.; Black L. P.; Williams K. J.; Graim K.; Datta S.; Reddy S. T.; Guirgis F. W. Lipidomic Changes in a Novel Sepsis Outcome-Based Analysis Reveals Potent pro-Inflammatory and pro-Resolving Signaling Lipids. Clin. Transl. Sci. 2024, 17 (3), e13745 10.1111/cts.13745. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ahmad N. S.; Tan T. L.; Arifin K. T.; Ngah W. Z. W.; Yusof Y. A. M. High SPLA2-IIA Level Is Associated with Eicosanoid Metabolism in Patients with Bacterial Sepsis Syndrome. PloS One 2020, 15 (3), e0230285 10.1371/journal.pone.0230285. [DOI] [PMC free article] [PubMed] [Google Scholar]
Tan T. L.; Goh Y. Y. The Role of Group IIA Secretory Phospholipase A2 (SPLA2-IIA) as a Biomarker for the Diagnosis of Sepsis and Bacterial Infection in Adults-A Systematic Review. PloS One 2017, 12 (7), e0180554 10.1371/journal.pone.0180554. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sala-Vila A.; Miles E. A.; Calder P. C. Fatty Acid Composition Abnormalities in Atopic Disease: Evidence Explored and Role in the Disease Process Examined. Clin. Exp. Allergy 2008, 38 (9), 1432–1450. 10.1111/j.1365-2222.2008.03072.x. [DOI] [PubMed] [Google Scholar]
Rucci E.; den Dekker H. T.; de Jongste J. C.; Steenweg-de-Graaff J.; Gaillard R.; Pasmans S. G.; Hofman A.; Tiemeier H.; Jaddoe V. W. V.; Duijts L. Maternal Fatty Acid Levels during Pregnancy, Childhood Lung Function and Atopic Diseases. The Generation R Study. Clin. Exp. Allergy 2016, 46 (3), 461–471. 10.1111/cea.12613. [DOI] [PubMed] [Google Scholar]
Miyake Y.; Tanaka K.; Sasaki S.; Arakawa M. Polyunsaturated Fatty Acid Intake and Prevalence of Eczema and Rhinoconjunctivitis in Japanese Children: The Ryukyus Child Health Study. BMC Public Health 2011, 11 (1), 358. 10.1186/1471-2458-11-358. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lundström S. L.; Yang J.; Källberg H. J.; Thunberg S.; Gafvelin G.; Haeggström J. Z.; Grönneberg R.; Grunewald J.; van Hage M.; Hammock B. D.; Eklund A.; Wheelock Å. M.; Wheelock C. E. Allergic Asthmatics Show Divergent Lipid Mediator Profiles from Healthy Controls Both at Baseline and Following Birch Pollen Provocation. PLoS One 2012, 7, e33780 10.1371/journal.pone.0033780. [DOI] [PMC free article] [PubMed] [Google Scholar]
Lee S.-Y.; Park Y. M.; Yoo H. J.; Suh D. I.; Shin Y. H.; Kim K. W.; Ahn K.; Hong S.-J. Gut Linoleic Acid Is Associated with the Severity of Atopic Dermatitis and Sensitization to Egg White/Milk in Infants. Pediatr. Allergy Immunol. 2021, 32 (2), 382–385. 10.1111/pai.13393. [DOI] [PubMed] [Google Scholar]
Zheng P.; Yan G.; Zhang Y.; Huang H.; Luo W.; Xue M.; Li N.; Wu J.-L.; Sun B. Metabolomics Reveals Process of Allergic Rhinitis Patients with Single- and Double-Species Mite Subcutaneous Immunotherapy. Metabolites 2021, 11 (9), 613. 10.3390/metabo11090613. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ellul S.; Marx W.; Collier F.; Saffery R.; Tang M.; Burgner D.; Carlin J.; Vuillermin P.; Ponsonby A.-L. Plasma Metabolomic Profiles Associated with Infant Food Allergy with Further Consideration of Other Early Life Factors. Prostaglandins Leukot. Essent. Fatty Acids 2020, 159, 102099. 10.1016/j.plefa.2020.102099. [DOI] [PubMed] [Google Scholar]
Yen C.-H.; Dai Y.-S.; Yang Y.-H.; Wang L.-C.; Lee J.-H.; Chiang B.-L. Linoleic Acid Metabolite Levels and Transepidermal Water Loss in Children with Atopic Dermatitis. Ann. Allergy. Asthma. Immunol. 2008, 100 (1), 66–73. 10.1016/S1081-1206(10)60407-3. [DOI] [PubMed] [Google Scholar]
Berth-Jones J.; Graham-Brown R. a. C. Placebo-Controlled Trial of Essential Fatty Acid Supplementation in Atopic Dermatitis. Lancet 1993, 341 (8860), 1557–1560. 10.1016/0140-6736(93)90697-F. [DOI] [PubMed] [Google Scholar]
Chiba T.; Nakahara T.; Kohda F.; Ichiki T.; Manabe M.; Furue M. Measurement of Trihydroxy-Linoleic Acids in Stratum Corneum by Tape-Stripping: Possible Biomarker of Barrier Function in Atopic Dermatitis. PLoS One 2019, 14 (1), e0210013 10.1371/journal.pone.0210013. [DOI] [PMC free article] [PubMed] [Google Scholar]
Kaikiri H.; Miyamoto J.; Kawakami T.; Park S.-B.; Kitamura N.; Kishino S.; Yonejima Y.; Hisa K.; Watanabe J.; Ogita T.; Ogawa J.; Tanabe S.; Suzuki T. Supplemental Feeding of a Gut Microbial Metabolite of Linoleic Acid, 10-Hydroxy-cis-12-Octadecenoic Acid, Alleviates Spontaneous Atopic Dermatitis and Modulates Intestinal Microbiota in NC/Nga Mice. Int. J. Food Sci. Nutr. 2017, 68 (8), 941–951. 10.1080/09637486.2017.1318116. [DOI] [PubMed] [Google Scholar]
Fujii M.; Nakashima H.; Tomozawa J.; Shimazaki Y.; Ohyanagi C.; Kawaguchi N.; Ohya S.; Kohno S.; Nabe T. Deficiency of n-6 Polyunsaturated Fatty Acids Is Mainly Responsible for Atopic Dermatitis-like Pruritic Skin Inflammation in Special Diet-Fed Hairless Mice. Exp. Dermatol. 2013, 22 (4), 272–277. 10.1111/exd.12120. [DOI] [PubMed] [Google Scholar]
Plötz S. G.; Traidl-Hoffmann C.; Feussner I.; Kasche A.; Feser A.; Ring J.; Jakob T.; Behrendt H. Chemotaxis and Activation of Human Peripheral Blood Eosinophils Induced by Pollen-Associated Lipid Mediators. J. Allergy Clin. Immunol. 2004, 113 (6), 1152–1160. 10.1016/j.jaci.2004.03.011. [DOI] [PubMed] [Google Scholar]
Traidl-Hoffmann C.; Kasche A.; Jakob T.; Huger M.; Plötz S.; Feussner I.; Ring J.; Behrendt H. Lipid Mediators from Pollen Act as Chemoattractants and Activators of Polymorphonuclear Granulocytes. J. Allergy Clin. Immunol. 2002, 109 (5), 831–838. 10.1067/mai.2002.124655. [DOI] [PubMed] [Google Scholar]
Traidl-Hoffmann C.; Mariani V.; Hochrein H.; Karg K.; Wagner H.; Ring J.; Mueller M. J.; Jakob T.; Behrendt H. Pollen-Associated Phytoprostanes Inhibit Dendritic Cell Interleukin-12 Production and Augment T Helper Type 2 Cell Polarization. J. Exp. Med. 2005, 201 (4), 627–636. 10.1084/jem.20041065. [DOI] [PMC free article] [PubMed] [Google Scholar]
Mariani V.; Gilles S.; Jakob T.; Thiel M.; Mueller M. J.; Ring J.; Behrendt H.; Traidl-Hoffmann C. Immunomodulatory Mediators from Pollen Enhance the Migratory Capacity of Dendritic Cells and License Them for Th2 Attraction. J. Immunol. 2007, 178 (12), 7623–7631. 10.4049/jimmunol.178.12.7623. [DOI] [PubMed] [Google Scholar]
Gilles S.; Mariani V.; Bryce M.; Mueller M. J.; Ring J.; Jakob T.; Pastore S.; Behrendt H.; Traidl-Hoffmann C. Pollen-Derived E1-Phytoprostanes Signal via PPAR-γ and NF-ΚB-Dependent Mechanisms. J. Immunol. 2009, 182 (11), 6653–6658. 10.4049/jimmunol.0802613. [DOI] [PubMed] [Google Scholar]
Gilles S.; Jacoby D.; Blume C.; Mueller M. J.; Jakob T.; Behrendt H.; Schaekel K.; Traidl-Hoffmann C. Pollen-Derived Low-Molecular Weight Factors Inhibit 6-Sulfo LacNAc+ Dendritic Cells’ Capacity to Induce T-Helper Type 1 Responses. Clin. Exp. Allergy 2010, 40 (2), 269–278. 10.1111/j.1365-2222.2009.03369.x. [DOI] [PubMed] [Google Scholar]
González Roldán N.; Engel R.; Düpow S.; Jakob K.; Koops F.; Orinska Z.; Vigor C.; Oger C.; Galano J.-M.; Durand T.; Jappe U.; Duda K. A. Lipid Mediators From Timothy Grass Pollen Contribute to the Effector Phase of Allergy and Prime Dendritic Cells for Glycolipid Presentation. Front. Immunol. 2019, 10, 974. 10.3389/fimmu.2019.00974. [DOI] [PMC free article] [PubMed] [Google Scholar]
Soh W. T.; Aglas L.; Mueller G. A.; Gilles S.; Weiss R.; Scheiblhofer S.; Huber S.; Scheidt T.; Thompson P. M.; Briza P.; London R. E.; Traidl-Hoffmann C.; Cabrele C.; Brandstetter H.; Ferreira F. Multiple Roles of Bet v 1 Ligands in Allergen Stabilization and Modulation of Endosomal Protease Activity. Allergy 2019, 74 (12), 2382–2393. 10.1111/all.13948. [DOI] [PMC free article] [PubMed] [Google Scholar]
Medina S.; Gil-Izquierdo Á.; Abu-Reidah I. M.; Durand T.; Bultel-Poncé V.; Galano J.-M.; Domínguez-Perles R. Evaluation of Phoenix dactylifera Edible Parts and Byproducts as Sources of Phytoprostanes and Phytofurans. J. Agric. Food Chem. 2020, 68 (33), 8942–8950. 10.1021/acs.jafc.0c03364. [DOI] [PubMed] [Google Scholar]
Bublin M.; Eiwegger T.; Breiteneder H. Do Lipids Influence the Allergic Sensitization Process?. J. Allergy Clin. Immunol. 2014, 134 (3), 521–529. 10.1016/j.jaci.2014.04.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hagemann P. M.; Nsiah-Dosu S.; Hundt J. E.; Hartmann K.; Orinska Z. Modulation of Mast Cell Reactivity by Lipids: The Neglected Side of Allergic Diseases. Front. Immunol. 2019, 10, 1174. 10.3389/fimmu.2019.01174. [DOI] [PMC free article] [PubMed] [Google Scholar]
Dreisbach A. W.; Rice J. C.; Japa S.; Newman J. W.; Sigel A.; Gill R. S.; Hess A. E.; Cemo A. C.; Fonseca J. P.; Hammock B. D.; Lertora J. J. L.; Hamm L. L. Salt Loading Increases Urinary Excretion of Linoleic Acid Diols and Triols in Healthy Human Subjects. Hypertension 2008, 51 (3), 755–761. 10.1161/HYPERTENSIONAHA.107.100123. [DOI] [PubMed] [Google Scholar]
Holla V. R.; Makita K.; Zaphiropoulos P. G.; Capdevila J. H. The Kidney Cytochrome P-450 2C23 Arachidonic Acid Epoxygenase Is Upregulated during Dietary Salt Loading. J. Clin. Invest. 1999, 104 (6), 751–760. 10.1172/JCI7013. [DOI] [PMC free article] [PubMed] [Google Scholar]
Sun P.; Lin D.-H.; Wang T.; Babilonia E.; Wang Z.; Jin Y.; Kemp R.; Nasjletti A.; Wang W.-H. Low Na Intake Suppresses Expression of CYP2C23 and Arachidonic Acid-Induced Inhibition of ENaC. Am. J. Physiol.-Ren. Physiol. 2006, 291 (6), F1192–F1200. 10.1152/ajprenal.00112.2006. [DOI] [PubMed] [Google Scholar]
Boldt C.; Röschel T.; Himmerkus N.; Plain A.; Bleich M.; Labes R.; Blum M.; Krause H.; Magheli A.; Giesecke T.; Mutig K.; Rothe M.; Weldon S. M.; Dragun D.; Schunck W.-H.; Bachmann S.; Paliege A. Vasopressin Lowers Renal Epoxyeicosatrienoic Acid Levels by Activating Soluble Epoxide Hydrolase. Am. J. Physiol.-Ren. Physiol. 2016, 311 (6), F1198–F1210. 10.1152/ajprenal.00062.2016. [DOI] [PubMed] [Google Scholar]
McMahon B. A.; Chawla L. S. The Furosemide Stress Test: Current Use and Future Potential. Ren. Fail. 2021, 43 (1), 830–839. 10.1080/0886022X.2021.1906701. [DOI] [PMC free article] [PubMed] [Google Scholar]
Bech A. P.; Wetzels J. F. M.; Nijenhuis T. Reference Values of Renal Tubular Function Tests Are Dependent on Age and Kidney Function. Physiol. Rep. 2017, 5 (23), e13542 10.14814/phy2.13542. [DOI] [PMC free article] [PubMed] [Google Scholar]
Grapov D.; Adams S. H.; Pedersen T. L.; Garvey W. T.; Newman J. W. Type 2 Diabetes Associated Changes in the Plasma Non-Esterified Fatty Acids, Oxylipins and Endocannabinoids. PLoS One 2012, 7 (11), e48852 10.1371/journal.pone.0048852. [DOI] [PMC free article] [PubMed] [Google Scholar]
Umeno A.; Shichiri M.; Ishida N.; Hashimoto Y.; Abe K.; Kataoka M.; Yoshino K.; Hagihara Y.; Aki N.; Funaki M.; Asada Y.; Yoshida Y. Singlet Oxygen Induced Products of Linoleates, 10- and 12-(Z,E)-Hydroxyoctadecadienoic Acids (HODE), Can Be Potential Biomarkers for Early Detection of Type 2 Diabetes. PLoS One 2013, 8 (5), e63542 10.1371/journal.pone.0063542. [DOI] [PMC free article] [PubMed] [Google Scholar]
Dalle C.; Tournayre J.; Mainka M.; Basiak-Rasała A.; Pétéra M.; Lefèvre-Arbogast S.; Dalloux-Chioccioli J.; Deschasaux-Tanguy M.; Lécuyer L.; Kesse-Guyot E.; Fezeu L. K.; Hercberg S.; Galan P.; Samieri C.; Zatońska K.; Calder P. C.; Fiil Hjorth M.; Astrup A.; Mazur A.; Bertrand-Michel J.; Schebb N. H.; Szuba A.; Touvier M.; Newman J. W.; Gladine C. The Plasma Oxylipin Signature Provides a Deep Phenotyping of Metabolic Syndrome Complementary to the Clinical Criteria. Int. J. Mol. Sci. 2022, 23 (19), 11688. 10.3390/ijms231911688. [DOI] [PMC free article] [PubMed] [Google Scholar]
Criscuolo A.; Nepachalovich P.; Garcia-del Rio D. F.; Lange M.; Ni Z.; Baroni M.; Cruciani G.; Goracci L.; Blüher M.; Fedorova M. Analytical and Computational Workflow for In-Depth Analysis of Oxidized Complex Lipids in Blood Plasma. Nat. Commun. 2022, 13 (1), 6547. 10.1038/s41467-022-33225-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
Hateley C.; Olona A.; Halliday L.; Edin M. L.; Ko J.-H.; Forlano R.; Terra X.; Lih F. B.; Beltrán-Debón R.; Manousou P.; Purkayastha S.; Moorthy K.; Thursz M. R.; Zhang G.; Goldin R. D.; Zeldin D. C.; Petretto E.; Behmoaras J. Multi-Tissue Profiling of Oxylipins Reveal a Conserved up-Regulation of Epoxide:Diol Ratio That Associates with White Adipose Tissue Inflammation and Liver Steatosis in Obesity. EBioMedicine 2024, 103, 105127. 10.1016/j.ebiom.2024.105127. [DOI] [PMC free article] [PubMed] [Google Scholar]
Ziboh V. A.; Cho Y.; Mani I.; Xi S. Biological Significance of Essential Fatty Acids/Prostanoids/ Lipoxygenase-Derived Monohydroxy Fatty Acids in the Skin. Arch. Pharm. Res. 2002, 25 (6), 747–758. 10.1007/BF02976988. [DOI] [PubMed] [Google Scholar]
Conconi A.; Smerdon M. J.; Howe G. A.; Ryan C. A. The Octadecanoid Signalling Pathway in Plants Mediates a Response to Ultraviolet Radiation. Nature 1996, 383 (6603), 826–829. 10.1038/383826a0. [DOI] [PubMed] [Google Scholar]
Menke F. L. H.; Parchmann S.; Mueller M. J.; Kijne J. W.; Memelink J. Involvement of the Octadecanoid Pathway and Protein Phosphorylation in Fungal Elicitor-Induced Expression of Terpenoid Indole Alkaloid Biosynthetic Genes in Catharanthus Roseus. Plant Physiol. 1999, 119 (4), 1289–1296. 10.1104/pp.119.4.1289. [DOI] [PMC free article] [PubMed] [Google Scholar]
Howe G. A. Cyclopentenone Signals for Plant Defense: Remodeling the Jasmonic Acid Response. Proc. Natl. Acad. Sci. U. S. A. 2001, 98 (22), 12317–12319. 10.1073/pnas.231480898. [DOI] [PMC free article] [PubMed] [Google Scholar]
Farmer E. E.; Ryan C. A. Octadecanoid Precursors of Jasmonic Acid Activate the Synthesis of Wound-Inducible Proteinase Inhibitors. Plant Cell 1992, 4 (2), 129–134. 10.1105/tpc.4.2.129. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref1] Chevreul M. E.; de Bouffe J.. Ou recueil des mémoires concernant la chimie et les arts qui en dépendent. Annales de Chimie 1813.881 [Google Scholar]

[ref2] Burdge G. C.; Calder P. C.. Introduction to Fatty Acids and Lipids. Intravenous Lipid Emulsions; World Review of Nutrition and Dietetics; Karger, 2015; Vol. 112, pp 1–16. 10.1159/000365423 [DOI] [PubMed] [Google Scholar]

[ref3] Benatti P.; Peluso G.; Nicolai R.; Calvani M. Polyunsaturated Fatty Acids: Biochemical, Nutritional and Epigenetic Properties. J. Am. Coll. Nutr. 2004, 23 (4), 281–302. 10.1080/07315724.2004.10719371. [DOI] [PubMed] [Google Scholar]

[ref4] Vance D. E.; Vance J. E.. Biochemistry of Lipids, Lipoproteins and Membranes; Elsevier Science, 2008. [Google Scholar]

[ref5] Burr G. O.; Burr M. M.; Miller E. S. On the Fatty Acids Essential in Nutrition. III. J. Biol. Chem. 1932, 97, 1–9. 10.1016/S0021-9258(18)76213-3. [DOI] [Google Scholar]

[ref6] Tocher D. R.; Leaver M. J.; Hodgson P. A. Recent Advances in the Biochemistry and Molecular Biology of Fatty Acyl Desaturases. Prog. Lipid Res. 1998, 37 (2), 73–117. 10.1016/S0163-7827(98)00005-8. [DOI] [PubMed] [Google Scholar]

[ref7] Holman R. T.; Mohrhauer H.; Porwit-Bobr Z.; Baptist J. N. A Hypothesis Involving Competitive Inhibitions in the Metabolism of Polyunsaturated Fatty Acids. Acta Chemica Scandinavia. 1963, S84–S90. 10.3891/acta.chem.scand.17s-0084. [DOI] [Google Scholar]

[ref8] Nakamura M. T.; Nara T. Y. Essential Fatty Acid Synthesis and Its Regulation in Mammals. Prostaglandins Leukot. Essent. Fatty Acids 2003, 68 (2), 145–150. 10.1016/S0952-3278(02)00264-8. [DOI] [PubMed] [Google Scholar]

[ref9] Cormier H.; Rudkowska I.; Lemieux S.; Couture P.; Julien P.; Vohl M.-C. Effects of FADS and ELOVL Polymorphisms on Indexes of Desaturase and Elongase Activities: Results from a Pre-Post Fish Oil Supplementation. Genes Nutr. 2014, 9 (6), 437. 10.1007/s12263-014-0437-z. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref10] Glick N. R.; Fischer M. H. The Role of Essential Fatty Acids in Human Health. J. Evid.-Based Complement. Altern. Med. 2013, 18 (4), 268–289. 10.1177/2156587213488788. [DOI] [Google Scholar]

[ref11] Gerwick W. H.; Moghaddam M.; Hamberg M. Oxylipin Metabolism in the Red Alga Gracilariopsis lemaneiformis: Mechanism of Formation of Vicinal Dihydroxy Fatty Acids. Arch. Biochem. Biophys. 1991, 290 (2), 436–444. 10.1016/0003-9861(91)90563-X. [DOI] [PubMed] [Google Scholar]

[ref12] Dyall S. C.; Balas L.; Bazan N. G.; Brenna J. T.; Chiang N.; da Costa Souza F.; Dalli J.; Durand T.; Galano J.-M.; Lein P. J.; Serhan C. N.; Taha A. Y. Polyunsaturated Fatty Acids and Fatty Acid-Derived Lipid Mediators: Recent Advances in the Understanding of Their Biosynthesis, Structures, and Functions. Prog. Lipid Res. 2022, 86, 101165. 10.1016/j.plipres.2022.101165. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref13] Harizi H.; Corcuff J.-B.; Gualde N. Arachidonic-Acid-Derived Eicosanoids: Roles in Biology and Immunopathology. Trends Mol. Med. 2008, 14 (10), 461–469. 10.1016/j.molmed.2008.08.005. [DOI] [PubMed] [Google Scholar]

[ref14] Calder P. C. Eicosanoids. Essays Biochem. 2020, 64 (3), 423–441. 10.1042/EBC20190083. [DOI] [PubMed] [Google Scholar]

[ref15] Samuelsson B.; Dahlén S.-E.; Lindgren J. Å.; Rouzer C. A.; Serhan C. N. Leukotrienes and Lipoxins: Structures, Biosynthesis, and Biological Effects. Science 1987, 237 (4819), 1171–1176. 10.1126/science.2820055. [DOI] [PubMed] [Google Scholar]

[ref16] Tourdot B.; Ahmed I.; Holinstat M. The Emerging Role of Oxylipins in Thrombosis and Diabetes. Front. Pharmacol. 2014, 4, 176. 10.3389/fphar.2013.00176. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref17] Misheva M.; Johnson J.; McCullagh J. Role of Oxylipins in the Inflammatory-Related Diseases NAFLD, Obesity, and Type 2 Diabetes. Metabolites 2022, 12 (12), 1238. 10.3390/metabo12121238. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref18] Barquissau V.; Ghandour R. A.; Ailhaud G.; Klingenspor M.; Langin D.; Amri E.-Z.; Pisani D. F. Control of Adipogenesis by Oxylipins, GPCRs and PPARs. Biochimie 2017, 136, 3–11. 10.1016/j.biochi.2016.12.012. [DOI] [PubMed] [Google Scholar]

[ref19] Liput K. P.; Lepczyński A.; Ogłuszka M.; Nawrocka A.; Poławska E.; Grzesiak A.; Ślaska B.; Pareek C. S.; Czarnik U.; Pierzchała M. Effects of Dietary n-3 and n-6 Polyunsaturated Fatty Acids in Inflammation and Cancerogenesis. Int. J. Mol. Sci. 2021, 22 (13), 6965. 10.3390/ijms22136965. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref20] Edin M. L.; Duval C.; Zhang G.; Zeldin D. C. Role of Linoleic Acid-Derived Oxylipins in Cancer. Cancer Metastasis Rev. 2020, 39 (3), 581–582. 10.1007/s10555-020-09904-8. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref21] Lundstrom S. L.; Balgoma D.; Wheelock A. M.; Haeggstrom J. Z.; Dahlen S.-E.; Wheelock C. E. Lipid Mediator Profiling in Pulmonary Disease. Curr. Pharm. Biotechnol. 2011, 12, 1026–1052. 10.2174/138920111795909087. [DOI] [PubMed] [Google Scholar]

[ref22] Swardfager W.; Hennebelle M.; Yu D.; Hammock B.; Levitt A.; Hashimoto K.; Taha A. Metabolic/Inflammatory/Vascular Comorbidity in Psychiatric Disorders; Soluble Epoxide Hydrolase (SEH) as a Possible New Target. Neurosci. Biobehav. Rev. 2018, 87, 56–66. 10.1016/j.neubiorev.2018.01.010. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref23] Duan J.; Song Y.; Zhang X.; Wang C. Effect of ω-3 Polyunsaturated Fatty Acids-Derived Bioactive Lipids on Metabolic Disorders. Front. Physiol. 2021, 12, 646491. 10.3389/fphys.2021.646491. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref24] Guimarães R. C.; Gonçalves T. T.; Leiria L. O. Exploiting Oxidized Lipids and the Lipid-Binding GPCRs against Cardiometabolic Diseases. Br. J. Pharmacol. 2021, 178 (3), 531–549. 10.1111/bph.15321. [DOI] [PubMed] [Google Scholar]

[ref25] Nayeem M. A. Role of Oxylipins in Cardiovascular Diseases. Acta Pharmacol. Sin. 2018, 39 (7), 1142–1154. 10.1038/aps.2018.24. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref26] He J.; Ma C.; Tang D.; Zhong S.; Yuan X.; Zheng F.; Zeng Z.; Chen Y.; Liu D.; Hong X.; Dai W.; Yin L.; Dai Y. Absolute Quantification and Characterization of Oxylipins in Lupus Nephritis and Systemic Lupus Erythematosus. Front. Immunol. 2022, 13, 964901. 10.3389/fimmu.2022.964901. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref27] Khanapure S. P.; Garvey D. S.; Janero D. R.; Letts L. G. Eicosanoids in Inflammation: Biosynthesis, Pharmacology, and Therapeutic Frontiers. Curr. Top. Med. Chem. 2007, 7, 311–340. 10.2174/156802607779941314. [DOI] [PubMed] [Google Scholar]

[ref28] Wang B.; Wu L.; Chen J.; Dong L.; Chen C.; Wen Z.; Hu J.; Fleming I.; Wang D. W. Metabolism Pathways of Arachidonic Acids: Mechanisms and Potential Therapeutic Targets. Signal Transduct. Target. Ther. 2021, 6 (1), 94. 10.1038/s41392-020-00443-w. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref29] Bazan N. G. Docosanoids and Elovanoids from Omega-3 Fatty Acids Are pro-Homeostatic Modulators of Inflammatory Responses, Cell Damage and Neuroprotection. Mol. Aspects Med. 2018, 64, 18–33. 10.1016/j.mam.2018.09.003. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref30] Serhan C. N. Novel Eicosanoid and Docosanoid Mediators: Resolvins, Docosatrienes, and Neuroprotectins. Curr. Opin. Clin. Nutr. Metab. Care 2005, 8 (2), 115–121. 10.1097/00075197-200503000-00003. [DOI] [PubMed] [Google Scholar]

[ref31] Hajeyah A. A.; Griffiths W. J.; Wang Y.; Finch A. J.; O’Donnell V. B. The Biosynthesis of Enzymatically Oxidized Lipids. Front. Endocrinol. 2020, 11, 591819. 10.3389/fendo.2020.591819. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref32] Gabbs M.; Leng S.; Devassy J. G.; Monirujjaman M.; Aukema H. M. Advances in Our Understanding of Oxylipins Derived from Dietary PUFAs. Adv. Nutr. 2015, 6 (5), 513–540. 10.3945/an.114.007732. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref33] Christie W. W.; Harwood J. L. Oxidation of Polyunsaturated Fatty Acids to Produce Lipid Mediators. Essays Biochem. 2020, 64 (3), 401–421. 10.1042/EBC20190082. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref34] Dennis E. A.; Norris P. C. Eicosanoid Storm in Infection and Inflammation. Nat. Rev. Immunol. 2015, 15 (8), 511–523. 10.1038/nri3859. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref35] Serhan C. N. Pro-Resolving Lipid Mediators Are Leads for Resolution Physiology. Nature 2014, 510 (7503), 92–101. 10.1038/nature13479. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref36] Chiang N.; Serhan C. N. Specialized Pro-Resolving Mediator Network: An Update on Production and Actions. Essays Biochem. 2020, 64 (3), 443–462. 10.1042/EBC20200018. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref37] Calder P. C. Eicosapentaenoic and Docosahexaenoic Acid Derived Specialised Pro-Resolving Mediators: Concentrations in Humans and the Effects of Age, Sex, Disease and Increased Omega-3 Fatty Acid Intake. Biochimie 2020, 178, 105–123. 10.1016/j.biochi.2020.08.015. [DOI] [PubMed] [Google Scholar]

[ref38] Vidar Hansen T.; Serhan C. N. Protectins: Their Biosynthesis, Metabolism and Structure-Functions. Biochem. Pharmacol. 2022, 206, 115330. 10.1016/j.bcp.2022.115330. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref39] Schebb N. H.; Kühn H.; Kahnt A. S.; Rund K. M.; O’Donnell V. B.; Flamand N.; Peters-Golden M.; Jakobsson P.-J.; Weylandt K. H.; Rohwer N.; Murphy R. C.; Geisslinger G.; FitzGerald G. A.; Hanson J.; Dahlgren C.; Alnouri M. W.; Offermanns S.; Steinhilber D. Formation, Signaling and Occurrence of Specialized Pro-Resolving Lipid Mediators—What Is the Evidence so Far?. Front. Pharmacol. 2022, 13, 838782. 10.3389/fphar.2022.838782. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref40] Andreou A.; Brodhun F.; Feussner I. Biosynthesis of Oxylipins in Non-Mammals. Prog. Lipid Res. 2009, 48 (3–4), 148–170. 10.1016/j.plipres.2009.02.002. [DOI] [PubMed] [Google Scholar]

[ref41] Parchem K.; Letsiou S.; Petan T.; Oskolkova O.; Medina I.; Kuda O.; O’Donnell V. B.; Nicolaou A.; Fedorova M.; Bochkov V.; Gladine C. Oxylipin Profiling for Clinical Research: Current Status and Future Perspectives. Prog. Lipid Res. 2024, 95, 101276. 10.1016/j.plipres.2024.101276. [DOI] [PubMed] [Google Scholar]

[ref42] Schaller F. Enzymes of the Biosynthesis of Octadecanoid-Derived Signalling Molecules. J. Exp. Bot. 2001, 52 (354), 11–23. 10.1093/jexbot/52.354.11. [DOI] [PubMed] [Google Scholar]

[ref43] Agrawal G. K.; Tamogami S.; Han O.; Iwahashi H.; Rakwal R. Rice Octadecanoid Pathway. Biochem. Biophys. Res. Commun. 2004, 317 (1), 1–15. 10.1016/j.bbrc.2004.03.020. [DOI] [PubMed] [Google Scholar]

[ref44] Carlson S. E.; Carver J. D.; House S. G. High Fat Diets Varying in Ratios of Polyunsaturated to Saturated Fatty Acid and Linoleic to Linolenic Acid: A Comparison of Rat Neural and Red Cell Membrane Phospholipids. J. Nutr. 1986, 116 (5), 718–725. 10.1093/jn/116.5.718. [DOI] [PubMed] [Google Scholar]

[ref45] Whelan J.; Fritsche K. Linoleic Acid. Adv. Nutr. 2013, 4 (3), 311–312. 10.3945/an.113.003772. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref46] Hutchins P. M.; Murphy R. C. Cholesteryl Ester Acyl Oxidation and Remodeling in Murine Macrophages: Formation of Oxidized Phosphatidylcholine. J. Lipid Res. 2012, 53 (8), 1588–1597. 10.1194/jlr.M026799. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref47] Liu G.-Y.; Moon S. H.; Jenkins C. M.; Sims H. F.; Guan S.; Gross R. W. Synthesis of Oxidized Phospholipids by Sn-1 Acyltransferase Using 2–15-HETE Lysophospholipids. J. Biol. Chem. 2019, 294 (26), 10146–10159. 10.1074/jbc.RA119.008766. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref48] Burke J. E.; Dennis E. A. Phospholipase A2 Biochemistry. Cardiovasc. Drugs Ther. 2009, 23 (1), 49–59. 10.1007/s10557-008-6132-9. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref49] Wang L.; Gill R.; Pedersen T. L.; Higgins L. J.; Newman J. W.; Rutledge J. C. Triglyceride-Rich Lipoprotein Lipolysis Releases Neutral and Oxidized FFAs That Induce Endothelial Cell Inflammation. J. Lipid Res. 2009, 50 (2), 204–213. 10.1194/jlr.M700505-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref50] Dennis E. A.; Cao J.; Hsu Y.-H.; Magrioti V.; Kokotos G. Phospholipase A2 Enzymes: Physical Structure, Biological Function, Disease Implication, Chemical Inhibition, and Therapeutic Intervention. Chem. Rev. 2011, 111 (10), 6130–6185. 10.1021/cr200085w. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref51] Blasbalg T. L.; Hibbeln J. R.; Ramsden C. E.; Majchrzak S. F.; Rawlings R. R. Changes in Consumption of Omega-3 and Omega-6 Fatty Acids in the United States during the 20th Century. Am. J. Clin. Nutr. 2011, 93 (5), 950–962. 10.3945/ajcn.110.006643. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref52] Wasternack C. Jasmonates: An Update on Biosynthesis, Signal Transduction and Action in Plant Stress Response, Growth and Development. Ann. Bot. 2007, 100 (4), 681–697. 10.1093/aob/mcm079. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref53] Feussner I.; Wasternack C. The Lipoxygenase Pathway. Annu. Rev. Plant Biol. 2002, 53 (1), 275–297. 10.1146/annurev.arplant.53.100301.135248. [DOI] [PubMed] [Google Scholar]

[ref54] Mosblech A.; Feussner I.; Heilmann I. Oxylipins: Structurally Diverse Metabolites from Fatty Acid Oxidation. Plant Physiol. Biochem. 2009, 47 (6), 511–517. 10.1016/j.plaphy.2008.12.011. [DOI] [PubMed] [Google Scholar]

[ref55] Sugiyama S.; Hayakawa M.; Nagai S.; Ajioka M.; Ozawa T. Leukotoxin, 9, 10-Epoxy-12-Octadecenoate, Causes Cardiac Failure in Dogs. Life Sci. 1987, 40 (3), 225–231. 10.1016/0024-3205(87)90336-5. [DOI] [PubMed] [Google Scholar]

[ref56] Greene J. F.; Hammock B. D. Toxicity of Linoleic Acid Metabolites. Adv. Exp. Med. Biol. 1999, 469, 471–477. 10.1007/978-1-4615-4793-8_69. [DOI] [PubMed] [Google Scholar]

[ref57] Newman J. W.; Morisseau C.; Hammock B. D. Epoxide Hydrolases: Their Roles and Interactions with Lipid Metabolism. Prog. Lipid Res. 2005, 44 (1), 1–51. 10.1016/j.plipres.2004.10.001. [DOI] [PubMed] [Google Scholar]

[ref58] Hildreth K.; Kodani S. D.; Hammock B. D.; Zhao L. Cytochrome P450-Derived Linoleic Acid Metabolites EpOMEs and DiHOMEs: A Review of Recent Studies. J. Nutr. Biochem. 2020, 86, 108484. 10.1016/j.jnutbio.2020.108484. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref59] Chiba T.; Thomas C. P.; Calcutt M. W.; Boeglin W. E.; O’Donnell V. B.; Brash A. R. The Precise Structures and Stereochemistry of Trihydroxy-Linoleates Esterified in Human and Porcine Epidermis and Their Significance in Skin Barrier Function. J. Biol. Chem. 2016, 291 (28), 14540–14554. 10.1074/jbc.M115.711267. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref60] Ramsden C. E.; Ringel A.; Majchrzak-Hong S. F.; Yang J.; Blanchard H.; Zamora D.; Loewke J. D.; Rapoport S. I.; Hibbeln J. R.; Davis J. M.; Hammock B. D.; Taha A. Y. Dietary Linoleic Acid-Induced Alterations in pro- and Anti-Nociceptive Lipid Autacoids: Implications for Idiopathic Pain Syndromes?. Mol. Pain 2016, 12, 174480691663638. 10.1177/1744806916636386. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref61] Ramsden C. E.; Domenichiello A. F.; Yuan Z.-X.; Sapio M. R.; Keyes G. S.; Mishra S. K.; Gross J. R.; Majchrzak-Hong S.; Zamora D.; Horowitz M. S.; Davis J. M.; Sorokin A. V.; Dey A.; LaPaglia D. M.; Wheeler J. J.; Vasko M. R.; Mehta N. N.; Mannes A. J.; Iadarola M. J. A Systems Approach for Discovering Linoleic Acid Derivatives That Potentially Mediate Pain and Itch. Sci. Signal. 2017, 10 (493), eaal5241 10.1126/scisignal.aal5241. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref62] Domenichiello A. F.; Sapio M. R.; Loydpierson A. J.; Maric D.; Goto T.; Horowitz M. S.; Keyes G. S.; Yuan Z.-X.; Majchrzak-Hong S. F.; Mannes A. J.; Iadarola M. J.; Ramsden C. E. Molecular Pathways Linking Oxylipins to Nociception in Rats. J. Pain 2021, 22, 275–299. 10.1016/j.jpain.2020.09.001. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref63] Stanford K. I.; Lynes M. D.; Takahashi H.; Baer L. A.; Arts P. J.; May F. J.; Lehnig A. C.; Middelbeek R. J. W.; Richard J. J.; So K.; Chen E. Y.; Gao F.; Narain N. R.; Distefano G.; Shettigar V. K.; Hirshman M. F.; Ziolo M. T.; Kiebish M. A.; Tseng Y.-H.; Coen P. M.; Goodyear L. J. 12,13-DiHOME: An Exercise-Induced Lipokine That Increases Skeletal Muscle Fatty Acid Uptake. Cell Metab. 2018, 27 (5), 1111–1120. 10.1016/j.cmet.2018.03.020. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref64] Lynes M. D.; Leiria L. O.; Lundh M.; Bartelt A.; Shamsi F.; Huang T. L.; Takahashi H.; Hirshman M. F.; Schlein C.; Lee A.; Baer L. A.; May F. J.; Gao F.; Narain N. R.; Chen E. Y.; Kiebish M. A.; Cypess A. M.; Blüher M.; Goodyear L. J.; Hotamisligil G. S.; Stanford K. I.; Tseng Y.-H. The Cold-Induced Lipokine 12,13-DiHOME Promotes Fatty Acid Transport into Brown Adipose Tissue. Nat. Med. 2017, 23 (5), 631–637. 10.1038/nm.4297. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref65] Cabral M.; Martín-Venegas R.; Moreno J. J. Differential Cell Growth/Apoptosis Behavior of 13-Hydroxyoctadecadienoic Acid Enantiomers in a Colorectal Cancer Cell Line. Am. J. Physiol.-Gastrointest. Liver Physiol. 2014, 307 (6), G664–G671. 10.1152/ajpgi.00064.2014. [DOI] [PubMed] [Google Scholar]

[ref66] Vangaveti V. N.; Jansen H.; Kennedy R. L.; Malabu U. H. Hydroxyoctadecadienoic Acids: Oxidised Derivatives of Linoleic Acid and Their Role in Inflammation Associated with Metabolic Syndrome and Cancer. Eur. J. Pharmacol. 2016, 785, 70–76. 10.1016/j.ejphar.2015.03.096. [DOI] [PubMed] [Google Scholar]

[ref67] Kumar N.; Gupta G.; Anilkumar K.; Fatima N.; Karnati R.; Reddy G. V.; Giri P. V.; Reddanna P. 15-Lipoxygenase Metabolites of α-Linolenic Acid, [13-(S)-HPOTrE and 13-(S)-HOTrE], Mediate Anti-Inflammatory Effects by Inactivating NLRP3 Inflammasome. Sci. Rep. 2016, 6 (1), 31649. 10.1038/srep31649. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref68] Bergmann C. B.; McReynolds C. B.; Wan D.; Singh N.; Goetzman H.; Caldwell C. C.; Supp D. M.; Hammock B. D. SEH-Derived Metabolites of Linoleic Acid Drive Pathologic Inflammation While Impairing Key Innate Immune Cell Function in Burn Injury. Proc. Natl. Acad. Sci. U. S. A. 2022, 119 (13), e2120691119 10.1073/pnas.2120691119. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref69] Gilroy D. W.; Edin M. L.; De Maeyer R. P. H.; Bystrom J.; Newson J.; Lih F. B.; Stables M.; Zeldin D. C.; Bishop-Bailey D. CYP450-Derived Oxylipins Mediate Inflammatory Resolution. Proc. Natl. Acad. Sci. U. S. A. 2016, 113 (23), E3240. 10.1073/pnas.1521453113. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref70] Fujimura K. E.; Sitarik A. R.; Havstad S.; Lin D. L.; Levan S.; Fadrosh D.; Panzer A. R.; LaMere B.; Rackaityte E.; Lukacs N. W.; Wegienka G.; Boushey H. A.; Ownby D. R.; Zoratti E. M.; Levin A. M.; Johnson C. C.; Lynch S. V. Neonatal Gut Microbiota Associates with Childhood Multisensitized Atopy and T Cell Differentiation. Nat. Med. 2016, 22 (10), 1187–1191. 10.1038/nm.4176. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref71] Levan S. R.; Stamnes K. A.; Lin D. L.; Panzer A. R.; Fukui E.; McCauley K.; Fujimura K. E.; McKean M.; Ownby D. R.; Zoratti E. M.; Boushey H. A.; Cabana M. D.; Johnson C. C.; Lynch S. V. Elevated Faecal 12,13-DiHOME Concentration in Neonates at High Risk for Asthma Is Produced by Gut Bacteria and Impedes Immune Tolerance. Nat. Microbiol. 2019, 4 (11), 1851–1861. 10.1038/s41564-019-0498-2. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref72] Bell M. V.; Dick J. R.; Pond D. W. Octadecapentaenoic Acid in a Raphidophyte Alga, Heterosigma akashiwo. Phytochemistry 1997, 45 (2), 303–306. 10.1016/S0031-9422(96)00867-9. [DOI] [Google Scholar]

[ref73] Trostchansky A.; Bonilla L.; González-Perilli L.; Rubbo H. Nitro-Fatty Acids: Formation, Redox Signaling, and Therapeutic Potential. Antioxid. Redox Signal. 2013, 19 (11), 1257–1265. 10.1089/ars.2012.5023. [DOI] [PubMed] [Google Scholar]

[ref74] Mata-Pérez C.; Sánchez-Calvo B.; Padilla M. N.; Begara-Morales J. C.; Valderrama R.; Corpas F. J.; Barroso J. B. Nitro-Fatty Acids in Plant Signaling: New Key Mediators of Nitric Oxide Metabolism. Redox Biol. 2017, 11, 554–561. 10.1016/j.redox.2017.01.002. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref75] Delmastro-Greenwood M.; Freeman B. A.; Wendell S. G. Redox-Dependent Anti-Inflammatory Signaling Actions of Unsaturated Fatty Acids. Annu. Rev. Physiol. 2014, 76 (1), 79–105. 10.1146/annurev-physiol-021113-170341. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref76] Schopfer F. J.; Cipollina C.; Freeman B. A. Formation and Signaling Actions of Electrophilic Lipids. Chem. Rev. 2011, 111 (10), 5997–6021. 10.1021/cr200131e. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref77] Quaranta A.; Revol-Cavalier J.; Wheelock C. E. The Octadecanoids: An Emerging Class of Lipid Mediators. Biochem. Soc. Trans. 2022, 50 (6), 1569–1582. 10.1042/BST20210644. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref78] Yamada Y.; Uemura H.; Nakaya H.; Sakata K.; Takatori T.; Nagao M.; Iwase H.; Iwadate K. Production of Hydroxy Fatty Acid (10-Hydroxy-12(Z)-Octadecenoic Acid) by Lactobacillus plantarum from Linoleic Acid and Its Cardiac Effects to Guinea Pig Papillary Muscles. Biochem. Biophys. Res. Commun. 1996, 226 (2), 391–395. 10.1006/bbrc.1996.1366. [DOI] [PubMed] [Google Scholar]

[ref79] Miyamoto J.; Igarashi M.; Watanabe K.; Karaki S.; Mukouyama H.; Kishino S.; Li X.; Ichimura A.; Irie J.; Sugimoto Y.; Mizutani T.; Sugawara T.; Miki T.; Ogawa J.; Drucker D. J.; Arita M.; Itoh H.; Kimura I. Gut Microbiota Confers Host Resistance to Obesity by Metabolizing Dietary Polyunsaturated Fatty Acids. Nat. Commun. 2019, 10 (1), 4007. 10.1038/s41467-019-11978-0. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref80] Tang L.; Shang J.; Song C.; Yang R.; Shang X.; Mao W.; Bao D.; Tan Q. Untargeted Metabolite Profiling of Antimicrobial Compounds in the Brown Film of Lentinula edodes Mycelium via LC-MS/MS Analysis. ACS Omega 2020, 5 (13), 7567–7575. 10.1021/acsomega.0c00398. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref81] Takahashi H.; Kamakari K.; Goto T.; Hara H.; Mohri S.; Suzuki H.; Shibata D.; Nakata R.; Inoue H.; Takahashi N.; Kawada T. 9-Oxo-10(E),12(Z),15(Z)-Octadecatrienoic Acid Activates Peroxisome Proliferator-Activated Receptor α in Hepatocytes. Lipids 2015, 50 (11), 1083–1091. 10.1007/s11745-015-4071-3. [DOI] [PubMed] [Google Scholar]

[ref82] Petta T.; Secatto A.; Faccioli L. H.; Moraes L. A. B. Inhibition of Inflammatory Response in LPS Induced Macrophages by 9-KOTE and 13-KOTE Produced by Biotransformation. Enzyme Microb. Technol. 2014, 58–59, 36–43. 10.1016/j.enzmictec.2014.02.011. [DOI] [PubMed] [Google Scholar]

[ref83] Fahy E.; Subramaniam S.; Murphy R. C.; Nishijima M.; Raetz C. R. H.; Shimizu T.; Spener F.; van Meer G.; Wakelam M. J. O.; Dennis E. A. Update of the LIPID MAPS Comprehensive Classification System for Lipids. J. Lipid Res. 2009, 50, S9–S14. 10.1194/jlr.R800095-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref84] Sud M.; Fahy E.; Cotter D.; Brown A.; Dennis E. A.; Glass C. K.; Merrill A. H. Jr; Murphy R. C.; Raetz C. R. H.; Russell D. W.; Subramaniam S. LMSD: LIPID MAPS Structure Database. Nucleic Acids Res. 2007, 35, D527–D532. 10.1093/nar/gkl838. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref85] Wishart D. S.; Guo A.; Oler E.; Wang F.; Anjum A.; Peters H.; Dizon R.; Sayeeda Z.; Tian S.; Lee B. L.; Berjanskii M.; Mah R.; Yamamoto M.; Jovel J.; Torres-Calzada C.; Hiebert-Giesbrecht M.; Lui V. W.; Varshavi D.; Varshavi D.; Allen D.; Arndt D.; Khetarpal N.; Sivakumaran A.; Harford K.; Sanford S.; Yee K.; Cao X.; Budinski Z.; Liigand J.; Zhang L.; Zheng J.; Mandal R.; Karu N.; Dambrova M.; Schiöth H. B.; Greiner R.; Gautam V. HMDB 5.0: The Human Metabolome Database for 2022. Nucleic Acids Res. 2022, 50 (D1), D622–D631. 10.1093/nar/gkab1062. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref86] Smith D. L.; Willis A. L. A Suggested Shorthand Nomenclature for the Eicosanoids. Lipids 1987, 22 (12), 983–986. 10.1007/BF02536436. [DOI] [PubMed] [Google Scholar]

[ref87] Taber D. F.; Morrow J. D.; Roberts L. J II. A Nomenclature System for the Isoprostanes. Prostaglandins 1997, 53, 63–67. 10.1016/S0090-6980(97)00005-1. [DOI] [PubMed] [Google Scholar]

[ref88] Taber D. F.; Roberts L. J. Nomenclature Systems for the Neuroprostanes and for the Neurofurans. Prostaglandins Other Lipid Mediat. 2005, 78 (1–4), 14–18. 10.1016/j.prostaglandins.2005.07.002. [DOI] [PubMed] [Google Scholar]

[ref89] Taber D. F.; Fessel J. P.; Roberts L. J. A Nomenclature System for Isofurans. Prostaglandins Other Lipid Mediat. 2004, 73 (1), 47–50. 10.1016/j.prostaglandins.2003.11.004. [DOI] [PubMed] [Google Scholar]

[ref90] Vick B. A.; Zimmerman D. C. Biosynthesis of Jasmonic Acid by Several Plant Species. Plant Physiol. 1984, 75 (2), 458–461. 10.1104/pp.75.2.458. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref91] Grechkin A. N.; Hamberg M. The “Heterolytic Hydroperoxide Lyase” Is an Isomerase Producing a Short-Lived Fatty Acid Hemiacetal. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2004, 1636 (1), 47–58. 10.1016/j.bbalip.2003.12.003. [DOI] [PubMed] [Google Scholar]

[ref92] Hamberg M. A Pathway for Biosynthesis of Divinyl Ether Fatty Acids in Green Leaves. Lipids 1998, 33 (11), 1061–1071. 10.1007/s11745-998-0306-7. [DOI] [PubMed] [Google Scholar]

[ref93] Toporkova Y. Y.; Smirnova E. O.; Lantsova N. V.; Mukhtarova L. S.; Grechkin A. N. Detection of the First Epoxyalcohol Synthase/Allene Oxide Synthase (CYP74 Clan) in the Lancelet (Branchiostoma belcheri, Chordata). Int. J. Mol. Sci. 2021, 22 (9), 4737. 10.3390/ijms22094737. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref94] Lee M.; Lenman M.; Banaś A.; Bafor M.; Singh S.; Schweizer M.; Nilsson R.; Liljenberg C.; Dahlqvist A.; Gummeson P.-O.; Sjödahl S.; Green A.; Stymne S. Identification of Non-Heme Diiron Proteins That Catalyze Triple Bond and Epoxy Group Formation. Science 1998, 280 (5365), 915–918. 10.1126/science.280.5365.915. [DOI] [PubMed] [Google Scholar]

[ref95] Laneuville O.; Breuer D. K.; Xu N.; Huang Z. H.; Gage D. A.; Watson J. T.; Lagarde M.; DeWitt D. L.; Smith W. L. Fatty Acid Substrate Specificities of Human Prostaglandin-Endoperoxide H Synthase-1 and – 2: FORMATION OF 12-HYDROXY-(9Z,13E/Z,15Z)-OCTADECATRIENOIC ACIDS FROM α-LINOLENIC ACID. J. Biol. Chem. 1995, 270 (33), 19330–19336. 10.1074/jbc.270.33.19330. [DOI] [PubMed] [Google Scholar]

[ref96] Brash A. R.; Boeglin W. E.; Chang M. S. Discovery of a Second 15S-Lipoxygenase in Humans. Proc. Natl. Acad. Sci. U. S. A. 1997, 94 (12), 6148–6152. 10.1073/pnas.94.12.6148. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref97] Bylund J.; Ericsson J.; Oliw E. H. Analysis of Cytochrome P450 Metabolites of Arachidonic and Linoleic Acids by Liquid Chromatography-Mass Spectrometry with Ion Trap MS2. Anal. Biochem. 1998, 265 (1), 55–68. 10.1006/abio.1998.2897. [DOI] [PubMed] [Google Scholar]

[ref98] Oliw E. H.; Bylund J.; Herman C. Bisallylic Hydroxylation and Epoxidation of Polyunsaturated Fatty Acids by Cytochrome P450. Lipids 1996, 31 (10), 1003–1021. 10.1007/BF02522457. [DOI] [PubMed] [Google Scholar]

[ref99] Schuchardt J. P.; Schmidt S.; Kressel G.; Dong H.; Willenberg I.; Hammock B. D.; Hahn A.; Schebb N. H. Comparison of Free Serum Oxylipin Concentrations in Hyper- vs. Normolipidemic Men. Prostaglandins Leukot. Essent. Fatty Acids 2013, 89 (1), 19–29. 10.1016/j.plefa.2013.04.001. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref100] Yu Z.; Schneider C.; Boeglin W. E.; Marnett L. J.; Brash A. R. The Lipoxygenase Gene ALOXE3 Implicated in Skin Differentiation Encodes a Hydroperoxide Isomerase. Proc. Natl. Acad. Sci. U. S. A. 2003, 100 (16), 9162–9167. 10.1073/pnas.1633612100. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref101] Liu M.; Chen P.; Véricel E.; Lelli M.; Béguin L.; Lagarde M.; Guichardant M. Characterization and Biological Effects of Di-Hydroxylated Compounds Deriving from the Lipoxygenation of ALA. J. Lipid Res. 2013, 54 (8), 2083–2094. 10.1194/jlr.M035139. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref102] Jahn U.; Galano J.-M.; Durand T. Beyond Prostaglandins—Chemistry and Biology of Cyclic Oxygenated Metabolites Formed by Free-Radical Pathways from Polyunsaturated Fatty Acids. Angew. Chem., Int. Ed. 2008, 47 (32), 5894–5955. 10.1002/anie.200705122. [DOI] [PubMed] [Google Scholar]

[ref103] Leung K. S.; Oger C.; Guy A.; Bultel-Poncé V.; Vigor C.; Durand T.; Gil-Izquierdo A.; Medina S.; Galano J.-M.; Lee J. C.-Y.. Alpha-Linolenic Acid, Phytoprostanes and Phytofurans in Plant, Algae and Food. In Lipids in Plants and Algae: From Fundamental Science to Industrial Applications; Advances in Botanical Research; Elsevier, 2022; Vol. 101, p 437. 10.1016/bs.abr.2021.09.005. [DOI] [Google Scholar]

[ref104] Mata-Pérez C.; Sánchez-Calvo B.; Padilla M. N.; Begara-Morales J. C.; Luque F.; Melguizo M.; Jiménez-Ruiz J.; Fierro-Risco J.; Peñas-Sanjuán A.; Valderrama R.; Corpas F. J.; Barroso J. B. Nitro-Fatty Acids in Plant Signaling: Nitro-Linolenic Acid Induces the Molecular Chaperone Network in Arabidopsis. Plant Physiol. 2016, 170 (2), 686–701. 10.1104/pp.15.01671. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref105] Koutoulogenis G. S.; Kokotos G. Nitro Fatty Acids (NO₂-FAs): An Emerging Class of Bioactive Fatty Acids. Molecules 2021, 26 (24), 7536. 10.3390/molecules26247536. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref106] Diczfalusy U. Beta-Oxidation of Eicosanoids. Prog. Lipid Res. 1994, 33 (4), 403–428. 10.1016/0163-7827(94)90025-6. [DOI] [PubMed] [Google Scholar]

[ref107] Misheva M.; Kotzamanis K.; Davies L. C.; Tyrrell V. J.; Rodrigues P. R. S.; Benavides G. A.; Hinz C.; Murphy R. C.; Kennedy P.; Taylor P. R.; Rosas M.; Jones S. A.; McLaren J. E.; Deshpande S.; Andrews R.; Schebb N. H.; Czubala M. A.; Gurney M.; Aldrovandi M.; Meckelmann S. W.; Ghazal P.; Darley-Usmar V.; White D. A.; O’Donnell V. B. Oxylipin Metabolism Is Controlled by Mitochondrial β-Oxidation during Bacterial Inflammation. Nat. Commun. 2022, 13 (1), 139. 10.1038/s41467-021-27766-8. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref108] Whelan J. The Health Implications of Changing Linoleic Acid Intakes. Prostaglandins Leukot. Essent. Fatty Acids 2008, 79 (3), 165–167. 10.1016/j.plefa.2008.09.013. [DOI] [PubMed] [Google Scholar]

[ref109] Montani J.-P. Ancel Keys: The Legacy of a Giant in Physiology, Nutrition, and Public Health. Obes. Rev. 2021, 22, e13196 10.1111/obr.13196. [DOI] [PubMed] [Google Scholar]

[ref110] Mercola J.; D’Adamo C. R. Linoleic Acid: A Narrative Review of the Effects of Increased Intake in the Standard American Diet and Associations with Chronic Disease. Nutrients 2023, 15 (14), 3129. 10.3390/nu15143129. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref111] Marangoni F.; Agostoni C.; Borghi C.; Catapano A. L.; Cena H.; Ghiselli A.; La Vecchia C.; Lercker G.; Manzato E.; Pirillo A.; Riccardi G.; Risé P.; Visioli F.; Poli A. Dietary Linoleic Acid and Human Health: Focus on Cardiovascular and Cardiometabolic Effects. Atherosclerosis 2020, 292, 90–98. 10.1016/j.atherosclerosis.2019.11.018. [DOI] [PubMed] [Google Scholar]

[ref112] Li J.; Guasch-Ferré M.; Li Y.; Hu F. B. Dietary Intake and Biomarkers of Linoleic Acid and Mortality: Systematic Review and Meta-Analysis of Prospective Cohort Studies. Am. J. Clin. Nutr. 2020, 112 (1), 150–167. 10.1093/ajcn/nqz349. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref113] Taha A. Y. Linoleic Acid-Good or Bad for the Brain?. NPJ. Sci. Food 2020, 4, 1. 10.1038/s41538-019-0061-9. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref114] Lankinen M. A.; Fauland A.; Shimizu B.-I.; Ågren J.; Wheelock C. E.; Laakso M.; Schwab U.; Pihlajamäki J. Inflammatory Response to Dietary Linoleic Acid Depends on FADS1 Genotype. Am. J. Clin. Nutr. 2019, 109 (1), 165–175. 10.1093/ajcn/nqy287. [DOI] [PubMed] [Google Scholar]

[ref115] Ramsden C. E.; Hennebelle M.; Schuster S.; Keyes G. S.; Johnson C. D.; Kirpich I. A.; Dahlen J. E.; Horowitz M. S.; Zamora D.; Feldstein A. E.; McClain C. J.; Muhlhausler B. S.; Makrides M.; Gibson R. A.; Taha A. Y. Effects of Diets Enriched in Linoleic Acid and Its Peroxidation Products on Brain Fatty Acids, Oxylipins, and Aldehydes in Mice. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2018, 1863 (10), 1206–1213. 10.1016/j.bbalip.2018.07.007. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref116] Ramsden C. E.; Ringel A.; Feldstein A. E.; Taha A. Y.; MacIntosh B. A.; Hibbeln J. R.; Majchrzak-Hong S. F.; Faurot K. R.; Rapoport S. I.; Cheon Y.; Chung Y.-M.; Berk M.; Mann J. D. Lowering Dietary Linoleic Acid Reduces Bioactive Oxidized Linoleic Acid Metabolites in Humans. Prostaglandins Leukot. Essent. Fatty Acids 2012, 87, 135–141. 10.1016/j.plefa.2012.08.004. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref117] Liang N.; Hennebelle M.; Gaul S.; Johnson C. D.; Zhang Z.; Kirpich I. A.; McClain C. J.; Feldstein A. E.; Ramsden C. E.; Taha A. Y. Feeding Mice a Diet High in Oxidized Linoleic Acid Metabolites Does Not Alter Liver Oxylipin Concentrations. Prostaglandins Leukot. Essent. Fatty Acids 2021, 172, 102316. 10.1016/j.plefa.2021.102316. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref118] Schuster S.; Johnson C. D.; Hennebelle M.; Holtmann T.; Taha A. Y.; Kirpich I. A.; Eguchi A.; Ramsden C. E.; Papouchado B. G.; McClain C. J.; Feldstein A. E. Oxidized Linoleic Acid Metabolites Induce Liver Mitochondrial Dysfunction, Apoptosis, and NLRP3 Activation in Mice. J. Lipid Res. 2018, 59 (9), 1597–1609. 10.1194/jlr.M083741. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref119] Caligiuri S. P. B.; Parikh M.; Stamenkovic A.; Pierce G. N.; Aukema H. M. Dietary Modulation of Oxylipins in Cardiovascular Disease and Aging. Am. J. Physiol. Heart Circ. Physiol. 2017, 313 (5), H903–H918. 10.1152/ajpheart.00201.2017. [DOI] [PubMed] [Google Scholar]

[ref120] Ostermann A. I.; Schebb N. H. Effects of Omega-3 Fatty Acid Supplementation on the Pattern of Oxylipins: A Short Review about the Modulation of Hydroxy-, Dihydroxy-, and Epoxy-Fatty Acids. Food Funct. 2017, 8 (7), 2355–2367. 10.1039/C7FO00403F. [DOI] [PubMed] [Google Scholar]

[ref121] Innes J. K.; Calder P. C. Omega-6 Fatty Acids and Inflammation. Prostaglandins Leukot. Essent. Fatty Acids 2018, 132, 41–48. 10.1016/j.plefa.2018.03.004. [DOI] [PubMed] [Google Scholar]

[ref122] Hamilton J. S.; Klett E. L. Linoleic Acid and the Regulation of Glucose Homeostasis: A Review of the Evidence. Prostaglandins Leukot. Essent. Fatty Acids 2021, 175, 102366. 10.1016/j.plefa.2021.102366. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref123] Belury M. A.; Cole R. M.; Bailey B. E.; Ke J.-Y.; Andridge R. R.; Kiecolt-Glaser J. K. Erythrocyte Linoleic Acid, but Not Oleic Acid, Is Associated with Improvements in Body Composition in Men and Women. Mol. Nutr. Food Res. 2016, 60 (5), 1206–1212. 10.1002/mnfr.201500744. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref124] Shahidi F.; Ambigaipalan P. Omega-3 Polyunsaturated Fatty Acids and Their Health Benefits. Annu. Rev. Food Sci. Technol. 2018, 9 (1), 345–381. 10.1146/annurev-food-111317-095850. [DOI] [PubMed] [Google Scholar]

[ref125] Spector A. A.; Kim H.-Y. Emergence of Omega-3 Fatty Acids in Biomedical Research. Prostaglandins Leukot. Essent. Fatty Acids 2019, 140, 47–50. 10.1016/j.plefa.2018.11.017. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref126] Galli C.; Marangoni F. N-3 Fatty Acids in the Mediterranean Diet. Prostaglandins Leukot. Essent. Fatty Acids 2006, 75 (3), 129–133. 10.1016/j.plefa.2006.05.007. [DOI] [PubMed] [Google Scholar]

[ref127] Naghshi S.; Aune D.; Beyene J.; Mobarak S.; Asadi M.; Sadeghi O. Dietary Intake and Biomarkers of Alpha Linolenic Acid and Risk of All Cause, Cardiovascular, and Cancer Mortality: Systematic Review and Dose-Response Meta-Analysis of Cohort Studies. BMJ. 2021, 375, n2213. 10.1136/bmj.n2213. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref128] Seidel U.; Eberhardt K.; Wiebel M.; Luersen K.; Ipharraguerre I. R.; Haegele F. A.; Winterhalter P.; Bosy-Westphal A.; Schebb N. H.; Rimbach G. Stearidonic Acid Improves Eicosapentaenoic Acid Status: Studies in Humans and Cultured Hepatocytes. Front. Nutr. 2024, 11, 1359958. 10.3389/fnut.2024.1359958. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref129] Benjamin S.; Spener F. Conjugated Linoleic Acids as Functional Food: An Insight into Their Health Benefits. Nutr. Metab. 2009, 6 (1), 36. 10.1186/1743-7075-6-36. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref130] Yuan G.-F.; Chen X.-E.; Li D. Conjugated Linolenic Acids and Their Bioactivities: A Review. Food Funct. 2014, 5 (7), 1360–1368. 10.1039/c4fo00037d. [DOI] [PubMed] [Google Scholar]

[ref131] Paul D.; Dey T. K.; Chakraborty A.; Dhar P.. Chapter 13 - Promising Functional Lipids for Therapeutic Applications. In Role of Materials Science in Food Bioengineering; Handbook of Food Bioengineering; Grumezescu A. M., Holban A. M., Eds.; Academic Press, 2018; pp 413–449. 10.1016/B978-0-12-811448-3.00013-9. [DOI] [Google Scholar]

[ref132] Yuan G.; Sinclair A. J.; Xu C.; Li D. Incorporation and Metabolism of Punicic Acid in Healthy Young Humans. Mol. Nutr. Food Res. 2009, 53 (10), 1336–1342. 10.1002/mnfr.200800520. [DOI] [PubMed] [Google Scholar]

[ref133] Shabbir M. A.; Khan M. R.; Saeed M.; Pasha I.; Khalil A. A.; Siraj N. Punicic Acid: A Striking Health Substance to Combat Metabolic Syndromes in Humans. Lipids Health Dis. 2017, 16 (1), 99. 10.1186/s12944-017-0489-3. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref134] De Caterina R. N-3 Fatty Acids in Cardiovascular Disease. N. Engl. J. Med. 2011, 364 (25), 2439–2450. 10.1056/NEJMra1008153. [DOI] [PubMed] [Google Scholar]

[ref135] Cao M.; Yang F.; McClements D. J.; Guo Y.; Liu R.; Chang M.; Wei W.; Jin J.; Wang X. Impact of Dietary N-6/n-3 Fatty Acid Ratio of Atherosclerosis Risk: A Review. Prog. Lipid Res. 2024, 95, 101289. 10.1016/j.plipres.2024.101289. [DOI] [PubMed] [Google Scholar]

[ref136] Risérus U.; Willett W. C.; Hu F. B. Dietary Fats and Prevention of Type 2 Diabetes. Prog. Lipid Res. 2009, 48 (1), 44–51. 10.1016/j.plipres.2008.10.002. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref137] Erkkilä A.; de Mello V. D. F.; Risérus U.; Laaksonen D. E. Dietary Fatty Acids and Cardiovascular Disease: An Epidemiological Approach. Prog. Lipid Res. 2008, 47 (3), 172–187. 10.1016/j.plipres.2008.01.004. [DOI] [PubMed] [Google Scholar]

[ref138] Wendell S. G.; Baffi C.; Holguin F. Fatty Acids, Inflammation, and Asthma. J. Allergy Clin. Immunol. 2014, 133 (5), 1255–1264. 10.1016/j.jaci.2013.12.1087. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref139] Brühl L.; Weisshaar R.; Matthäus B. Epoxy Fatty Acids in Used Frying Fats and Oils, Edible Oils and Chocolate and Their Formation in Oils during Heating. Eur. J. Lipid Sci. Technol. 2016, 118 (3), 425–434. 10.1002/ejlt.201500235. [DOI] [Google Scholar]

[ref140] Emami S.; Zhang Z.; Taha A. Y. Quantitation of Oxylipins in Fish and Algae Oil Supplements Using Optimized Hydrolysis Procedures and Ultra-High Performance Liquid Chromatography Coupled to Tandem Mass-Spectrometry. J. Agric. Food Chem. 2020, 68 (35), 9329–9344. 10.1021/acs.jafc.0c02461. [DOI] [PubMed] [Google Scholar]

[ref141] Koch E.; Wiebel M.; Löwen A.; Willenberg I.; Schebb N. H. Characterization of the Oxylipin Pattern and Other Fatty Acid Oxidation Products in Freshly Pressed and Stored Plant Oils. J. Agric. Food Chem. 2022, 70 (40), 12935–12945. 10.1021/acs.jafc.2c04987. [DOI] [PubMed] [Google Scholar]

[ref142] Richardson C. E.; Hennebelle M.; Otoki Y.; Zamora D.; Yang J.; Hammock B. D.; Taha A. Y. Lipidomic Analysis of Oxidized Fatty Acids in Plant and Algae Oils. J. Agric. Food Chem. 2017, 65 (9), 1941–1951. 10.1021/acs.jafc.6b05559. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref143] Devassy J. G.; Yamaguchi T.; Monirujjaman M.; Gabbs M.; Ravandi A.; Zhou J.; Aukema H. M. Distinct Effects of Dietary Flax Compared to Fish Oil, Soy Protein Compared to Casein, and Sex on the Renal Oxylipin Profile in Models of Polycystic Kidney Disease. Prostaglandins Leukot. Essent. Fatty Acids 2017, 123, 1–13. 10.1016/j.plefa.2017.07.002. [DOI] [PubMed] [Google Scholar]

[ref144] Domínguez-Perles R.; Abellán Á.; León D.; Ferreres F.; Guy A.; Oger C.; Galano J. M.; Durand T.; Gil-Izquierdo Á. Sorting out the Phytoprostane and Phytofuran Profile in Vegetable Oils. Food Res. Int. 2018, 107, 619–628. 10.1016/j.foodres.2018.03.013. [DOI] [PubMed] [Google Scholar]

[ref145] Mubiru E.; Jacxsens L.; Papastergiadis A.; Lachat C.; Shrestha K.; Mozumder N. H. M. R.; De Meulenaer B. Exposure Assessment of Epoxy Fatty Acids through Consumption of Specific Foods Available in Belgium. Food Addit. Contam. Part A 2017, 34 (6), 1000–1011. 10.1080/19440049.2017.1310399. [DOI] [PubMed] [Google Scholar]

[ref146] Koch E.; Löwen A.; Schebb N. H. Do Meals Contain a Relevant Amount of Oxylipins? LC-MS-Based Analysis of Oxidized Fatty Acids in Food. Food Chem. 2024, 438, 137941. 10.1016/j.foodchem.2023.137941. [DOI] [PubMed] [Google Scholar]

[ref147] Abbattista R.; Feinberg N. G.; Snodgrass I. F.; Newman J. W.; Dandekar A. M. Unveiling the “Hidden Quality” of the Walnut Pellicle: A Precious Source of Bioactive Lipids. Front. Plant Sci. 2024, 15, 1395543. 10.3389/fpls.2024.1395543. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref148] Nourooz-Zadeh J.; Tajaddini-Sarmadi J.; Wolff S. P. Measurement of Hydroperoxides in Edible Oils Using the Ferrous Oxidation in Xylenol Orange Assay. J. Agric. Food Chem. 1995, 43 (1), 17–21. 10.1021/jf00049a005. [DOI] [Google Scholar]

[ref149] Choe E.; Min D. B. Mechanisms and Factors for Edible Oil Oxidation. Compr. Rev. Food Sci. Food Saf. 2006, 5 (4), 169–186. 10.1111/j.1541-4337.2006.00009.x. [DOI] [Google Scholar]

[ref150] Domínguez R.; Pateiro M.; Gagaoua M.; Barba F. J.; Zhang W.; Lorenzo J. M. A Comprehensive Review on Lipid Oxidation in Meat and Meat Products. Antioxidants 2019, 8 (10), 429. 10.3390/antiox8100429. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref151] Kanner J.; Lapidot T. The Stomach as a Bioreactor: Dietary Lipid Peroxidation in the Gastric Fluid and the Effects of Plant-Derived Antioxidants. Free Radic. Biol. Med. 2001, 31 (11), 1388–1395. 10.1016/S0891-5849(01)00718-3. [DOI] [PubMed] [Google Scholar]

[ref152] Márquez-Ruiz G.; Holgado F.; Ruiz-Méndez M. V.; Velasco J. Chemical Changes of Hydroperoxy-, Epoxy-, Keto- and Hydroxy-Model Lipids under Simulated Gastric Conditions. Foods 2021, 10 (9), 2035. 10.3390/foods10092035. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref153] Zhang Z.; Emami S.; Hennebelle M.; Morgan R. K.; Lerno L. A.; Slupsky C. M.; Lein P. J.; Taha A. Y. Linoleic Acid-Derived 13-Hydroxyoctadecadienoic Acid Is Absorbed and Incorporated into Rat Tissues. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2021, 1866 (3), 158870. 10.1016/j.bbalip.2020.158870. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref154] Kanazawa K.; Ashida H. Catabolic Fate of Dietary Trilinoleoylglycerol Hydroperoxides in Rat Gastrointestines. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1998, 1393 (2–3), 336–348. 10.1016/S0005-2760(98)00088-5. [DOI] [PubMed] [Google Scholar]

[ref155] Ringseis R.; Eder K. Regulation of Genes Involved in Lipid Metabolism by Dietary Oxidized Fat. Mol. Nutr. Food Res. 2011, 55 (1), 109–121. 10.1002/mnfr.201000424. [DOI] [PubMed] [Google Scholar]

[ref156] Zaunschirm M.; Pignitter M.; Kopic A.; Keßler C.; Hochkogler C.; Kretschy N.; Somoza M. M.; Somoza V. Exposure of Human Gastric Cells to Oxidized Lipids Stimulates Pathways of Amino Acid Biosynthesis on a Genomic and Metabolomic Level. Molecules 2019, 24 (22), 4111. 10.3390/molecules24224111. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref157] Fazzari M.; Trostchansky A.; Schopfer F. J.; Salvatore S. R.; Sánchez-Calvo B.; Vitturi D.; Valderrama R.; Barroso J. B.; Radi R.; Freeman B. A.; Rubbo H. Olives and Olive Oil Are Sources of Electrophilic Fatty Acid Nitroalkenes. PLoS One 2014, 9 (1), e84884 10.1371/journal.pone.0084884. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref158] Khoo N. K.; Schopfer F. J. Nitrated Fatty Acids: From Diet to Disease. Curr. Opin. Physiol. 2019, 9, 67–72. 10.1016/j.cophys.2019.04.013. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref159] Karwowska M.; Kononiuk A. Nitrates/Nitrites in Food—Risk for Nitrosative Stress and Benefits. Antioxidants 2020, 9 (3), 241. 10.3390/antiox9030241. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref160] Bonacci G.; Baker P. R. S.; Salvatore S. R.; Shores D.; Khoo N. K. H.; Koenitzer J. R.; Vitturi D. A.; Woodcock S. R.; Golin-Bisello F.; Cole M. P.; Watkins S.; St Croix C.; Batthyany C. I.; Freeman B. A.; Schopfer F. J. Conjugated Linoleic Acid Is a Preferential Substrate for Fatty Acid Nitration. J. Biol. Chem. 2012, 287, 44071–44082. 10.1074/jbc.M112.401356. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref161] Charles R. L.; Rudyk O.; Prysyazhna O.; Kamynina A.; Yang J.; Morisseau C.; Hammock B. D.; Freeman B. A.; Eaton P. Protection from Hypertension in Mice by the Mediterranean Diet Is Mediated by Nitro Fatty Acid Inhibition of Soluble Epoxide Hydrolase. Proc. Natl. Acad. Sci. U. S. A. 2014, 111 (22), 8167–8172. 10.1073/pnas.1402965111. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref162] Duarte S.; Melo T.; Domingues R.; de Dios Alché J.; Pérez-Sala D. Insight into the Cellular Effects of Nitrated Phospholipids: Evidence for Pleiotropic Mechanisms of Action. Free Radic. Biol. Med. 2019, 144, 192–202. 10.1016/j.freeradbiomed.2019.06.003. [DOI] [PubMed] [Google Scholar]

[ref163] O’Neill G. P.; Ford-Hutchinson A. W. Expression of MRNA for Cyclooxygenase-1 and Cyclooxygenase-2 in Human Tissues. FEBS Lett. 1993, 330 (2), 157–160. 10.1016/0014-5793(93)80263-T. [DOI] [PubMed] [Google Scholar]

[ref164] Zidar N.; Odar K.; Glavac D.; Jerse M.; Zupanc T.; Stajer D. Cyclooxygenase in Normal Human Tissues - Is COX-1 Really a Constitutive Isoform, and COX-2 an Inducible Isoform?. J. Cell. Mol. Med. 2009, 13 (9b), 3753–3763. 10.1111/j.1582-4934.2008.00430.x. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref165] Kirkby N. S.; Chan M. V.; Zaiss A. K.; Garcia-Vaz E.; Jiao J.; Berglund L. M.; Verdu E. F.; Ahmetaj-Shala B.; Wallace J. L.; Herschman H. R.; Gomez M. F.; Mitchell J. A. Systematic Study of Constitutive Cyclooxygenase-2 Expression: Role of NF-ΚB and NFAT Transcriptional Pathways. Proc. Natl. Acad. Sci. U. S. A. 2016, 113 (2), 434–439. 10.1073/pnas.1517642113. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref166] Ringbom T.; Huss U.; Stenholm Å.; Flock S.; Skattebøl L.; Perera P.; Bohlin L. COX-2 Inhibitory Effects of Naturally Occurring and Modified Fatty Acids. J. Nat. Prod. 2001, 64 (6), 745–749. 10.1021/np000620d. [DOI] [PubMed] [Google Scholar]

[ref167] Laneuville O.; Breuer D. K.; Xu N.; Huang Z. H.; Gage D. A.; Watson J. T.; Lagarde M.; DeWitt D. L.; Smith W. L. Fatty Acid Substrate Specificities of Human Prostaglandin-Endoperoxide H Synthase-1 and – 2. J. Biol. Chem. 1995, 270 (33), 19330–19336. 10.1074/jbc.270.33.19330. [DOI] [PubMed] [Google Scholar]

[ref168] Marnett L. J. Cyclooxygenase Mechanisms. Curr. Opin. Chem. Biol. 2000, 4 (5), 545–552. 10.1016/S1367-5931(00)00130-7. [DOI] [PubMed] [Google Scholar]

[ref169] Hamberg M. Stereochemistry of Oxygenation of Linoleic Acid Catalyzed by Prostaglandin-Endoperoxide H Synthase-2. Arch. Biochem. Biophys. 1998, 349 (2), 376–380. 10.1006/abbi.1997.0443. [DOI] [PubMed] [Google Scholar]

[ref170] Altmann R.; Hausmann M.; Spöttl T.; Gruber M.; Bull A. W.; Menzel K.; Vogl D.; Herfarth H.; Schölmerich J.; Falk W.; Rogler G. 13-Oxo-ODE Is an Endogenous Ligand for PPARγ in Human Colonic Epithelial Cells. Biochem. Pharmacol. 2007, 74 (4), 612–622. 10.1016/j.bcp.2007.05.027. [DOI] [PubMed] [Google Scholar]

[ref171] Funk C. D.; Powell W. S. Metabolism of Linoleic Acid by Prostaglandin Endoperoxide Synthase from Adult and Fetal Blood Vessels. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1983, 754 (1), 57–71. 10.1016/0005-2760(83)90082-6. [DOI] [PubMed] [Google Scholar]

[ref172] Fuchs D.; Tang X.; Johnsson A.-K.; Dahlén S.-E.; Hamberg M.; Wheelock C. E. Eosinophils Synthesize Trihydroxyoctadecenoic Acids (TriHOMEs) via a 15-Lipoxygenase Dependent Process. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2020, 1865 (4), 158611. 10.1016/j.bbalip.2020.158611. [DOI] [PubMed] [Google Scholar]

[ref173] Kuhn H.; Banthiya S.; van Leyen K. Mammalian Lipoxygenases and Their Biological Relevance. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2015, 1851 (4), 308–330. 10.1016/j.bbalip.2014.10.002. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref174] Wuest S. J. A.; Crucet M.; Gemperle C.; Loretz C.; Hersberger M. Expression and Regulation of 12/15-Lipoxygenases in Human Primary Macrophages. Atherosclerosis 2012, 225 (1), 121–127. 10.1016/j.atherosclerosis.2012.07.022. [DOI] [PubMed] [Google Scholar]

[ref175] Jisaka M.; Kim R. B.; Boeglin W. E.; Nanney L. B.; Brash A. R. Molecular Cloning and Functional Expression of a Phorbol Ester-Inducible 8S-Lipoxygenase from Mouse Skin. J. Biol. Chem. 1997, 272 (39), 24410–24416. 10.1074/jbc.272.39.24410. [DOI] [PubMed] [Google Scholar]

[ref176] Lehnert N.; Solomon E. I. Density-Functional Investigation on the Mechanism of H-Atom Abstraction by Lipoxygenase. J. Biol. Inorg. Chem. 2003, 8 (3), 294–305. 10.1007/s00775-002-0415-6. [DOI] [PubMed] [Google Scholar]

[ref177] Gardner H. W.; Kleiman R.; Weisleder D. Homolytic Decomposition of Linoleic Acid Hydroperoxide: Identification of Fatty Acid Products. Lipids 1974, 9 (9), 696–706. 10.1007/BF02532178. [DOI] [Google Scholar]

[ref178] Soberman R. J.; Harper T. W.; Betteridge D.; Lewis R. A.; Austen K. F. Characterization and Separation of the Arachidonic Acid 5-Lipoxygenase and Linoleic Acid Omega-6 Lipoxygenase (Arachidonic Acid 15-Lipoxygenase) of Human Polymorphonuclear Leukocytes. J. Biol. Chem. 1985, 260 (7), 4508–4515. 10.1016/S0021-9258(18)89293-6. [DOI] [PubMed] [Google Scholar]

[ref179] Baer A. N.; Costello P. B.; Green F. A. Stereospecificity of the Products of the Fatty Acid Oxygenases Derived from Psoriatic Scales. J. Lipid Res. 1991, 32 (2), 341–347. 10.1016/S0022-2275(20)42094-2. [DOI] [PubMed] [Google Scholar]

[ref180] Zheng Y.; Yin H.; Boeglin W. E.; Elias P. M.; Crumrine D.; Beier D. R.; Brash A. R. Lipoxygenases Mediate the Effect of Essential Fatty Acid in Skin Barrier Formation: A Proposed Role In Releasing Omega-Hydroxyceramide For Construction Of The Corneocyte Lipid Envelope. J. Biol. Chem. 2011, 286 (27), 24046–24056. 10.1074/jbc.M111.251496. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref181] Haeggström J. Z.; Funk C. D. Lipoxygenase and Leukotriene Pathways: Biochemistry, Biology, and Roles in Disease. Chem. Rev. 2011, 111 (10), 5866–5898. 10.1021/cr200246d. [DOI] [PubMed] [Google Scholar]

[ref182] Reinaud O.; Delaforge M.; Boucher J. L.; Rocchiccioli F.; Mansuy D. Oxidative Metabolism of Linoleic Acid by Human Leukocytes. Biochem. Biophys. Res. Commun. 1989, 161 (2), 883–891. 10.1016/0006-291X(89)92682-X. [DOI] [PubMed] [Google Scholar]

[ref183] Kühn H.; Belkner J.; Wiesner R.; Alder L. Occurrence of 9- and 13-Keto-Octadecadienoic Acid in Biological Membranes Oxygenated by the Reticulocyte Lipoxygenase. Arch. Biochem. Biophys. 1990, 279 (2), 218–224. 10.1016/0003-9861(90)90484-G. [DOI] [PubMed] [Google Scholar]

[ref184] Bull A. W.; Earles S. M.; Bronstein J. C. Metabolism of Oxidized Linoleic Acid: Distribution of Activity for the Enzymatic Oxidation of 13-Hydroxyoctadecadienoic Acid to 13-Oxooctadecadienoic Acid in Rat Tissues. Prostaglandins 1991, 41 (1), 43–50. 10.1016/0090-6980(91)90103-M. [DOI] [PubMed] [Google Scholar]

[ref185] Goodfriend T. L.; Ball D. L.; Egan B. M.; Campbell W. B.; Nithipatikom K. Epoxy-Keto Derivative of Linoleic Acid Stimulates Aldosterone Secretion. Hypertension 2004, 43 (2), 358–363. 10.1161/01.HYP.0000113294.06704.64. [DOI] [PubMed] [Google Scholar]

[ref186] Brash A. R.; Noguchi S.; Boeglin W. E.; Calcutt M. W.; Stec D. F.; Schneider C.; Meyer J. M. Two C18 Hydroxy-Cyclohexenone Fatty Acids from Mammalian Epidermis: Potential Relation to 12R-Lipoxygenase and Covalent Binding of Ceramides. J. Biol. Chem. 2023, 299 (6), 104739. 10.1016/j.jbc.2023.104739. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref187] Edin M. L.; Yamanashi H.; Boeglin W. E.; Graves J. P.; DeGraff L. M.; Lih F. B.; Zeldin D. C.; Brash A. R. Epoxide Hydrolase 3 (Ephx3) Gene Disruption Reduces Ceramide Linoleate Epoxide Hydrolysis and Impairs Skin Barrier Function. J. Biol. Chem. 2021, 296, 100198. 10.1074/jbc.RA120.016570. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref188] Hamberg M. An Epoxy Alcohol Synthase Pathway in Higher Plants: Biosynthesis of Antifungal Trihydroxy Oxylipins in Leaves of Potato. Lipids 1999, 34 (11), 1131–1142. 10.1007/s11745-999-0464-7. [DOI] [PubMed] [Google Scholar]

[ref189] Collins J. F.; Hu Z.; Ranganathan P. N.; Feng D.; Garrick L. M.; Garrick M. D.; Browne R. W. Induction of Arachidonate 12-Lipoxygenase (Alox15) in Intestine of Iron-Deficient Rats Correlates with the Production of Biologically Active Lipid Mediators. Am. J. Physiol.-Gastrointest. Liver Physiol. 2008, 294 (4), G948–G962. 10.1152/ajpgi.00274.2007. [DOI] [PubMed] [Google Scholar]

[ref190] Caligiuri S. P. B.; Love K.; Winter T.; Gauthier J.; Taylor C. G.; Blydt-Hansen T.; Zahradka P.; Aukema H. M. Dietary Linoleic Acid and α-Linolenic Acid Differentially Affect Renal Oxylipins and Phospholipid Fatty Acids in Diet-Induced Obese Rats. J. Nutr. 2013, 143 (9), 1421–1431. 10.3945/jn.113.177360. [DOI] [PubMed] [Google Scholar]

[ref191] Cossette C.; Patel P.; Anumolu J. R.; Sivendran S.; Lee G. J.; Gravel S.; Graham F. D.; Lesimple A.; Mamer O. A.; Rokach J.; Powell W. S. Human Neutrophils Convert the Sebum-Derived Polyunsaturated Fatty Acid Sebaleic Acid to a Potent Granulocyte Chemoattractant. J. Biol. Chem. 2008, 283 (17), 11234–11243. 10.1074/jbc.M709531200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref192] Arterburn L. M.; Barclay W.; Dangi B.; Flatt J.; Lee J.; Vinjamoori D.. Oxylipins From Stearidonic Acid and Gamma-Linolenic Acid and Methods Of Making and Using The Same. US Patent US 2007/0248586 A1, 2007.

[ref193] Danielson P. B. The Cytochrome P450 Superfamily: Biochemistry, Evolution and Drug Metabolism in Humans. Curr. Drug Metab. 2002, 3, 561–597. 10.2174/1389200023337054. [DOI] [PubMed] [Google Scholar]

[ref194] Nebert D. W.; Wikvall K.; Miller W. L. Human Cytochromes P450 in Health and Disease. Philos. Trans. R. Soc. B Biol. Sci. 2013, 368 (1612), 20120431. 10.1098/rstb.2012.0431. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref195] Kells P. M.; Ouellet H.; Santos-Aberturas J.; Aparicio J. F.; Podust L. M. Structure of Cytochrome P450 PimD Suggests Epoxidation of the Polyene Macrolide Pimaricin Occurs via a Hydroperoxoferric Intermediate. Chem. Biol. 2010, 17 (8), 841–851. 10.1016/j.chembiol.2010.05.026. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref196] Meunier B.; de Visser S. P.; Shaik S. Mechanism of Oxidation Reactions Catalyzed by Cytochrome P450 Enzymes. Chem. Rev. 2004, 104 (9), 3947–3980. 10.1021/cr020443g. [DOI] [PubMed] [Google Scholar]

[ref197] Morisseau C.; Kodani S. D.; Kamita S. G.; Yang J.; Lee K. S. S.; Hammock B. D. Relative Importance of Soluble and Microsomal Epoxide Hydrolases for the Hydrolysis of Epoxy-Fatty Acids in Human Tissues. Int. J. Mol. Sci. 2021, 22 (9), 4993. 10.3390/ijms22094993. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref198] Moran J. H.; Mitchell L. A.; Bradbury J. A.; Qu W.; Zeldin D. C.; Schnellmann R. G.; Grant D. F. Analysis of the Cytotoxic Properties of Linoleic Acid Metabolites Produced by Renal and Hepatic P450s. Toxicol. Appl. Pharmacol. 2000, 168 (3), 268–279. 10.1006/taap.2000.9053. [DOI] [PubMed] [Google Scholar]

[ref199] Draper A. J.; Hammock B. D. Identification of CYP2C9 as a Human Liver Microsomal Linoleic Acid Epoxygenase. Arch. Biochem. Biophys. 2000, 376 (1), 199–205. 10.1006/abbi.2000.1705. [DOI] [PubMed] [Google Scholar]

[ref200] Bylund J.; Kunz T.; Valmsen K.; Oliw E. H. Cytochromes P450 with Bisallylic Hydroxylation Activity on Arachidonic and Linoleic Acids Studied with Human Recombinant Enzymes and with Human and Rat Liver Microsomes. J. Pharmacol. Exp. Ther. 1998, 284, 51–60. [PubMed] [Google Scholar]

[ref201] Laethem R. M.; Balazy M.; Koop D. R. Epoxidation of C18 Unsaturated Fatty Acids by Cytochromes P4502C2 and P4502CAA. Drug Metab. Dispos. 1996, 24, 664–668. [PubMed] [Google Scholar]

[ref202] Williamson K. C.; Morisseau C.; Maxwell J. E.; Hammock B. D. Regio- and Enantioselective Hydrolysis of Phenyloxiranes Catalyzed by Soluble Epoxide Hydrolase. Tetrahedron Asymmetry 2000, 11 (22), 4451–4462. 10.1016/S0957-4166(00)00437-7. [DOI] [Google Scholar]

[ref203] Teder T.; Boeglin W. E.; Brash A. R. Lipoxygenase-Catalyzed Transformation of Epoxy Fatty Acids to Hydroxy-Endoperoxides: A Potential P450 and Lipoxygenase Interaction. J. Lipid Res. 2014, 55 (12), 2587–2596. 10.1194/jlr.M054072. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref204] Tsikas D.; Sawa M.; Brunner G.; Gutzki F.-M.; Meyer H. H.; Frölich J. C. Gas Chromatography-Mass Spectrometry of cis-9,10-Epoxyoctadecanoic Acid (cis-EODA): I. Direct Evidence for cis-EODA Formation from Oleic Acid Oxidation by Liver Microsomes and Isolated Hepatocytes. J. Chromatogr. B 2003, 784 (2), 351–365. 10.1016/S1570-0232(02)00821-8. [DOI] [PubMed] [Google Scholar]

[ref205] Thum T.; Batkai S.; Malinski P. G.; Becker T.; Mevius I.; Klempnauer J.; Meyer H. H.; Frölich J. C.; Borlak J.; Tsikas D. Measurement and Diagnostic Use of Hepatic Cytochrome P450 Metabolism of Oleic Acid in Liver Disease. Liver Int. 2010, 30 (8), 1181–1188. 10.1111/j.1478-3231.2010.02310.x. [DOI] [PubMed] [Google Scholar]

[ref206] Oliw E. H. Bis-Allylic Hydroxylation of Linoleic Acid and Arachidonic Acid by Human Hepatic Monooxygenases. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1993, 1166 (2), 258–263. 10.1016/0005-2760(93)90106-J. [DOI] [PubMed] [Google Scholar]

[ref207] Adas F.; Salaün J. P.; Berthou F.; Picart D.; Simon B.; Amet Y. Requirement for ω and (ω-1)-Hydroxylations of Fatty Acids by Human Cytochromes P450 2E1 and 4A11. J. Lipid Res. 1999, 40 (11), 1990–1997. 10.1016/S0022-2275(20)32422-6. [DOI] [PubMed] [Google Scholar]

[ref208] Nguyen X.; Wang M.-H.; Reddy K. M.; Falck J. R.; Schwartzman M. L. Kinetic Profile of the Rat CYP4A Isoforms: Arachidonic Acid Metabolism and Isoform-Specific Inhibitors. Am. J. Physiol.-Regul. Integr. Comp. Physiol. 1999, 276 (6), R1691–R1700. 10.1152/ajpregu.1999.276.6.R1691. [DOI] [PubMed] [Google Scholar]

[ref209] Chuang S. S.; Helvig C.; Taimi M.; Ramshaw H. A.; Collop A. H.; Amad M.; White J. A.; Petkovich M.; Jones G.; Korczak B. CYP2U1, a Novel Human Thymus- and Brain-Specific Cytochrome P450, Catalyzes ω- and (ω-1)-Hydroxylation of Fatty Acids. J. Biol. Chem. 2004, 279 (8), 6305–6314. 10.1074/jbc.M311830200. [DOI] [PubMed] [Google Scholar]

[ref210] Bjorkhem I.; Danielsson H. Omega- and (Omega-1)-Oxidation of Fatty Acids by Rat Liver Microsomes. Eur. J. Biochem. 1970, 17 (3), 450–459. 10.1111/j.1432-1033.1970.tb01186.x. [DOI] [PubMed] [Google Scholar]

[ref211] Le Quéré V.; Plée-Gautier E.; Potin P.; Madec S.; Salaün J.-P. Human CYP4F3s Are the Main Catalysts in the Oxidation of Fatty Acid Epoxides. J. Lipid Res. 2004, 45 (8), 1446–1458. 10.1194/jlr.M300463-JLR200. [DOI] [PubMed] [Google Scholar]

[ref212] Dhar M.; Sepkovic D. W.; Hirani V.; Magnusson R. P.; Lasker J. M. Omega Oxidation of 3-Hydroxy Fatty Acids by the Human CYP4F Gene Subfamily Enzyme CYP4F11. J. Lipid Res. 2008, 49 (3), 612–624. 10.1194/jlr.M700450-JLR200. [DOI] [PubMed] [Google Scholar]

[ref213] Aggarwal N.; Kitano S.; Puah G. R. Y.; Kittelmann S.; Hwang I. Y.; Chang M. W. Microbiome and Human Health: Current Understanding, Engineering, and Enabling Technologies. Chem. Rev. 2023, 123 (1), 31–72. 10.1021/acs.chemrev.2c00431. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref214] Schoeler M.; Caesar R. Dietary Lipids, Gut Microbiota and Lipid Metabolism. Rev. Endocr. Metab. Disord. 2019, 20 (4), 461–472. 10.1007/s11154-019-09512-0. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref215] Schoeler M.; Ellero-Simatos S.; Birkner T.; Mayneris-Perxachs J.; Olsson L.; Brolin H.; Loeber U.; Kraft J. D.; Polizzi A.; Martí-Navas M.; Puig J.; Moschetta A.; Montagner A.; Gourdy P.; Heymes C.; Guillou H.; Tremaroli V.; Fernández-Real J. M.; Forslund S. K.; Burcelin R.; Caesar R. The Interplay between Dietary Fatty Acids and Gut Microbiota Influences Host Metabolism and Hepatic Steatosis. Nat. Commun. 2023, 14 (1), 5329. 10.1038/s41467-023-41074-3. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref216] Zhang Q.; Zhang L.; Chen C.; Li P.; Lu B. The Gut Microbiota-Artery Axis: A Bridge between Dietary Lipids and Atherosclerosis?. Prog. Lipid Res. 2023, 89, 101209. 10.1016/j.plipres.2022.101209. [DOI] [PubMed] [Google Scholar]

[ref217] Jonsson A. L.; Bäckhed F. Role of Gut Microbiota in Atherosclerosis. Nat. Rev. Cardiol. 2017, 14 (2), 79–87. 10.1038/nrcardio.2016.183. [DOI] [PubMed] [Google Scholar]

[ref218] Fan C.; Xu J.; Tong H.; Fang Y.; Chen Y.; Lin Y.; Chen R.; Chen F.; Wu G. Gut-Brain Communication Mediates the Impact of Dietary Lipids on Cognitive Capacity. Food Funct. 2024, 15 (4), 1803–1824. 10.1039/D3FO05288E. [DOI] [PubMed] [Google Scholar]

[ref219] Özçam M.; Lynch S. V. The Gut-Airway Microbiome Axis in Health and Respiratory Diseases. Nat. Rev. Microbiol. 2024, 22, 492. 10.1038/s41579-024-01048-8. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref220] Budden K. F.; Gellatly S. L.; Wood D. L. A.; Cooper M. A.; Morrison M.; Hugenholtz P.; Hansbro P. M. Emerging Pathogenic Links between Microbiota and the Gut-Lung Axis. Nat. Rev. Microbiol. 2017, 15 (1), 55–63. 10.1038/nrmicro.2016.142. [DOI] [PubMed] [Google Scholar]

[ref221] Pinckard K. M.; Shettigar V. K.; Wright K. R.; Abay E.; Baer L. A.; Vidal P.; Dewal R. S.; Das D.; Duarte-Sanmiguel S.; Hernández-Saavedra D.; Arts P. J.; Lehnig A. C.; Bussberg V.; Narain N. R.; Kiebish M. A.; Yi F.; Sparks L. M.; Goodpaster B. H.; Smith S. R.; Pratley R. E.; Lewandowski E. D.; Raman S. V.; Wold L. E.; Gallego-Perez D.; Coen P. M.; Ziolo M. T.; Stanford K. I. A Novel Endocrine Role for the BAT-Released Lipokine 12,13-DiHOME to Mediate Cardiac Function. Circulation 2021, 143 (2), 145–159. 10.1161/CIRCULATIONAHA.120.049813. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref222] Zimmer B.; Angioni C.; Osthues T.; Toewe A.; Thomas D.; Pierre S. C.; Geisslinger G.; Scholich K.; Sisignano M. The Oxidized Linoleic Acid Metabolite 12,13-DiHOME Mediates Thermal Hyperalgesia during Inflammatory Pain. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2018, 1863 (7), 669–678. 10.1016/j.bbalip.2018.03.012. [DOI] [PubMed] [Google Scholar]

[ref223] Kishino S.; Takeuchi M.; Park S.-B.; Hirata A.; Kitamura N.; Kunisawa J.; Kiyono H.; Iwamoto R.; Isobe Y.; Arita M.; Arai H.; Ueda K.; Shima J.; Takahashi S.; Yokozeki K.; Shimizu S.; Ogawa J. Polyunsaturated Fatty Acid Saturation by Gut Lactic Acid Bacteria Affecting Host Lipid Composition. Proc. Natl. Acad. Sci. U. S. A. 2013, 110 (44), 17808–17813. 10.1073/pnas.1312937110. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref224] Polan C. E.; McNeill J. J.; Tove S. B. Biohydrogenation of Unsaturated Fatty Acids by Rumen Bacteria. J. Bacteriol. 1964, 88 (4), 1056–1064. 10.1128/jb.88.4.1056-1064.1964. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref225] Mir P. S.; McAllister T. A.; Scott S.; Aalhus J.; Baron V.; McCartney D.; Charmley E.; Goonewardene L.; Basarab J.; Okine E.; Weselake R. J.; Mir Z. Conjugated Linoleic Acid-Enriched Beef Production. Am. J. Clin. Nutr. 2004, 79, 1207S–1211S. 10.1093/ajcn/79.6.1207S. [DOI] [PubMed] [Google Scholar]

[ref226] Ferlay A.; Bernard L.; Meynadier A.; Malpuech-Brugère C. Production of Trans and Conjugated Fatty Acids in Dairy Ruminants and Their Putative Effects on Human Health: A Review. Biochimie 2017, 141, 107–120. 10.1016/j.biochi.2017.08.006. [DOI] [PubMed] [Google Scholar]

[ref227] Lee K. W.; Lee H. J.; Cho H. Y.; Kim Y. J. Role of the Conjugated Linoleic Acid in the Prevention of Cancer. Crit. Rev. Food Sci. Nutr. 2005, 45 (2), 135–144. 10.1080/10408690490911800. [DOI] [PubMed] [Google Scholar]

[ref228] Wahle K. W. J.; Heys S. D.; Rotondo D. Conjugated Linoleic Acids: Are They Beneficial or Detrimental to Health?. Prog. Lipid Res. 2004, 43 (6), 553–587. 10.1016/j.plipres.2004.08.002. [DOI] [PubMed] [Google Scholar]

[ref229] Dhar Dubey K. K.; Sharma G.; Kumar A. Conjugated Linolenic Acids: Implication in Cancer. J. Agric. Food Chem. 2019, 67 (22), 6091–6101. 10.1021/acs.jafc.9b01379. [DOI] [PubMed] [Google Scholar]

[ref230] Tanaka T.; Hosokawa M.; Yasui Y.; Ishigamori R.; Miyashita K. Cancer Chemopreventive Ability of Conjugated Linolenic Acids. Int. J. Mol. Sci. 2011, 12 (11), 7495–7509. 10.3390/ijms12117495. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref231] Scollan N. D.; Price E. M.; Morgan S. A.; Huws S. A.; Shingfield K. J. Can We Improve the Nutritional Quality of Meat?. Proc. Nutr. Soc. 2017, 76 (4), 603–618. 10.1017/S0029665117001112. [DOI] [PubMed] [Google Scholar]

[ref232] Ponnampalam E.; Mann N.; Sinclair A. Effect of Feeding Systems on Omega-3 Fatty Acids, Conjugated Linoleic Acid and Trans Fatty Acids in Australian Beef Cuts: Potential Impact on Humnan Health. Asia Pac. J. Clin. Nutr. 2006, 15, 21–29. [PubMed] [Google Scholar]

[ref233] Vahmani P.; Ponnampalam E. N.; Kraft J.; Mapiye C.; Bermingham E. N.; Watkins P. J.; Proctor S. D.; Dugan M. E. R. Bioactivity and Health Effects of Ruminant Meat Lipids. Invited Review. Meat Sci. 2020, 165, 108114. 10.1016/j.meatsci.2020.108114. [DOI] [PubMed] [Google Scholar]

[ref234] Hira T.; Ogasawara S.; Yahagi A.; Kamachi M.; Li J.; Nishimura S.; Sakaino M.; Yamashita T.; Kishino S.; Ogawa J.; Hara H. Novel Mechanism of Fatty Acid Sensing in Enteroendocrine Cells: Specific Structures in Oxo-Fatty Acids Produced by Gut Bacteria Are Responsible for CCK Secretion in STC-1 Cells via GPR40. Mol. Nutr. Food Res. 2018, 62 (19), 1800146. 10.1002/mnfr.201800146. [DOI] [PubMed] [Google Scholar]

[ref235] Hirata A.; Kishino S.; Park S.-B.; Takeuchi M.; Kitamura N.; Ogawa J. A Novel Unsaturated Fatty Acid Hydratase toward C16 to C22 Fatty Acids from Lactobacillus acidophilus. J. Lipid Res. 2015, 56 (7), 1340–1350. 10.1194/jlr.M059444. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref236] Kishimoto N.; Yamamoto I.; Toraishi K.; Yoshioka S.; Saito K.; Masuda H.; Fujita T. Two Distinct Pathways for the Formation of Hydroxy FA from Linoleic Acid by Lactic Acid Bacteria. Lipids 2003, 38 (12), 1269–1274. 10.1007/s11745-003-1188-4. [DOI] [PubMed] [Google Scholar]

[ref237] Takeuchi M.; Kishino S.; Tanabe K.; Hirata A.; Park S.-B.; Shimizu S.; Ogawa J. Hydroxy Fatty Acid Production by Pediococcus sp. Eur. J. Lipid Sci. Technol. 2013, 115 (4), 386–393. 10.1002/ejlt.201200414. [DOI] [Google Scholar]

[ref238] Hudson J. A.; Morvan B.; Joblin K. N. Hydration of Linoleic Acid by Bacteria Isolated from Ruminants. FEMS Microbiol. Lett. 1998, 169 (2), 277–282. 10.1111/j.1574-6968.1998.tb13329.x. [DOI] [PubMed] [Google Scholar]

[ref239] Koritala S.; Hou C. T.; Hesseltine C. W.; Bagby M. O. Microbial Conversion of Oleic Acid to 10-Hydroxystearic Acid. Appl. Microbiol. Biotechnol. 1989, 32 (3), 299–304. 10.1007/BF00184978. [DOI] [Google Scholar]

[ref240] Hou C. T. Conversion of Linoleic Acid to 10-Hydroxy-12(Z)-Octadecenoic Acid by Flavobacterium sp. J. Am. Oil Chem. Soc. 1994, 71 (9), 975–978. 10.1007/BF02542264. [DOI] [Google Scholar]

[ref241] Hou C. T. New Bioactive Fatty Acids. Asia Pac. J. Clin. Nutr. 2008, 17, 192–195. [PubMed] [Google Scholar]

[ref242] Hou C. T. A Novel Compound, 12,13,17-Trihydroxy-9(Z)-Octadecenoic Acid, from Linoleic Acid by a New Microbial Isolate Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 1996, 73 (11), 1359–1362. 10.1007/BF02523497. [DOI] [Google Scholar]

[ref243] Iwasaki Y.; Brown W.; Hou C. T. Biosynthetic Pathway of Diepoxy Bicyclic FA from Linoleic Acid by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2002, 79 (4), 369–372. 10.1007/s11746-002-0490-x. [DOI] [Google Scholar]

[ref244] Gardner H. W.; Hou C. T.; Weisleder D.; Brown W. Biotransformation of Linoleic Acid by Clavibacter sp. ALA2: Heterocyclic and Heterobicyclic Fatty Acids. Lipids 2000, 35 (10), 1055–1060. 10.1007/s11745-000-0618-7. [DOI] [PubMed] [Google Scholar]

[ref245] Hosokawa M.; Hou C. T.; Weisleder D.; Brown W. Biosynthesis of Tetrahydrofuranyl Fatty Acids from Linoleic Acid by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2003, 80 (2), 145–149. 10.1007/s11746-003-0667-3. [DOI] [Google Scholar]

[ref246] Hou C. T.; Gardner H. W.; Brown W. 12,13,16-Trihydroxy-9(Z)-Octadecenoic Acid, a Possible Intermediate in the Bioconversion of Linoleic Acid to Tetrahydrofuranyl Fatty Acids by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2001, 78 (11), 1167–1169. 10.1007/s11746-001-0407-8. [DOI] [Google Scholar]

[ref247] Hosokawa M.; Hou C. T.; Weisleder D. Production of Novel Tetrahydroxyfuranyl Fatty Acids from α-Linolenic Acid by Clavibacter sp. Strain ALA2. Appl. Environ. Microbiol. 2003, 69 (7), 3868–3873. 10.1128/AEM.69.7.3868-3873.2003. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref248] Hosokawa M.; Hou C. T.; Weisleder D. Bioconversion of N–3 and N–6 PUFA by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 2003, 80 (11), 1085–1091. 10.1007/s11746-003-0824-8. [DOI] [Google Scholar]

[ref249] Andreou A.; Göbel C.; Hamberg M.; Feussner I. A Bisallylic Mini-Lipoxygenase from Cyanobacterium Cyanothece sp. That Has an Iron as Cofactor. J. Biol. Chem. 2010, 285 (19), 14178–14186. 10.1074/jbc.M109.094771. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref250] Gao B.; Boeglin W. E.; Brash A. R. Omega-3 Fatty Acids Are Oxygenated at the n-7 Carbon by the Lipoxygenase Domain of a Fusion Protein in the Cyanobacterium Acaryochloris marina. Biochim. Biophys. Acta 2010, 1801 (1), 58–63. 10.1016/j.bbalip.2009.09.004. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref251] el-Sharkawy S. H.; Yang W.; Dostal L.; Rosazza J. P. Microbial Oxidation of Oleic Acid. Appl. Environ. Microbiol. 1992, 58 (7), 2116–2122. 10.1128/aem.58.7.2116-2122.1992. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref252] Hou C. T. Production of Hydroxy Fatty Acids from Unsaturated Fatty Acids by Flavobacterium sp. DS5 Hydratase, a C-10 Positional- and Cis Unsaturation-Specific Enzyme. J. Am. Oil Chem. Soc. 1995, 72 (11), 1265–1270. 10.1007/BF02546197. [DOI] [Google Scholar]

[ref253] Hou C. T. Production of 10-Ketostearic Acid from Oleic Acid by Flavobacterium sp. Strain DS5 (NRRL B-14859). Appl. Environ. Microbiol. 1994, 60 (10), 3760–3763. 10.1128/aem.60.10.3760-3763.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref254] Hou C. T.; Bagby M. O.; Plattner R. D.; Koritala S. A Novel Compound, 7,10-Dihydroxy-8(E)-Octadecenoic Acid from Oleic Acid by Bioconversion. J. Am. Oil Chem. Soc. 1991, 68 (2), 99–101. 10.1007/BF02662326. [DOI] [Google Scholar]

[ref255] Hou C. T.; Bagby M. O. 10-Hydroxy-8(Z)-Octadecenoic Acid, an Intermediate in the Bioconversion of Oleic Acid to 7,10-Dihydroxy-8(E)-Octadecenoic Acid. J. Ind. Microbiol. 1992, 9 (2), 103–107. 10.1007/BF01569740. [DOI] [Google Scholar]

[ref256] Guerrero A.; Casals I.; Busquets M.; Leon Y.; Manresa A. Oxydation of Oleic Acid to (E)-10-Hydroperoxy-8-Octadecenoic and (E)-10-Hydroxy-8-Octadecenoic Acids by Pseudomonas sp. 42A2. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1997, 1347 (1), 75–81. 10.1016/S0005-2760(97)00056-8. [DOI] [PubMed] [Google Scholar]

[ref257] Busquets M.; Deroncelé V.; Vidal-Mas J.; Rodríguez E.; Guerrero A.; Manresa A. Isolation and Characterization of a Lipoxygenase from Pseudomonas 42A2 Responsible for the Biotransformation of Oleic Acid into (S)-(E)-10-Hydroxy-8-Octadecenoic Acid. Antonie Van Leeuwenhoek 2004, 85 (2), 129–139. 10.1023/B:ANTO.0000020152.15440.65. [DOI] [PubMed] [Google Scholar]

[ref258] Bhattarai S.; Niraula N. P.; Sohng J. K.; Oh T.-J. In-Silico and In-Vitro Based Studies of Streptomyces peucetius CYP107N3 for Oleic Acid Epoxidation. BMB Rep. 2012, 45 (12), 736–741. 10.5483/BMBRep.2012.45.12.080. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref259] Hintikka E.; Holopainen R.; Asola A.; Jestoi M.; Peitzsch M.; Kalso S.; Larsson L.; Reijula K.; Tuomi T. Mycotoxins in the Ventilation Systems of Four Schools in Finland. World Mycotoxin J. 2009, 2 (4), 369–379. 10.3920/WMJ2009.1155. [DOI] [Google Scholar]

[ref260] Niu M.; Keller N. P. Co-opting Oxylipin Signals in Microbial Disease. Cell. Microbiol. 2019, 21 (6), e13025 10.1111/cmi.13025. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref261] Tsitsigiannis D. I.; Keller N. P. Oxylipins as Developmental and Host-Fungal Communication Signals. Trends Microbiol. 2007, 15 (3), 109–118. 10.1016/j.tim.2007.01.005. [DOI] [PubMed] [Google Scholar]

[ref262] Brodhun F.; Feussner I. Oxylipins in Fungi. FEBS J. 2011, 278 (7), 1047–1063. 10.1111/j.1742-4658.2011.08027.x. [DOI] [PubMed] [Google Scholar]

[ref263] Christensen S. A.; Kolomiets M. V. The Lipid Language of Plant-Fungal Interactions. Fungal Genet. Biol. 2011, 48 (1), 4–14. 10.1016/j.fgb.2010.05.005. [DOI] [PubMed] [Google Scholar]

[ref264] Wurzenberger M.; Grosch W. Origin of the Oxygen in the Products of the Enzymatic Cleavage Reaction of Linoleic Acid to 1-Octen-3-Ol and 10-Oxo-Trans-8-Decenoic Acid in Mushrooms (Psalliota bispora). Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1984, 794 (1), 18–24. 10.1016/0005-2760(84)90292-3. [DOI] [Google Scholar]

[ref265] Wurzenberger M.; Grosch W. Stereochemistry of the Cleavage of the 10-Hydroperoxide Isomer of Linoleic Acid to 1-Octen-3-Ol by a Hydroperoxide Lyase from Mushrooms (Psalliota bispora). Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1984, 795 (1), 163–165. 10.1016/0005-2760(84)90117-6. [DOI] [Google Scholar]

[ref266] Wurzenberger M.; Grosch W. The Formation of 1-Octen-3-Ol from the 10-Hydroperoxide Isomer of Linoleic Acid by a Hydroperoxide Lyase in Mushrooms (Psalliota bispora). Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1984, 794 (1), 25–30. 10.1016/0005-2760(84)90293-5. [DOI] [Google Scholar]

[ref267] Wurzenberger M.; Grosch W. Enzymic Oxidation of Linolenic Acid to 1,Z-5-Octadien-3-Ol, Z-2,Z-5-Octadien-1-Ol and 10-Oxo-E-8-Decenoic Acid by a Protein Fraction from Mushrooms (Psalliota bispora). Lipids 1986, 21 (4), 261–266. 10.1007/BF02536408. [DOI] [PubMed] [Google Scholar]

[ref268] Brash A. R.; Niraula N. P.; Boeglin W. E.; Mashhadi Z. An Ancient Relative of Cyclooxygenase in Cyanobacteria Is a Linoleate 10S-Dioxygenase That Works in Tandem with a Catalase-Related Protein with Specific 10S-Hydroperoxide Lyase Activity. J. Biol. Chem. 2014, 289 (19), 13101–13111. 10.1074/jbc.M114.555904. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref269] Teshima T.; Funai R.; Nakazawa T.; Ito J.; Utsumi T.; Kakumyan P.; Mukai H.; Yoshiga T.; Murakami R.; Nakagawa K.; Honda Y.; Matsui K. Coprinopsis Cinerea Dioxygenase Is an Oxygenase Forming 10(S)-Hydroperoxide of Linoleic Acid, Essential for Mushroom Alcohol, 1-Octen-3-Ol, Synthesis. J. Biol. Chem. 2022, 298 (11), 102507. 10.1016/j.jbc.2022.102507. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref270] Oliw E. H. Fatty Acid Dioxygenase-Cytochrome P450 Fusion Enzymes of Filamentous Fungal Pathogens. Fungal Genet. Biol. 2021, 157, 103623. 10.1016/j.fgb.2021.103623. [DOI] [PubMed] [Google Scholar]

[ref271] Wadman M. W.; van Zadelhoff G.; Hamberg M.; Visser T.; Veldink G. A.; Vliegenthart J. F. G. Conversion of Linoleic Acid into Novel Oxylipins by the Mushroom Agaricus bisporus. Lipids 2005, 40 (11), 1163–1170. 10.1007/s11745-005-1481-2. [DOI] [PubMed] [Google Scholar]

[ref272] Hoffmann I.; Jernerén F.; Garscha U.; Oliw E. H. Expression of 5,8-LDS of Aspergillus fumigatus and Its Dioxygenase Domain: A Comparison with 7,8-LDS, 10-Dioxygenase, and Cyclooxygenase. Arch. Biochem. Biophys. 2011, 506 (2), 216–222. 10.1016/j.abb.2010.11.022. [DOI] [PubMed] [Google Scholar]

[ref273] Garscha U.; Jernerén F.; Chung D.; Keller N. P.; Hamberg M.; Oliw E. H. Identification of Dioxygenases Required for Aspergillus Development. J. Biol. Chem. 2007, 282 (48), 34707–34718. 10.1074/jbc.M705366200. [DOI] [PubMed] [Google Scholar]

[ref274] Su C.; Sahlin M.; Oliw E. H. A Protein Radical and Ferryl Intermediates Are Generated by Linoleate Diol Synthase, a Ferric Hemeprotein with Dioxygenase and Hydroperoxide Isomerase Activities. J. Biol. Chem. 1998, 273 (33), 20744–20751. 10.1074/jbc.273.33.20744. [DOI] [PubMed] [Google Scholar]

[ref275] Brodowsky I. D.; Oliw E. H. Metabolism of 18:2(n – 6), 18:3(n – 3), 20:4(n – 6) and 20:5(n – 3) by the Fungus Gaeumannomyces graminis: Identification of Metabolites Formed by 8-Hydroxylation and by W2 and W3 Oxygenation. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1992, 1124 (1), 59–65. 10.1016/0005-2760(92)90126-G. [DOI] [PubMed] [Google Scholar]

[ref276] Cristea M.; Osbourn A. E.; Oliw E. H. Linoleate Diol Synthase of the Rice Blast Fungus Magnaporthe grisea. Lipids 2003, 38 (12), 1275–1280. 10.1007/s11745-003-1189-3. [DOI] [PubMed] [Google Scholar]

[ref277] Jernerén F.; Garscha U.; Hoffmann I.; Hamberg M.; Oliw E. H. Reaction Mechanism of 5,8-Linoleate Diol Synthase, 10R-Dioxygenase, and 8,11-Hydroperoxide Isomerase of Aspergillus clavatus. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2010, 1801 (4), 503–507. 10.1016/j.bbalip.2009.12.012. [DOI] [PubMed] [Google Scholar]

[ref278] Shin K.-C.; Seo M.-J.; Oh D.-K. Characterization of a Novel 8R,11S-Linoleate Diol Synthase from Penicillium chrysogenum by Identification of Its Enzymatic Products. J. Lipid Res. 2016, 57 (2), 207–218. 10.1194/jlr.M061341. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref279] Hoffmann I.; Jernerén F.; Oliw E. H. Epoxy Alcohol Synthase of the Rice Blast Fungus Represents a Novel Subfamily of Dioxygenase-Cytochrome P450 Fusion Enzymes. J. Lipid Res. 2014, 55 (10), 2113–2123. 10.1194/jlr.M051755. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref280] Su C.; Oliw E. H. Manganese Lipoxygenase: Purification And Characterization. J. Biol. Chem. 1998, 273 (21), 13072–13079. 10.1074/jbc.273.21.13072. [DOI] [PubMed] [Google Scholar]

[ref281] Hamberg M.; Su C.; Oliw E. Manganese Lipoxygenase. Discovery Of A Bis-Allylic Hydroperoxide As Product And Intermediate In A Lipoxygenase Reaction. J. Biol. Chem. 1998, 273 (21), 13080–13088. 10.1074/jbc.273.21.13080. [DOI] [PubMed] [Google Scholar]

[ref282] Wennman A.; Oliw E. H. Secretion of Two Novel Enzymes, Manganese 9S-Lipoxygenase and Epoxy Alcohol Synthase, by the Rice Pathogen Magnaporthe salvinii. J. Lipid Res. 2013, 54 (3), 762–775. 10.1194/jlr.M033787. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref283] Brodhun F.; Cristobal-Sarramian A.; Zabel S.; Newie J.; Hamberg M.; Feussner I. An Iron 13S-Lipoxygenase with an α-Linolenic Acid Specific Hydroperoxidase Activity from Fusarium oxysporum. PLoS One 2013, 8 (5), e64919 10.1371/journal.pone.0064919. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref284] Teder T.; Boeglin W. E.; Schneider C.; Brash A. R. A Fungal Catalase Reacts Selectively with the 13S Fatty Acid Hydroperoxide Products of the Adjacent Lipoxygenase Gene and Exhibits 13S-Hydroperoxide-Dependent Peroxidase Activity. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2017, 1862 (7), 706–715. 10.1016/j.bbalip.2017.03.011. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref285] González-Benjumea A.; Carro J.; Renau-Mínguez C.; Linde D.; Fernández-Fueyo E.; Gutiérrez A.; Martínez A. T. Fatty Acid Epoxidation by Collariella virescens Peroxygenase and Heme-Channel Variants. Catal. Sci. Technol. 2020, 10 (3), 717–725. 10.1039/C9CY02332A. [DOI] [Google Scholar]

[ref286] Yin H.; Xu L.; Porter N. A. Free Radical Lipid Peroxidation: Mechanisms and Analysis. Chem. Rev. 2011, 111 (10), 5944–5972. 10.1021/cr200084z. [DOI] [PubMed] [Google Scholar]

[ref287] Viswanathan S.; Hammock B. D.; Newman J. W.; Meerarani P.; Toborek M.; Hennig B. Involvement of CYP 2C9 in Mediating the Proinflammatory Effects of Linoleic Acid in Vascular Endothelial Cells. J. Am. Coll. Nutr. 2003, 22 (6), 502–510. 10.1080/07315724.2003.10719328. [DOI] [PubMed] [Google Scholar]

[ref288] Kellogg E.; Fridovich I. Superoxide, Hydrogen Peroxide, and Singlet Oxygen in Lipid Peroxidation by a Xanthine Oxidase System. J. Biol. Chem. 1975, 250 (22), 8812–8817. 10.1016/S0021-9258(19)40745-X. [DOI] [PubMed] [Google Scholar]

[ref289] Li Y.; Mouche S.; Sajic T.; Veyrat-Durebex C.; Supale R.; Pierroz D.; Ferrari S.; Negro F.; Hasler U.; Feraille E.; Moll S.; Meda P.; Deffert C.; Montet X.; Krause K.-H.; Szanto I. Deficiency in the NADPH Oxidase 4 Predisposes towards Diet-Induced Obesity. Int. J. Obes. 2012, 36 (12), 1503–1513. 10.1038/ijo.2011.279. [DOI] [PubMed] [Google Scholar]

[ref290] Kurutas E. B. The Importance of Antioxidants Which Play the Role in Cellular Response against Oxidative/Nitrosative Stress: Current State. Nutr. J. 2015, 15 (1), 71. 10.1186/s12937-016-0186-5. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref291] Oh J. M.; Choi J. M.; Lee J. Y.; Oh S. J.; Kim B. H.; Kim S. K. Role of NADPH Oxidase-4 in Saturated Fatty Acid-Induced Insulin Resistance in SK-Hep-1 Cells. Food Chem. Toxicol. 2014, 63, 128–135. 10.1016/j.fct.2013.10.049. [DOI] [PubMed] [Google Scholar]

[ref292] Hatanaka E.; Dermargos A.; Hirata A. E.; Vinolo M. A. R.; Carpinelli A. R.; Newsholme P.; Armelin H. A.; Curi R. Oleic, Linoleic and Linolenic Acids Increase ROS Production by Fibroblasts via NADPH Oxidase Activation. PLoS One 2013, 8 (4), e58626 10.1371/journal.pone.0058626. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref293] Badwey J. A.; Curnutte J. T.; Karnovsky M. L. cis-Polyunsaturated Fatty Acids Induce High Levels of Superoxide Production by Human Neutrophils. J. Biol. Chem. 1981, 256 (24), 12640–12643. 10.1016/S0021-9258(18)42939-0. [DOI] [PubMed] [Google Scholar]

[ref294] Tyurina Y. Y.; Poloyac S. M.; Tyurin V. A.; Kapralov A. A.; Jiang J.; Anthonymuthu T. S.; Kapralova V. I.; Vikulina A. S.; Jung M.-Y.; Epperly M. W.; Mohammadyani D.; Klein-Seetharaman J.; Jackson T. C.; Kochanek P. M.; Pitt B. R.; Greenberger J. S.; Vladimirov Y. A.; Bayır H.; Kagan V. E. A Mitochondrial Pathway for Biosynthesis of Lipid Mediators. Nat. Chem. 2014, 6 (6), 542–552. 10.1038/nchem.1924. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref295] Sies H.; Berndt C.; Jones D. P. Oxidative Stress. Annu. Rev. Biochem. 2017, 86 (1), 715–748. 10.1146/annurev-biochem-061516-045037. [DOI] [PubMed] [Google Scholar]

[ref296] Gutteridge J. M. C.; Bannister J. V. Copper + Zinc and Manganese Superoxide Dismutases Inhibit Deoxyribose Degradation by the Superoxide-Driven Fenton Reaction at Two Different Stages. Implications for the Redox States of Copper and Manganese. Biochem. J. 1986, 234 (1), 225–228. 10.1042/bj2340225. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref297] Pryor W. A. Oxy-Radicals and Related Species: Their Formation, Lifetimes, and Reactions. Annu. Rev. Physiol. 1986, 48, 657–667. 10.1146/annurev.ph.48.030186.003301. [DOI] [PubMed] [Google Scholar]

[ref298] Wilcox A. L.; Marnett L. J. Polyunsaturated Fatty Acid Alkoxyl Radicals Exist as Carbon-Centered Epoxyallylic Radicals: A Key Step in Hydroperoxide-Amplified Lipid Peroxidation. Chem. Res. Toxicol. 1993, 6 (4), 413–416. 10.1021/tx00034a003. [DOI] [PubMed] [Google Scholar]

[ref299] Gardner H. W. Oxygen Radical Chemistry of Polyunsaturated Fatty Acids. Free Radic. Biol. Med. 1989, 7 (1), 65–86. 10.1016/0891-5849(89)90102-0. [DOI] [PubMed] [Google Scholar]

[ref300] Spiteller G. Linoleic Acid Peroxidation-the Dominant Lipid Peroxidation Process in Low Density Lipoprotein-and Its Relationship to Chronic Diseases. Chem. Phys. Lipids 1998, 95 (2), 105–162. 10.1016/S0009-3084(98)00091-7. [DOI] [PubMed] [Google Scholar]

[ref301] Kaplan E.; Ansari K. Reduction of Polyunsaturated Fatty Acid Hydroperoxides by Human Brain Glutathione Peroxidase. Lipids 1984, 19 (10), 784–789. 10.1007/BF02534472. [DOI] [PubMed] [Google Scholar]

[ref302] Christophersen B. O. Formation of Monohydroxy-Polyenic Fatty Acids from Lipid Peroxides by a Glutathione Peroxidase. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1968, 164 (1), 35–46. 10.1016/0005-2760(68)90068-4. [DOI] [PubMed] [Google Scholar]

[ref303] Little C.; O’Brien P. J. An Intracellular GSH-Peroxidase with a Lipid Peroxide Substrate. Biochem. Biophys. Res. Commun. 1968, 31 (2), 145–150. 10.1016/0006-291X(68)90721-3. [DOI] [PubMed] [Google Scholar]

[ref304] Yoshida Y.; Umeno A.; Akazawa Y.; Shichiri M.; Murotomi K.; Horie M. Chemistry of Lipid Peroxidation Products and Their Use as Biomarkers in Early Detection of Diseases. J. Oleo Sci. 2015, 64 (4), 347–356. 10.5650/jos.ess14281. [DOI] [PubMed] [Google Scholar]

[ref305] Frankel E. N.; Neff W. E.; Rohwedder W. K.; Khambay B. P. S.; Garwood R. F.; Weedon B. C. L. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry: II. Methyl Linoleate. Lipids 1977, 12 (11), 908–913. 10.1007/BF02533310. [DOI] [PubMed] [Google Scholar]

[ref306] Frankel E. N.; Neff W. E.; Rohwedder W. K.; Khambay B. P. S.; Garwood R. F.; Weedon B. C. L. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry: III. Methyl Linolenate. Lipids 1977, 12 (12), 1055–1061. 10.1007/BF02533334. [DOI] [PubMed] [Google Scholar]

[ref307] Frankel E. N.; Neff W. E.; Rohwedder W. K.; Khambay B. P. S.; Garwood R. F.; Weedon B. C. L. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry: I. Methyl Oleate. Lipids 1977, 12 (11), 901–907. 10.1007/BF02533309. [DOI] [PubMed] [Google Scholar]

[ref308] Watabe N.; Ishida Y.; Ochiai A.; Tokuoka Y.; Kawashima N. Oxidation Decomposition of Unsaturated Fatty Acids by Singlet Oxygen in Phospholipid Bilayer Membranes. J. Oleo Sci. 2007, 56 (2), 73–80. 10.5650/jos.56.73. [DOI] [PubMed] [Google Scholar]

[ref309] Frimer A. A. The Reaction of Singlet Oxygen with Olefins: The Question of Mechanism. Chem. Rev. 1979, 79 (5), 359–387. 10.1021/cr60321a001. [DOI] [Google Scholar]

[ref310] Minami Y.; Yokoyama K.; Bando N.; Kawai Y.; Terao J. Occurrence of Singlet Oxygen Oxygenation of Oleic Acid and Linoleic Acid in the Skin of Live Mice. Free Radic. Res. 2008, 42 (3), 197–204. 10.1080/10715760801948088. [DOI] [PubMed] [Google Scholar]

[ref311] Frankel E. N.; Neff W. E.; Selke E. Analysis of Autoxidized Fats by Gas Chromatography-Mass Spectrometry. IX. Homolytic vs. Heterolytic Cleavage of Primary and Secondary Oxidation Products. Lipids 1984, 19 (10), 790–800. 10.1007/BF02534473. [DOI] [Google Scholar]

[ref312] Radi R.; Beckman J. S.; Bush K. M.; Freeman B. A. Peroxynitrite-Induced Membrane Lipid Peroxidation: The Cytotoxic Potential of Superoxide and Nitric Oxide. Arch. Biochem. Biophys. 1991, 288 (2), 481–487. 10.1016/0003-9861(91)90224-7. [DOI] [PubMed] [Google Scholar]

[ref313] Rubbo H.; Parthasarathy S.; Barnes S.; Kirk M.; Kalyanaraman B.; Freeman B. A. Nitric Oxide Inhibition of Lipoxygenase-Dependent Liposome and Low-Density Lipoprotein Oxidation: Termination of Radical Chain Propagation Reactions and Formation of Nitrogen-Containing Oxidized Lipid Derivatives. Arch. Biochem. Biophys. 1995, 324 (1), 15–25. 10.1006/abbi.1995.9935. [DOI] [PubMed] [Google Scholar]

[ref314] Freeman B. A.; Baker P. R. S.; Schopfer F. J.; Woodcock S. R.; Napolitano A.; d’Ischia M. Nitro-Fatty Acid Formation and Signaling. J. Biol. Chem. 2008, 283 (23), 15515–15519. 10.1074/jbc.R800004200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref315] Cantu-Medellin N.; Kelley E. E. Xanthine Oxidoreductase-Catalyzed Reduction of Nitrite to Nitric Oxide: Insights Regarding Where, When and How. Nitric Oxide 2013, 34, 19–26. 10.1016/j.niox.2013.02.081. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref316] Kim-Shapiro D. B.; Schechter A. N.; Gladwin M. T. Unraveling the Reactions of Nitric Oxide, Nitrite, and Hemoglobin in Physiology and Therapeutics. Arterioscler. Thromb. Vasc. Biol. 2006, 26 (4), 697–705. 10.1161/01.ATV.0000204350.44226.9a. [DOI] [PubMed] [Google Scholar]

[ref317] Spiteller P.; Spiteller G. 9-Hydroxy-10,12-Octadecadienoic Acid (9-HODE) and 13-Hydroxy-9,11-Octadecadienoic Acid (13-HODE): Excellent Markers for Lipid Peroxidation. Chem. Phys. Lipids 1997, 89 (2), 131–139. 10.1016/S0009-3084(97)00070-4. [DOI] [Google Scholar]

[ref318] Bergström S.; Holman R. T. Total Conjugation of Linoleic Acid in Oxidation with Lipoxidase. Nature 1948, 161 (4080), 55. 10.1038/161055a0. [DOI] [PubMed] [Google Scholar]

[ref319] Bergström S.; Holman R. T.. Lipoxidase and the Autoxidation of Unsaturated Fatty Acids. In Advances in Enzymology and Related Areas of Molecular Biology; John Wiley & Sons, 1948; pp 425–457. 10.1002/9780470122532.ch9 [DOI] [Google Scholar]

[ref320] Russell G. A. Deuterium-Isotope Effects in the Autoxidation of Aralkyl Hydrocarbons. Mechanism of the Interaction of Peroxy Radicals. J. Am. Chem. Soc. 1957, 79 (14), 3871–3877. 10.1021/ja01571a068. [DOI] [Google Scholar]

[ref321] Kanofsky J. R. Singlet Oxygen Production from the Reactions of Alkylperoxy Radicals. Evidence from 1268-nm Chemiluminescence. J. Org. Chem. 1986, 51 (17), 3386–3388. 10.1021/jo00367a032. [DOI] [Google Scholar]

[ref322] Miyamoto S.; Martinez G. R.; Medeiros M. H. G.; Di Mascio P. Singlet Molecular Oxygen Generated from Lipid Hydroperoxides by the Russell Mechanism: Studies Using 18(O)-Labeled Linoleic Acid Hydroperoxide and Monomol Light Emission Measurements. J. Am. Chem. Soc. 2003, 125 (20), 6172–6179. 10.1021/ja029115o. [DOI] [PubMed] [Google Scholar]

[ref323] Miyamoto S.; Martinez G. R.; Rettori D.; Augusto O.; Medeiros M. H. G.; Di Mascio P. Linoleic Acid Hydroperoxide Reacts with Hypochlorous Acid, Generating Peroxyl Radical Intermediates and Singlet Molecular Oxygen. Proc. Natl. Acad. Sci. U. S. A. 2006, 103 (2), 293–298. 10.1073/pnas.0508170103. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref324] Takigawa Y.; Koshiishi I. Catalytic Production of Oxo-Fatty Acids by Lipoxygenases Is Mediated by the Radical-Radical Dismutation between Fatty Acid Alkoxyl Radicals and Fatty Acid Peroxyl Radicals in Fatty Acid Assembly. Chem. Pharm. Bull. (Tokyo) 2020, 68 (3), 258–264. 10.1248/cpb.c19-00975. [DOI] [PubMed] [Google Scholar]

[ref325] Zhu X.; Tang X.; Anderson V. E.; Sayre L. M. Mass Spectrometric Characterization of Protein Modification by the Products of Nonenzymatic Oxidation of Linoleic Acid. Chem. Res. Toxicol. 2009, 22 (8), 1386–1397. 10.1021/tx9000072. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref326] Labeque R.; Marnett L. J. Reaction of Hematin with Allylic Fatty Acid Hydroperoxides: Identification of Products and Implications for Pathways of Hydroperoxide-Dependent Epoxidation of 7,8-Dihydroxy-7,8-Dihydrobenzo[a]Pyrene. Biochemistry 1988, 27 (18), 7060–7070. 10.1021/bi00418a059. [DOI] [PubMed] [Google Scholar]

[ref327] Dix T. A.; Marnett L. J. Conversion of Linoleic Acid Hydroperoxide to Hydroxy, Keto, Epoxyhydroxy, and Trihydroxy Fatty Acids by Hematin. J. Biol. Chem. 1985, 260 (9), 5351–5357. 10.1016/S0021-9258(18)89028-7. [DOI] [PubMed] [Google Scholar]

[ref328] Neff W. E.; Frankel E. N.; Selke E.; Weisleder D. Photosensitized Oxidation of Methyl Linoleate Monohydroperoxides: Hydroperoxy Cyclic Peroxides, Dihydroperoxides, Keto Esters and Volatile Thermal Decomposition Products. Lipids 1983, 18 (12), 868–876. 10.1007/BF02534564. [DOI] [Google Scholar]

[ref329] Hamberg M. A Novel Transformation of 13-Ls-Hydroperoxy-9,11-Octadecadienoic acid. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1983, 752 (2), 191–197. 10.1016/0005-2760(83)90112-1. [DOI] [PubMed] [Google Scholar]

[ref330] Hamberg M. Autoxidation of Linoleic Acid: Isolation and Structure of Four Dihydroxyoctadecadienoic Acids. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1983, 752 (2), 353–356. 10.1016/0005-2760(83)90134-0. [DOI] [Google Scholar]

[ref331] Niisuke K.; Boeglin W. E.; Murray J. J.; Schneider C.; Brash A. R. Biosynthesis of a Linoleic Acid Allylic Epoxide: Mechanistic Comparison with Its Chemical Synthesis and Leukotriene A Biosynthesis. J. Lipid Res. 2009, 50 (7), 1448–1455. 10.1194/jlr.M900025-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref332] Frankel E. N.; Neff W. E.; Selke E.; Weisleder D. Photosensitized Oxidation of Methyl Linoleate: Secondary and Volatile Thermal Decomposition Products. Lipids 1982, 17 (1), 11–18. 10.1007/BF02535116. [DOI] [PubMed] [Google Scholar]

[ref333] Neff W. E.; Frankel E. N.; Weisleder D. Photosensitized Oxidation of Methyl Linolenate. Secondary Products. Lipids 1982, 17 (11), 780–790. 10.1007/BF02535354. [DOI] [PubMed] [Google Scholar]

[ref334] Hamberg M.; Gotthammar B. A New Reaction of Unsaturated Fatty Acid Hydroperoxides: Formation of 11-Hydroxy-12,13-Epoxy-9-Octadecenoic Acid from 13-Hydroperoxy-9,11-Octadecadienoic Acid. Lipids 1973, 8 (12), 737–744. 10.1007/BF02531842. [DOI] [Google Scholar]

[ref335] Gardner H. W.; Kleiman R. Degradation of Linoleic Acid Hydroperoxides by a Cysteine FeCl3 Catalyst as a Model for Similar Biochemical Reactions: II. Specificity in Formation of Fatty Acid Epoxides. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1981, 665 (1), 113–125. 10.1016/0005-2760(81)90239-3. [DOI] [PubMed] [Google Scholar]

[ref336] Gardner H. W.; Plattner R. D.; Weisleder D. The Epoxyallylic Radical from Homolysis and Rearrangement of Methyl Linoleate Hydroperoxide Combines with the Thiyl Radical of N-Acetylcysteine. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1985, 834 (1), 65–74. 10.1016/0005-2760(85)90177-8. [DOI] [Google Scholar]

[ref337] Gardner H. W.; Weisleder D.; Kleiman R. Formation of trans-12,13-Epoxy-9-Hydroperoxy-trans-10-Octadecenoic Acid from 13-L-Hydroperoxy-cis-9, trans-11-Octadecadienoic Acid Catalyzed by Either a Soybean Extract or Cysteine-FeCl3. Lipids 1978, 13 (4), 246–252. 10.1007/BF02533664. [DOI] [PubMed] [Google Scholar]

[ref338] Gardner H. W.; Selke E. Volatiles from Thermal Decomposition of Isomeric Methyl (12S, 13S)-(E)-12,13-Epoxy-9-Hydroperoxy-10-Octadecenoates. Lipids 1984, 19 (6), 375–380. 10.1007/BF02537398. [DOI] [Google Scholar]

[ref339] Lin D.; Zhang J.; Sayre L. M. Synthesis of Six Epoxyketooctadecenoic Acid (EKODE) Isomers, Their Generation from Nonenzymatic Oxidation of Linoleic Acid, and Their Reactivity with Imidazole Nucleophiles. J. Org. Chem. 2007, 72 (25), 9471–9480. 10.1021/jo701373f. [DOI] [PubMed] [Google Scholar]

[ref340] Szabó C.; Ischiropoulos H.; Radi R. Peroxynitrite: Biochemistry, Pathophysiology and Development of Therapeutics. Nat. Rev. Drug Discovery 2007, 6 (8), 662–680. 10.1038/nrd2222. [DOI] [PubMed] [Google Scholar]

[ref341] Lundberg J. O.; Gladwin M. T.; Ahluwalia A.; Benjamin N.; Bryan N. S.; Butler A.; Cabrales P.; Fago A.; Feelisch M.; Ford P. C.; Freeman B. A.; Frenneaux M.; Friedman J.; Kelm M.; Kevil C. G.; Kim-Shapiro D. B.; Kozlov A. V.; Lancaster J. R.; Lefer D. J.; McColl K.; McCurry K.; Patel R. P.; Petersson J.; Rassaf T.; Reutov V. P.; Richter-Addo G. B.; Schechter A.; Shiva S.; Tsuchiya K.; van Faassen E. E.; Webb A. J.; Zuckerbraun B. S.; Zweier J. L.; Weitzberg E. Nitrate and Nitrite in Biology, Nutrition and Therapeutics. Nat. Chem. Biol. 2009, 5 (12), 865–869. 10.1038/nchembio.260. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref342] Trujillo M.; Folkes L.; Bartesaghi S.; Kalyanaraman B.; Wardman P.; Radi R. Peroxynitrite-Derived Carbonate and Nitrogen Dioxide Radicals Readily React with Lipoic and Dihydrolipoic Acid. Free Radic. Biol. Med. 2005, 39 (2), 279–288. 10.1016/j.freeradbiomed.2005.03.014. [DOI] [PubMed] [Google Scholar]

[ref343] Meli R.; Nauser T.; Latal P.; Koppenol W. H. Reaction of Peroxynitrite with Carbon Dioxide: Intermediates and Determination of the Yield of CO3•- and NO2•. J. Biol. Inorg. Chem. 2002, 7 (1), 31–36. 10.1007/s007750100262. [DOI] [PubMed] [Google Scholar]

[ref344] Baker P. R. S.; Lin Y.; Schopfer F. J.; Woodcock S. R.; Groeger A. L.; Batthyany C.; Sweeney S.; Long M. H.; Iles K. E.; Baker L. M. S.; Branchaud B. P.; Chen Y. E.; Freeman B. A. Fatty Acid Transduction of Nitric Oxide Signaling: Multiple Nitrated Unsaturated Fatty Acid Derivatives Exist in Human Blood and Urine and Serve As Endogenous Peroxisome Proliferator-Activated Receptor Ligands. J. Biol. Chem. 2005, 280 (51), 42464–42475. 10.1074/jbc.M504212200. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref345] Baker P. R. S.; Schopfer F. J.; Sweeney S.; Freeman B. A. Red Cell Membrane and Plasma Linoleic Acid Nitration Products: Synthesis, Clinical Identification, and Quantitation. Proc. Natl. Acad. Sci. U. S. A. 2004, 101 (32), 11577–11582. 10.1073/pnas.0402587101. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref346] Buchan G. J.; Bonacci G.; Fazzari M.; Salvatore S. R.; Gelhaus Wendell S. Nitro-Fatty Acid Formation and Metabolism. Nitric Oxide 2018, 79, 38–44. 10.1016/j.niox.2018.07.003. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref347] Napolitano A.; Camera E.; Picardo M.; d’Ischia M. Acid-Promoted Reactions of Ethyl Linoleate with Nitrite Ions: Formation and Structural Characterization of Isomeric Nitroalkene, Nitrohydroxy, and Novel 3-Nitro-1,5-Hexadiene and 1,5-Dinitro-1,3-Pentadiene Products. J. Org. Chem. 2000, 65 (16), 4853–4860. 10.1021/jo000090q. [DOI] [PubMed] [Google Scholar]

[ref348] Onyango A. N. Endogenous Generation of Singlet Oxygen and Ozone in Human and Animal Tissues: Mechanisms, Biological Significance, and Influence of Dietary Components. Oxid. Med. Cell. Longev. 2016, 2016, 1–22. 10.1155/2016/2398573. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref349] Ahmed O. S.; Galano J.-M.; Pavlickova T.; Revol-Cavalier J.; Vigor C.; Lee J. C.-Y.; Oger C.; Durand T. Moving Forward with Isoprostanes, Neuroprostanes and Phytoprostanes: Where Are We Now?. Essays Biochem. 2020, 64 (3), 463–484. 10.1042/EBC20190096. [DOI] [PubMed] [Google Scholar]

[ref350] Cuyamendous C.; de la Torre A.; Lee Y. Y.; Leung K. S.; Guy A.; Bultel-Poncé V.; Galano J.-M.; Lee J. C.-Y.; Oger C.; Durand T. The Novelty of Phytofurans, Isofurans, Dihomo-Isofurans and Neurofurans: Discovery, Synthesis and Potential Application. Biochimie 2016, 130, 49–62. 10.1016/j.biochi.2016.08.002. [DOI] [PubMed] [Google Scholar]

[ref351] Samuelsson B. On the Incorporation of Oxygen in the Conversion of 8,11,14-Eicosatrienoic Acid to Prostaglandin E ₁. J. Am. Chem. Soc. 1965, 87 (13), 3011–3013. 10.1021/ja01091a043. [DOI] [PubMed] [Google Scholar]

[ref352] Hamberg M.; Svensson J.; Wakabayashi T.; Samuelsson B. Isolation and Structure of Two Prostaglandin Endoperoxides That Cause Platelet Aggregation. Proc. Natl. Acad. Sci. U. S. A. 1974, 71 (2), 345–349. 10.1073/pnas.71.2.345. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref353] Porter N. A.; Funk M. O. Peroxy Radical Cyclization as a Model for Prostaglandin Biosynthesis. J. Org. Chem. 1975, 40 (24), 3614–3615. 10.1021/jo00912a037. [DOI] [PubMed] [Google Scholar]

[ref354] Porter N. A.; Funk M. O.; Gilmore D.; Isaac R.; Nixon J. The Formation of Cyclic Peroxides from Unsaturated Hydroperoxides: Models for Prostaglandin Biosynthesis. J. Am. Chem. Soc. 1976, 98, 6000–6005. 10.1021/ja00435a037. [DOI] [PubMed] [Google Scholar]

[ref355] Pryor W. A.; Porter N. A. Suggested Mechanisms for the Production of 4-Hydroxy-2-Nonenal from the Autoxidation of Polyunsaturated Fatty Acids. Free Radic. Biol. Med. 1990, 8 (6), 541–543. 10.1016/0891-5849(90)90153-A. [DOI] [PubMed] [Google Scholar]

[ref356] Roza M.; Francke A. Cyclic Peroxides from a Soya Lipoxygenase-Catalysed Oxygenation of Methyl Linolenate. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1978, 528 (1), 119–126. 10.1016/0005-2760(78)90057-7. [DOI] [PubMed] [Google Scholar]

[ref357] Mihelich E. D. Structure and Stereochemistry of Novel Endoperoxides Isolated from the Sensitized Photooxidation of Methyl Linoleate. Implications for Prostaglandin Biosynthesis. J. Am. Chem. Soc. 1980, 102 (23), 7141–7143. 10.1021/ja00543a061. [DOI] [Google Scholar]

[ref358] Chan H. W.-S.; Matthew J. A.; Coxon D. T. A Hydroperoxy-Epidioxide from the Autoxidation of a Hydroperoxide of Methyl Linolenate. J. Chem. Soc. Chem. Commun. 1980, (5), 235–236. 10.1039/c39800000235. [DOI] [Google Scholar]

[ref359] Neff W. E.; Frankel E. N.; Weisleder D. High Pressure Liquid Chromatography of Autoxidized Lipids: II. Hydroperoxy-Cyclic Peroxides and Other Secondary Products from Methyl Linolenate. Lipids 1981, 16 (6), 439–448. 10.1007/BF02535012. [DOI] [Google Scholar]

[ref360] Coxon D. T.; Price K. R.; Chan H. W.-S. Formation, Isolation and Structure Determination of Methyl Linolenate Diperoxides. Chem. Phys. Lipids 1981, 28 (4), 365–378. 10.1016/0009-3084(81)90022-0. [DOI] [Google Scholar]

[ref361] Gunstone F. D.; Wijesundera R. C. Fatty Acids, Part 54. Some Reactions of Long-Chain Oxygenated Acids with Special Reference to Those Furnishing Furanoid Acids. Chem. Phys. Lipids 1979, 24 (2), 193–208. 10.1016/0009-3084(79)90088-4. [DOI] [Google Scholar]

[ref362] Barden A. E.; Croft K. D.; Durand T.; Guy A.; Mueller M. J.; Mori T. A. Flaxseed Oil Supplementation Increases Plasma F₁-Phytoprostanes in Healthy Men. J. Nutr. 2009, 139 (10), 1890–1895. 10.3945/jn.109.108316. [DOI] [PubMed] [Google Scholar]

[ref363] Dahle L. K.; Hill E. G.; Holman R. T. The Thiobarbituric Acid Reaction and the Autoxidations of Polyunsaturated Fatty Acid Methyl Esters. Arch. Biochem. Biophys. 1962, 98 (2), 253–261. 10.1016/0003-9861(62)90181-9. [DOI] [PubMed] [Google Scholar]

[ref364] Porter N. A. Mechanisms for the Autoxidation of Polyunsaturated Lipids. Acc. Chem. Res. 1986, 19 (9), 262–268. 10.1021/ar00129a001. [DOI] [Google Scholar]

[ref365] Corey E. J.; Shimoji K.; Shih C. Synthesis of Prostaglandins via a 1,2-Dioxabicyclo[2.2.1]Heptane (Endoperoxide) Intermediate. Stereochemical Divergence of Enzymic and Biomimetic Chemical Cyclization Reactions. J. Am. Chem. Soc. 1984, 106 (21), 6425–6427. 10.1021/ja00333a057. [DOI] [Google Scholar]

[ref366] O’Connor D. E.; Mihelich E. D.; Coleman M. C. Isolation and Characterization of Bicycloendoperoxides Derived from Methyl Linolenate. J. Am. Chem. Soc. 1981, 103 (1), 223–224. 10.1021/ja00391a056. [DOI] [Google Scholar]

[ref367] O’Connor D. E.; Mihelich E. D.; Coleman M. C. Stereochemical Course of the Autooxidative Cyclization of Lipid Hydroperoxides to Prostaglandin-like Bicyclic Endoperoxides. J. Am. Chem. Soc. 1984, 106 (12), 3577–3584. 10.1021/ja00324a028. [DOI] [Google Scholar]

[ref368] Montine T. J.; Montine K. S.; Reich E. E.; Terry E. S.; Porter N. A.; Morrow J. D. Antioxidants Significantly Affect the Formation of Different Classes of Isoprostanes and Neuroprostanes in Rat Cerebral Synaptosomes. Biochem. Pharmacol. 2003, 65 (4), 611–617. 10.1016/S0006-2952(02)01607-6. [DOI] [PubMed] [Google Scholar]

[ref369] Nugteren D. H.; Hazelhof E. Isolation and Properties of Intermediates in Prostaglandin Biosynthesis. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1973, 326 (3), 448–461. 10.1016/0005-2760(73)90145-8. [DOI] [PubMed] [Google Scholar]

[ref370] Porter N. A.; Byers J. D.; Mebane R. C.; Gilmore D. W.; Nixon J. R. Prostaglandin H₂ Methyl Ester. J. Org. Chem. 1978, 43 (10), 2088–2090. 10.1021/jo00404a067. [DOI] [Google Scholar]

[ref371] Chen Y.; Morrow J. D.; Roberts L. J. Formation of Reactive Cyclopentenone Compounds in Vivo as Products of the Isoprostane Pathway. J. Biol. Chem. 1999, 274 (16), 10863–10868. 10.1074/jbc.274.16.10863. [DOI] [PubMed] [Google Scholar]

[ref372] Thoma I.; Loeffler C.; Sinha A. K.; Gupta M.; Krischke M.; Steffan B.; Roitsch T.; Mueller M. J. Cyclopentenone Isoprostanes Induced by Reactive Oxygen Species Trigger Defense Gene Activation and Phytoalexin Accumulation in Plants. Plant J. 2003, 34 (3), 363–375. 10.1046/j.1365-313X.2003.01730.x. [DOI] [PubMed] [Google Scholar]

[ref373] Fessel J. P.; Porter N. A.; Moore K. P.; Sheller J. R.; Roberts L. J. Discovery of Lipid Peroxidation Products Formed in Vivo with a Substituted Tetrahydrofuran Ring (Isofurans) That Are Favored by Increased Oxygen Tension. Proc. Natl. Acad. Sci. U. S. A. 2002, 99 (26), 16713–16718. 10.1073/pnas.252649099. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref374] Roberts L. J.; Fessel J. P. The Biochemistry of the Isoprostane, Neuroprostane, and Isofuran Pathways of Lipid Peroxidation. Chem. Phys. Lipids 2004, 128 (1), 173–186. 10.1016/j.chemphyslip.2003.09.016. [DOI] [PubMed] [Google Scholar]

[ref375] Shearer G. C.; Newman J. W. Lipoprotein Lipase Releases Esterified Oxylipins from Very Low-Density Lipoproteins. Prostaglandins Leukot. Essent. Fatty Acids 2008, 79 (6), 215–222. 10.1016/j.plefa.2008.09.023. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref376] Liang N.; Harsch B. A.; Zhou S.; Borkowska A.; Shearer G. C.; Kaddurah-Daouk R.; Newman J. W.; Borkowski K. Oxylipin Transport by Lipoprotein Particles and Its Functional Implications for Cardiometabolic and Neurological Disorders. Prog. Lipid Res. 2024, 93, 101265. 10.1016/j.plipres.2023.101265. [DOI] [PubMed] [Google Scholar]

[ref377] O’Donnell V. B.; Murphy R. C. New Families of Bioactive Oxidized Phospholipids Generated by Immune Cells: Identification and Signaling Actions. Blood 2012, 120 (10), 1985–1992. 10.1182/blood-2012-04-402826. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref378] Fazzari M.; Vitturi D. A.; Woodcock S. R.; Salvatore S. R.; Freeman B. A.; Schopfer F. J. Electrophilic Fatty Acid Nitroalkenes Are Systemically Transported and Distributed upon Esterification to Complex Lipids. J. Lipid Res. 2019, 60 (2), 388–399. 10.1194/jlr.M088815. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref379] Fazzari M.; Khoo N. H.; Woodcock S. R.; Jorkasky D. K.; Li L.; Schopfer F. J.; Freeman B. A. Nitro-Fatty Acid Pharmacokinetics in the Adipose Tissue Compartment. J. Lipid Res. 2017, 58 (2), 375–385. 10.1194/jlr.M072058. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref380] Annevelink C. E.; Walker R. E.; Shearer G. C. Esterified Oxylipins: Do They Matter?. Metabolites 2022, 12 (11), 1007. 10.3390/metabo12111007. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref381] Carpanedo L.; Rund K. M.; Wende L. M.; Kampschulte N.; Schebb N. H. LC-HRMS Analysis of Phospholipids Bearing Oxylipins. Analytica Chimica Acta 2024, 1326, 343139. 10.1016/j.aca.2024.343139. [DOI] [PubMed] [Google Scholar]

[ref382] Genva M.; Obounou Akong F.; Andersson M. X.; Deleu M.; Lins L.; Fauconnier M.-L. New Insights into the Biosynthesis of Esterified Oxylipins and Their Involvement in Plant Defense and Developmental Mechanisms. Phytochem. Rev. 2019, 18 (1), 343–358. 10.1007/s11101-018-9595-8. [DOI] [Google Scholar]

[ref383] Astudillo A. M.; Balgoma D.; Balboa M. A.; Balsinde J. Dynamics of Arachidonic Acid Mobilization by Inflammatory Cells. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2012, 1821 (2), 249–256. 10.1016/j.bbalip.2011.11.006. [DOI] [PubMed] [Google Scholar]

[ref384] Samuelsson B. Role of Basic Science in the Development of New Medicines: Examples from the Eicosanoid Field. J. Biol. Chem. 2012, 287 (13), 10070–10080. 10.1074/jbc.X112.351437. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref385] Kobe M. J.; Neau D. B.; Mitchell C. E.; Bartlett S. G.; Newcomer M. E. The Structure of Human 15-Lipoxygenase-2 with a Substrate Mimic. J. Biol. Chem. 2014, 289 (12), 8562–8569. 10.1074/jbc.M113.543777. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref386] Belkner J.; Stender H.; Kühn H. The Rabbit 15-Lipoxygenase Preferentially Oxygenates LDL Cholesterol Esters, and This Reaction Does Not Require Vitamin E. J. Biol. Chem. 1998, 273 (36), 23225–23232. 10.1074/jbc.273.36.23225. [DOI] [PubMed] [Google Scholar]

[ref387] O’Donnell V. B. New Appreciation for an Old Pathway: The Lands Cycle Moves into New Arenas in Health and Disease. Biochem. Soc. Trans. 2022, 50 (1), 1–11. 10.1042/BST20210579. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref388] Yuan H.; Li X.; Zhang X.; Kang R.; Tang D. Identification of ACSL4 as a Biomarker and Contributor of Ferroptosis. Biochem. Biophys. Res. Commun. 2016, 478 (3), 1338–1343. 10.1016/j.bbrc.2016.08.124. [DOI] [PubMed] [Google Scholar]

[ref389] Klett E. L.; Chen S.; Yechoor A.; Lih F. B.; Coleman R. A. Long-Chain Acyl-CoA Synthetase Isoforms Differ in Preferences for Eicosanoid Species and Long-Chain Fatty Acids. J. Lipid Res. 2017, 58 (5), 884–894. 10.1194/jlr.M072512. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref390] Kanter J. E.; Tang C.; Oram J. F.; Bornfeldt K. E. Acyl-CoA Synthetase 1 Is Required for Oleate and Linoleate Mediated Inhibition of Cholesterol Efflux through ATP-Binding Cassette Transporter A1 in Macrophages. Biochim. Biophys. Acta BBA - Mol. Cell Biol. Lipids 2012, 1821 (3), 358–364. 10.1016/j.bbalip.2011.10.008. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref391] Golej D. L.; Askari B.; Kramer F.; Barnhart S.; Vivekanandan-Giri A.; Pennathur S.; Bornfeldt K. E. Long-Chain Acyl-CoA Synthetase 4 Modulates Prostaglandin E₂ Release from Human Arterial Smooth Muscle Cells. J. Lipid Res. 2011, 52 (4), 782–793. 10.1194/jlr.M013292. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref392] Walker R. E.; Richter C. K.; Skulas-Ray A. C.; Flock M. R.; Harsch B. A.; Annevelink C. E.; Kris-Etherton P. M.; Jensen G. L.; Shearer G. C. Effect of Omega-3 Ethyl Esters on the Triglyceride-Rich Lipoprotein Response to Endotoxin Challenge in Healthy Young Men. J. Lipid Res. 2023, 64 (5), 100353. 10.1016/j.jlr.2023.100353. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref393] Wopereis S.; Wolvers D.; van Erk M.; Gribnau M.; Kremer B.; van Dorsten F. A.; Boelsma E.; Garczarek U.; Cnubben N.; Frenken L.; van der Logt P.; Hendriks H. F. J.; Albers R.; van Duynhoven J.; van Ommen B.; Jacobs D. M. Assessment of Inflammatory Resilience in Healthy Subjects Using Dietary Lipid and Glucose Challenges. BMC Med. Genomics 2013, 6, 44. 10.1186/1755-8794-6-44. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref394] Wright D. N.; Katundu K. G. H.; Viscarra J. A.; Crocker D. E.; Newman J. W.; La Frano M. R.; Ortiz R. M. Oxylipin Responses to Fasting and Insulin Infusion in a Large Mammalian Model of Fasting-Induced Insulin Resistance, the Northern Elephant Seal. Am. J. Physiol. - Regul. Integr. Comp. Physiol. 2021, 321 (4), R537–R546. 10.1152/ajpregu.00016.2021. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref395] Shearer G. C.; Borkowski K.; Puumala S. L.; Harris W. S.; Pedersen T. L.; Newman J. W. Abnormal Lipoprotein Oxylipins in Metabolic Syndrome and Partial Correction by Omega-3 Fatty Acids. Prostaglandins Leukot. Essent. Fatty Acids 2018, 128, 1–10. 10.1016/j.plefa.2017.10.006. [DOI] [PubMed] [Google Scholar]

[ref396] Schebb N. H.; Ostermann A. I.; Yang J.; Hammock B. D.; Hahn A.; Schuchardt J. P. Comparison of the Effects of Long-Chain Omega-3 Fatty Acid Supplementation on Plasma Levels of Free and Esterified Oxylipins. Prostaglandins Other Lipid Mediat. 2014, 113–115, 21–29. 10.1016/j.prostaglandins.2014.05.002. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref397] Slatter D. A.; Aldrovandi M.; O’Connor A.; Allen S. M.; Brasher C. J.; Murphy R. C.; Mecklemann S.; Ravi S.; Darley-Usmar V.; O’Donnell V. B. Mapping the Human Platelet Lipidome Reveals Cytosolic Phospholipase A2 as a Regulator of Mitochondrial Bioenergetics during Activation. Cell Metab. 2016, 23 (5), 930–944. 10.1016/j.cmet.2016.04.001. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref398] Kolmert J.; Fauland A.; Fuchs D.; Säfholm J.; Gómez C.; Adner M.; Dahlén S.-E.; Wheelock C. E. Lipid Mediator Quantification in Isolated Human and Guinea Pig Airways: An Expanded Approach for Respiratory Research. Anal. Chem. 2018, 90 (17), 10239–10248. 10.1021/acs.analchem.8b01651. [DOI] [PubMed] [Google Scholar]

[ref399] Pedersen T. L.; Gray I. J.; Newman J. W. Plasma and Serum Oxylipin, Endocannabinoid, Bile Acid, Steroid, Fatty Acid and Nonsteroidal Anti-Inflammatory Drug Quantification in a 96-Well Plate Format. Anal. Chim. Acta 2021, 1143, 189–200. 10.1016/j.aca.2020.11.019. [DOI] [PubMed] [Google Scholar]

[ref400] Rajan M. R.; Sotak M.; Barrenäs F.; Shen T.; Borkowski K.; Ashton N. J.; Biörserud C.; Lindahl T. L.; Ramström S.; Schöll M.; Lindahl P.; Fiehn O.; Newman J. W.; Perkins R.; Wallenius V.; Lange S.; Börgeson E. Comparative Analysis of Obesity-Related Cardiometabolic and Renal Biomarkers in Human Plasma and Serum. Sci. Rep. 2019, 9 (1), 15385. 10.1038/s41598-019-51673-0. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref401] Matthew J. A.; Chan H. W. S.; Galliard T. A Simple Method for the Preparation of Pure 9-D-Hydroperoxide of Linoleic Acid and Methyl Linoleate Based on the Positional Specificity of Lipoxygenase in Tomato Fruit. Lipids 1977, 12 (3), 324–326. 10.1007/BF02533358. [DOI] [PubMed] [Google Scholar]

[ref402] Gardner H. W.Analysis of Plant Lipoxygenase Metabolites in Advances in Lipid Methodology - Four; W.W. Christie, Oily Press, Dundee, 1997. [Google Scholar]

[ref403] Iacazio G.; Langrand G.; Baratti J.; Buono G.; Triantaphylides C. Preparative, Enzymic Synthesis of Linoleic Acid (13S)-Hydroperoxide Using Soybean Lipoxygenase-1. J. Org. Chem. 1990, 55 (5), 1690–1691. 10.1021/jo00292a056. [DOI] [Google Scholar]

[ref404] Corey E. J.; Lansbury P. T. Jr Stereochemical Course of 5-Lipoxygenation of Arachidonate by Rat Basophil Leukemic Cell (RBL-1) and Potato Enzymes. J. Am. Chem. Soc. 1983, 105 (12), 4093–4094. 10.1021/ja00350a059. [DOI] [Google Scholar]

[ref405] Boeglin W. E.; Kim R. B.; Brash A. R. A 12R-Lipoxygenase in Human Skin: Mechanistic Evidence, Molecular Cloning, and Expression. Proc. Natl. Acad. Sci. U. S. A. 1998, 95 (12), 6744–6749. 10.1073/pnas.95.12.6744. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref406] Hamberg M. Fatty Acid Hydroperoxide Isomerase in Saprolegnia parasitica: Structural Studies of Epoxy Alcohols Formed from Isomeric Hydroperoxyoctadecadienoates. Lipids 1989, 24 (4), 249–255. 10.1007/BF02535158. [DOI] [Google Scholar]

[ref407] Peers K. E.; Coxon D. T. Controlled Synthesis of Monohydroperoxides by α-Tocopherol Inhibited Autoxidation of Polyunsaturated Lipids. Chem. Phys. Lipids 1983, 32 (1), 49–56. 10.1016/0009-3084(83)90069-5. [DOI] [Google Scholar]

[ref408] Hamberg M. Stereochemistry of Hydrogen Removal During Oxygenation of Linoleic Acid by Singlet Oxygen and Synthesis of 11(S)-Deuterium-Labeled Linoleic Acid. Lipids 2011, 46 (2), 201–206. 10.1007/s11745-010-3510-4. [DOI] [PubMed] [Google Scholar]

[ref409] Przybyla D.; Göbel C.; Imboden A.; Hamberg M.; Feussner I.; Apel K. Enzymatic, but Not Non-Enzymatic, ¹O₂-Mediated Peroxidation of Polyunsaturated Fatty Acids Forms Part of the EXECUTER1-Dependent Stress Response Program in the Flu Mutant of Arabidopsis thaliana. Plant J. 2008, 54 (2), 236–248. 10.1111/j.1365-313X.2008.03409.x. [DOI] [PubMed] [Google Scholar]

[ref410] Hamberg M. Steric Analysis of Hydroperoxides Formed by Lipoxygenase Oxygenation of Linoleic Acid. Anal. Biochem. 1971, 43 (2), 515–526. 10.1016/0003-2697(71)90282-X. [DOI] [PubMed] [Google Scholar]

[ref411] Kai K.; Takeuchi J.; Kataoka T.; Yokoyama M.; Watanabe N. Structure-Activity Relationship Study of Flowering-Inducer FN against Lemna paucicostata. Tetrahedron 2008, 64 (28), 6760–6769. 10.1016/j.tet.2008.04.115. [DOI] [Google Scholar]

[ref412] Bergström S.; Aulin-Erdtman G.; Rolander B.; Stenhagen E.; Östling S. The Monoketo- and Monohydroxyoctadecanoic Acids. Preparation and Characterization by Thermal and X-Ray Methods. Acta Chem. Scand. 1952, 6, 1157–1174. 10.3891/acta.chem.scand.06-1157. [DOI] [Google Scholar]

[ref413] Iacazio G. Easy Access to Various Natural Keto Polyunsaturated Fatty Acids and Their Corresponding Racemic Alcohols. Chem. Phys. Lipids 2003, 125 (2), 115–121. 10.1016/S0009-3084(03)00087-2. [DOI] [PubMed] [Google Scholar]

[ref414] Armstrong M. M.; Diaz G.; Kenyon V.; Holman T. R. Inhibitory and Mechanistic Investigations of Oxo-Lipids with Human Lipoxygenase Isozymes. Bioorg. Med. Chem. 2014, 22 (15), 4293–4297. 10.1016/j.bmc.2014.05.025. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref415] Maerker G.; Haeberer E. T.; Ault W. C. Epoxidation of Methyl Linoleate. I. The Question of Positional Selectivity in Monoepoxidation. J. Am. Oil Chem. Soc. 1966, 43 (2), 100–104. 10.1007/BF02641026. [DOI] [PubMed] [Google Scholar]

[ref416] Ferrari M.; Ghisalberti E. L.; Pagnoni U. M.; Pelizzoni F. Monoepoxidation of Methyl Linoleate: (±)-Methyl Vernolate and (±)-Methyl Coronarate. J. Am. Oil Chem. Soc. 1968, 45 (10), 649–651. 10.1007/BF02541248. [DOI] [Google Scholar]

[ref417] Newman J. W.; Watanabe T.; Hammock B. D. The Simultaneous Quantification of Cytochrome P450 Dependent Linoleate and Arachidonate Metabolites in Urine by HPLC-MS/MS. J. Lipid Res. 2002, 43 (9), 1563–1578. 10.1194/jlr.D200018-JLR200. [DOI] [PubMed] [Google Scholar]

[ref418] Teder T.; Boeglin W. E.; Brash A. R. Oxidation of C18 Hydroxy-Polyunsaturated Fatty Acids to Epoxide or Ketone by Catalase-Related Hemoproteins Activated with Iodosylbenzene. Lipids 2017, 52 (7), 587–597. 10.1007/s11745-017-4271-0. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref419] Municoy M.; González-Benjumea A.; Carro J.; Aranda C.; Linde D.; Renau-Mínguez C.; Ullrich R.; Hofrichter M.; Guallar V.; Gutiérrez A.; Martínez A. T. Fatty-Acid Oxygenation by Fungal Peroxygenases: From Computational Simulations to Preparative Regio- and Stereoselective Epoxidation. ACS Catal. 2020, 10 (22), 13584–13595. 10.1021/acscatal.0c03165. [DOI] [Google Scholar]

[ref420] Gao B.; Boeglin W. E.; Zheng Y.; Schneider C.; Brash A. R. Evidence for an Ionic Intermediate in the Transformation of Fatty Acid Hydroperoxide by a Catalase-Related Allene Oxide Synthase from the Cyanobacterium Acaryochloris marina. J. Biol. Chem. 2009, 284 (33), 22087–22098. 10.1074/jbc.M109.013151. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref421] SMITH C. R. Jr; BAGBY M. O.; LOHMAR R. L.; GLASS C. A.; WOLFF I. A. The Epoxy Acids of Chrysanthemum coronarium and Clarkia elegans Seed Oils. J. Org. Chem. 1960, 25 (2), 218–222. 10.1021/jo01072a019. [DOI] [Google Scholar]

[ref422] Gunstone F. D. Fatty Acids. Part II. The Nature of the Oxygenated Acid Present in Vernonia anthelmintica(Willd.) Seed Oil. J. Chem. Soc. Resumed 1954, (0), 1611–1616. 10.1039/jr9540001611. [DOI] [Google Scholar]

[ref423] Osbond J. M. 1043. Essential Fatty Acids. Part II. Synthesis of (±)-Vernolic, Linoleic, and γ-Linolenic Acid. J. Chem. Soc. Resumed 1961, 0 (0), 5270–5275. 10.1039/JR9610005270. [DOI] [Google Scholar]

[ref424] Morris L. J.; Wharry D. M. Naturally Occurring Epoxy Acids. IV. The Absolute Optical Configuration of Vernolic Acid. Lipids 1966, 1 (1), 41–46. 10.1007/BF02668123. [DOI] [PubMed] [Google Scholar]

[ref425] VanRheenen V.; Cha D. Y.; Hartley W. M.. Catalytic Osmium Tetroxide Oxidation of Olefins: cis-1,2-Cyclohexanediol. In Organic Syntheses; John Wiley & Sons, 2003; pp 43–43. 10.1002/0471264180.os058.08. [DOI] [Google Scholar]

[ref426] Corey E. J.; Su W.; Mehrotra M. M. An Efficient and Simple Method for the Conversion of 15-HPETE to 14,15-EPETE (Lipotriene A) and 5-HPETE to Leukotriene a as the Methyl Esters. Tetrahedron Lett. 1984, 25 (45), 5123–5126. 10.1016/S0040-4039(01)81540-9. [DOI] [Google Scholar]

[ref427] Hamberg M. Vanadium-Catalyzed Transformation of 13(S)-Hydroperoxy-9(Z), 11(E)-Octadecadienoic Acid: Structural Studies on Epoxy Alcohols and Trihydroxy Acids. Chem. Phys. Lipids 1987, 43 (1), 55–67. 10.1016/0009-3084(87)90017-X. [DOI] [Google Scholar]

[ref428] Hamberg M.; Herman C. A.; Herman R. P. Novel Biological Transformations of 15-Ls-Hydroperoxy-5,8,11,13-Eicosatetraenoic Acid. Biochim. Biophys. Acta BBA - Lipids Lipid Metab. 1986, 877 (3), 447–457. 10.1016/0005-2760(86)90211-0. [DOI] [PubMed] [Google Scholar]

[ref429] Hamberg M.; Olsson U. Efficient and Specific Conversion of 9-Lipoxygenase Hydroperoxides in the Beetroot. Formation of Pinellic Acid. Lipids 2011, 46 (9), 873–878. 10.1007/s11745-011-3592-7. [DOI] [PubMed] [Google Scholar]

[ref430] Thomas C. P.; Boeglin W. E.; Garcia-Diaz Y.; O’Donnell V. B.; Brash A. R. Steric Analysis of Epoxyalcohol and Trihydroxy Derivatives of 9-Hydroperoxy-Linoleic Acid from Hematin and Enzymatic Synthesis. Chem. Phys. Lipids 2013, 167–168, 21–32. 10.1016/j.chemphyslip.2013.01.002. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref431] Hamberg M. Regio- and Stereochemical Analysis of Trihydroxyoctadecenoic Acids Derived from Linoleic Acid 9- and 13-Hydroperoxides. Lipids 1991, 26 (6), 407–415. 10.1007/BF02536065. [DOI] [PubMed] [Google Scholar]

[ref432] Fuchs D.; Hamberg M.; Sköld C. M.; Wheelock Å. M.; Wheelock C. E. An LC-MS/MS Workflow to Characterize 16 Regio- and Stereoisomeric Trihydroxyoctadecenoic Acids. J. Lipid Res. 2018, 59 (10), 2025–2033. 10.1194/jlr.D087429. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref433] Hamberg M.; Hamberg G. Peroxygenase-Catalyzed Fatty Acid Epoxidation in Cereal Seeds (Sequential Oxidation of Linoleic Acid into 9(S),12(S),13(S)-Trihydroxy-10(E)-Octadecenoic Acid). Plant Physiol. 1996, 110 (3), 807–815. 10.1104/pp.110.3.807. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref434] Davis R. W.; Allweil A.; Tian J.; Brash A. R.; Sulikowski G. A. Stereocontrolled Synthesis of Four Isomeric Linoleate Triols of Relevance to Skin Barrier Formation and Function. Tetrahedron Lett. 2018, 59 (52), 4571–4573. 10.1016/j.tetlet.2018.11.033. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref435] Sabitha G.; Bhikshapathi M.; Reddy E. V.; Yadav J. S. Synthesis of (−)-Pinellic Acid and Its (9R,12S,13S)-Diastereoisomer. Helv. Chim. Acta 2009, 92 (10), 2052–2057. 10.1002/hlca.200900095. [DOI] [Google Scholar]

[ref436] Woodcock S. R.; Bonacci G.; Gelhaus S. L.; Schopfer F. J. Nitrated Fatty Acids: Synthesis and Measurement. Free Radic. Biol. Med. 2013, 59, 14–26. 10.1016/j.freeradbiomed.2012.11.015. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref437] Kaya K.; Miura T. Effects of Nitrogen Dioxide on Fatty Acid Compositions of Red Cell Membranes, Sera, and Livers in Rats. Environ. Res. 1982, 27 (1), 24–35. 10.1016/0013-9351(82)90054-8. [DOI] [PubMed] [Google Scholar]

[ref438] Kunimoto M.; Mochitate K.; Kaya K.; Miura T.; Kubota K. Effects of Nitrogen Dioxide on Red Blood Cells of Rats: Alterations of Cell Membrane Components and Populational Changes of Red Blood Cells during in Vivo Exposure to NO2. Environ. Res. 1984, 33 (2), 361–369. 10.1016/0013-9351(84)90034-3. [DOI] [PubMed] [Google Scholar]

[ref439] Kaya K.; Miura T.; Kubota K. Effects of Nitrogen Dioxide on Red Blood Cells of Rats: Changes in Components of Red Cell Membranes during in Vivo Exposure to NO2. Environ. Res. 1980, 23 (2), 397–409. 10.1016/0013-9351(80)90074-2. [DOI] [PubMed] [Google Scholar]

[ref440] O’Donnell V. B.; Eiserich J. P.; Chumley P. H.; Jablonsky M. J.; Krishna N. R.; Kirk M.; Barnes S.; Darley-Usmar V. M.; Freeman B. A. Nitration of Unsaturated Fatty Acids by Nitric Oxide-Derived Reactive Nitrogen Species Peroxynitrite, Nitrous Acid, Nitrogen Dioxide, and Nitronium Ion. Chem. Res. Toxicol. 1999, 12 (1), 83–92. 10.1021/tx980207u. [DOI] [PubMed] [Google Scholar]

[ref441] Gorczynski M. J.; Huang J.; King S. B. Regio- and Stereospecific Syntheses and Nitric Oxide Donor Properties of (E)-9- and (E)-10-Nitrooctadec-9-Enoic Acids. Org. Lett. 2006, 8 (11), 2305–2308. 10.1021/ol060548w. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref442] Woodcock S. R.; Marwitz A. J. V.; Bruno P.; Branchaud B. P. Synthesis of Nitrolipids. All Four Possible Diastereomers of Nitrooleic Acids: (E)- and (Z)-, 9- and 10-Nitro-Octadec-9-Enoic Acids. Org. Lett. 2006, 8 (18), 3931–3934. 10.1021/ol0613463. [DOI] [PubMed] [Google Scholar]

[ref443] Dunny E.; Evans P. Stereocontrolled Synthesis of the PPAR-γ Agonist 10-Nitrolinoleic Acid. J. Org. Chem. 2010, 75 (15), 5334–5336. 10.1021/jo1007493. [DOI] [PubMed] [Google Scholar]

[ref444] Zanoni G.; Valli M.; Bendjeddou L.; Porta A.; Bruno P.; Vidari G. Improved Synthesis of (E)-12-Nitrooctadec-12-Enoic Acid, a Potent PPARγ Activator. Development of a “Buffer-Free” Enzymatic Method for Hydrolysis of Methyl Esters. J. Org. Chem. 2010, 75 (23), 8311–8314. 10.1021/jo101806m. [DOI] [PubMed] [Google Scholar]

[ref445] Hassan M.; Krieg S.-C.; Ndefo Nde C.; Roos J.; Maier T. J.; El Rady E. A.; Raslan M. A.; Sadek K. U.; Manolikakes G. Streamlined One-Pot Synthesis of Nitro Fatty Acids. Eur. J. Org. Chem. 2021, 2021 (15), 2239–2252. 10.1002/ejoc.202100247. [DOI] [Google Scholar]

[ref446] Elix J. A.; Sargent M. V. Studies in Furan Chemistry. Part VI. The Synthesis of 8-(5-Hexyl-2-Furyl)Octanoic Acid, a Fatty Acid Found in Exocarpus Seed Oil. J. Chem. Soc. C Org. 1968, (0), 595–596. 10.1039/j39680000595. [DOI] [Google Scholar]

[ref447] Lie Ken Jie M. S. F.; Wong K. P. Synthesis of Dimethyl-Substituted C₁₈ Furanoid Fatty Ester. J. Am. Oil Chem. Soc. 1992, 69 (5), 485–487. 10.1007/BF02540955. [DOI] [Google Scholar]

[ref448] Lie Ken Jie M. S. F.; Lam C. H. Fatty Acids, Part XVI: The Synthesis of All Isomeric C18 Furan-Containing Fatty Acids. Chem. Phys. Lipids 1978, 21 (3), 275–287. 10.1016/0009-3084(78)90073-7. [DOI] [Google Scholar]

[ref449] Alaiz M.; Hidalgo F. J.; Zamora R.; Millán F.; Maza M. P.; Vioque E. Synthesis of 9,12-Epoxyoctadeca-9,11-Dienoic Acid. Chem. Phys. Lipids 1988, 48 (3), 289–292. 10.1016/0009-3084(88)90099-0. [DOI] [Google Scholar]

[ref450] El Fangour S.; Guy A.; Vidal J.-P.; Rossi J.-C.; Durand T. Total Synthesis of Phytoprostane F₁ and Its 16 Epimer. Tetrahedron Lett. 2003, 44 (10), 2105–2108. 10.1016/S0040-4039(03)00171-0. [DOI] [Google Scholar]

[ref451] El Fangour S.; Guy A.; Despres V.; Vidal J.-P.; Rossi J.-C.; Durand T. Total Synthesis of the Eight Diastereomers of the Syn-Anti-Syn Phytoprostanes F₁ Types I and II. J. Org. Chem. 2004, 69 (7), 2498–2503. 10.1021/jo035638i. [DOI] [PubMed] [Google Scholar]

[ref452] Roland A.; Durand T.; Egron D.; Vidal J.-P.; Rossi J.-C. Radical Cyclization of Highly Functionalized Precursors: Stereocontrol of Ring Closure of Acyclic 1-Substituted-2,4-Dihydroxylated Hex-5-Enyl Radicals. J. Chem. Soc. Perkin 1 2000, (2), 245–251. 10.1039/a904206g. [DOI] [Google Scholar]

[ref453] Durand T.; Guy A.; Henry O.; Vidal J.-P.; Rossi J.-C.; Rivalta C.; Valagussa A.; Chiabrando C. Total Syntheses of Four Metabolites of 15-F_2t-Isoprostane. Eur. J. Org. Chem. 2001, 2001 (4), 809–819. . [DOI] [Google Scholar]

[ref454] Smrček J.; Hájek M.; Hodek O.; Čížek K.; Pohl R.; Jahn E.; Galano J.; Oger C.; Durand T.; Cvačka J.; Jahn U. First Total Synthesis of Phytoprostanes with Prostaglandin-Like Configuration, Evidence for Their Formation in Edible Vegetable Oils and Orienting Study of Their Biological Activity. Chem. - Eur. J. 2021, 27 (37), 9556–9562. 10.1002/chem.202100872. [DOI] [PubMed] [Google Scholar]

[ref455] Rodrıguez A. R.; Spur B. W. First Total Synthesis of the E Type I Phytoprostanes. Tetrahedron Lett. 2003, 44 (40), 7411–7415. 10.1016/j.tetlet.2003.08.042. [DOI] [Google Scholar]

[ref456] Brinkmann Y.; Oger C.; Guy A.; Durand T.; Galano J.-M. Total Synthesis of 15-D2t- and 15-Epi-15-E2t-Isoprostanes. J. Org. Chem. 2010, 75 (7), 2411–2414. 10.1021/jo1000274. [DOI] [PubMed] [Google Scholar]

[ref457] Oger C.; Brinkmann Y.; Bouazzaoui S.; Durand T.; Galano J.-M. Stereocontrolled Access to Isoprostanes via a Bicyclo[3.3.0]Octene Framework. Org. Lett. 2008, 10 (21), 5087–5090. 10.1021/ol802104z. [DOI] [PubMed] [Google Scholar]

[ref458] Oger C.; Marton Z.; Brinkmann Y.; Bultel-Poncé V.; Durand T.; Graber M.; Galano J.-M. Lipase-Catalyzed Regioselective Monoacetylation of Unsymmetrical 1,5-Primary Diols. J. Org. Chem. 2010, 75 (6), 1892–1897. 10.1021/jo902541c. [DOI] [PubMed] [Google Scholar]

[ref459] Porta A.; Chiesa F.; Quaroni M.; Persico M.; Moratti R.; Zanoni G.; Vidari G. A Divergent Enantioselective Synthesis of 9-J1-Phytoprostane and 9-A1-Phytoprostane Methyl Ester. Eur. J. Org. Chem. 2014, 2014 (10), 2111–2119. 10.1002/ejoc.201301703. [DOI] [Google Scholar]

[ref460] Zanoni G.; Porta A.; Meriggi A.; Franzini M.; Vidari G. 1,2-Oxopalladation versus π-Allyl Palladium Route. A Regioconvergent Approach to a Key Intermediate for Cyclopentanoids Synthesis. New Insights into the Pd(II)-Catalyzed Lactonization Reaction. J. Org. Chem. 2002, 67 (17), 6064–6069. 10.1021/jo025722i. [DOI] [PubMed] [Google Scholar]

[ref461] Zanoni G.; Agnelli F.; Meriggi A.; Vidari G. Enantioselective Syntheses of Isoprostane and Iridoid Lactones Intermediates by Enzymatic Transesterification. Tetrahedron Asymmetry 2001, 12 (12), 1779–1784. 10.1016/S0957-4166(01)00311-1. [DOI] [Google Scholar]

[ref462] Schmidt A.; Boland W. General Strategy for the Synthesis of B1 Phytoprostanes, Dinor Isoprostanes, and Analogs. J. Org. Chem. 2007, 72 (5), 1699–1706. 10.1021/jo062359x. [DOI] [PubMed] [Google Scholar]

[ref463] Perlikowska W.; Mikołajczyk M. A Short Synthesis of Enantiomeric Phytoprostanes B₁ Type I. Synthesis 2009, 2009 (16), 2715–2718. 10.1055/s-0029-1216883. [DOI] [Google Scholar]

[ref464] Perlikowska W.; Mikołajczyk M. A Concise Approach to Both Enantiomers of Phytoprostane B₁ Type II. Tetrahedron Asymmetry 2011, 22 (18), 1767–1771. 10.1016/j.tetasy.2011.10.005. [DOI] [Google Scholar]

[ref465] Vázquez-Romero A.; Verdaguer X.; Riera A. General Approach to Prostanes B₁ by Intermolecular Pauson-Khand Reaction: Syntheses of Methyl Esters of Prostaglandin B₁ and Phytoprostanes 16-B₁-PhytoP and 9-L₁-PhytoP. Eur. J. Org. Chem. 2013, 2013 (9), 1716–1725. 10.1002/ejoc.201201442. [DOI] [Google Scholar]

[ref466] Vázquez-Romero A.; Cárdenas L.; Blasi E.; Verdaguer X.; Riera A. Synthesis of Prostaglandin and Phytoprostane B₁ Via Regioselective Intermolecular Pauson–Khand Reactions. Org. Lett. 2009, 11 (14), 3104–3107. 10.1021/ol901213d. [DOI] [PubMed] [Google Scholar]

[ref467] Guy A.; Flanagan S.; Durand T.; Oger C.; Galano J.-M. Facile Synthesis of Cyclopentenone B₁- and L₁-Type Phytoprostanes. Front. Chem. 2015, 3, 1–10. 10.3389/fchem.2015.00041. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref468] Beretta R.; Giambelli Gallotti M.; Pennè U.; Porta A.; Gil Romero J. F.; Zanoni G.; Vidari G. General Strategy for the Synthesis of B₁ and L₁ Prostanoids: Synthesis of Phytoprostanes (RS)-9-L₁-PhytoP, (R)-9-L₁-PhytoP, (RS)-16-B₁-PhytoP, and (RS)-16-L₁-PhytoP. J. Org. Chem. 2015, 80 (3), 1601–1609. 10.1021/jo502538b. [DOI] [PubMed] [Google Scholar]

[ref469] Cuyamendous C.; Leung K. S.; Durand T.; Lee J. C.-Y.; Oger C.; Galano J.-M. Synthesis and Discovery of Phytofurans: Metabolites of α-Linolenic Acid Peroxidation. Chem. Commun. 2015, 51 (86), 15696–15699. 10.1039/C5CC05736A. [DOI] [PubMed] [Google Scholar]

[ref470] Cuyamendous C.; Leung K. S.; Bultel-Poncé V.; Guy A.; Durand T.; Galano J.-M.; Lee J. C.-Y.; Oger C. Total Synthesis and in Vivo Quantitation of Phytofurans Derived from α-Linolenic Acid. Eur. J. Org. Chem. 2017, 2017 (17), 2486–2490. 10.1002/ejoc.201700270. [DOI] [Google Scholar]

[ref471] Kelly R. W. Physical Methods of Measurement of Prostaglandins. Clin. Endocrinol. Metab. 1973, 2 (3), 375–392. 10.1016/S0300-595X(73)80005-2. [DOI] [PubMed] [Google Scholar]

[ref472] Borgeat P.; Samuelsson B. Metabolism of Arachidonic Acid in Polymorphonuclear Leukocytes. Structural Analysis of Novel Hydroxylated Compounds. J. Biol. Chem. 1979, 254 (16), 7865–7869. 10.1016/S0021-9258(18)36026-5. [DOI] [PubMed] [Google Scholar]

[ref473] Ingram C. D.; Brash A. R. Characterization of HETEs and Related Conjugated Dienes by UV Spectroscopy. Lipids 1988, 23 (4), 340–344. 10.1007/BF02537345. [DOI] [PubMed] [Google Scholar]

[ref474] Jin J.; Boeglin W. E.; Brash A. R. Analysis of 12/15-Lipoxygenase Metabolism of EPA and DHA with Special Attention to Authentication of Docosatrienes. J. Lipid Res. 2021, 62, 100088. 10.1016/j.jlr.2021.100088. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref475] Ryhage R. The Mass Spectrometry Laboratory at the Karolinska Institute 1944–1987. Mass Spectrom. Rev. 1993, 12 (1), 1–49. 10.1002/mas.1280120102. [DOI] [Google Scholar]

[ref476] Tsikas D.; Zoerner A. A. Analysis of Eicosanoids by LC-MS/MS and GC-MS/MS: A Historical Retrospect and a Discussion. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2014, 964, 79–88. 10.1016/j.jchromb.2014.03.017. [DOI] [PubMed] [Google Scholar]

[ref477] Tsikas D. Application of Gas Chromatography-Mass Spectrometry and Gas Chromatography-Tandem Mass Spectrometry to Assess in Vivo Synthesis of Prostaglandins, Thromboxane, Leukotrienes, Isoprostanes and Related Compounds in Humans. J. Chromatogr. B. Biomed. Sci. Appl. 1998, 717 (1–2), 201–245. 10.1016/S0378-4347(98)00210-2. [DOI] [PubMed] [Google Scholar]

[ref478] Karara A.; Dishman E.; Blair I.; Falck J. R.; Capdevila J. H. Endogenous Epoxyeicosatrienoic Acids. Cytochrome P-450 Controlled Stereoselectivity of the Hepatic Arachidonic Acid Epoxygenase. J. Biol. Chem. 1989, 264 (33), 19822–19827. 10.1016/S0021-9258(19)47185-8. [DOI] [PubMed] [Google Scholar]

[ref479] Schulze B.; Lauchli R.; Sonwa M. M.; Schmidt A.; Boland W. Profiling of Structurally Labile Oxylipins in Plants by in Situ Derivatization with Pentafluorobenzyl Hydroxylamine. Anal. Biochem. 2006, 348 (2), 269–283. 10.1016/j.ab.2005.10.021. [DOI] [PubMed] [Google Scholar]

[ref480] Newman J. W.; Hammock B. D. Optimized Thiol Derivatizing Reagent for the Mass Spectral Analysis of Disubstituted Epoxy Fatty Acids. J. Chromatogr. A 2001, 925 (1–2), 223–240. 10.1016/S0021-9673(01)00998-0. [DOI] [PubMed] [Google Scholar]

[ref481] Murphy R. C.; Barkley R. M.; Zemski Berry K.; Hankin J.; Harrison K.; Johnson C.; Krank J.; McAnoy A.; Uhlson C.; Zarini S. Electrospray Ionization and Tandem Mass Spectrometry of Eicosanoids. Anal. Biochem. 2005, 346 (1), 1–42. 10.1016/j.ab.2005.04.042. [DOI] [PubMed] [Google Scholar]

[ref482] Mesaros C.; Lee S. H.; Blair I. A. Targeted Quantitative Analysis of Eicosanoid Lipids in Biological Samples Using Liquid Chromatography-Tandem Mass Spectrometry. J. Chromatogr. B Analyt. Technol. Biomed. Life. Sci. 2009, 877 (26), 2736–2745. 10.1016/j.jchromb.2009.03.011. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref483] Astarita G.; Kendall A. C.; Dennis E. A.; Nicolaou A. Targeted Lipidomic Strategies for Oxygenated Metabolites of Polyunsaturated Fatty Acids. Biochim. Biophys. Acta 2015, 1851 (4), 456–468. 10.1016/j.bbalip.2014.11.012. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref484] Chhonker Y. S.; Bala V.; Murry D. J. Quantification of Eicosanoids and Their Metabolites in Biological Matrices: A Review. Bioanalysis 2018, 10 (24), 2027–2046. 10.4155/bio-2018-0173. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref485] Kita Y.; Tokuoka S. M.; Shimizu T. Mediator Lipidomics by Liquid Chromatography-Tandem Mass Spectrometry. Biochim. Biophys. Acta Mol. Cell Biol. Lipids 2017, 1862 (8), 777–781. 10.1016/j.bbalip.2017.03.008. [DOI] [PubMed] [Google Scholar]

[ref486] Norris P. C.; Serhan C. N. Metabololipidomic Profiling of Functional Immunoresolvent Clusters and Eicosanoids in Mammalian Tissues. Biochem. Biophys. Res. Commun. 2018, 504 (3), 553–561. 10.1016/j.bbrc.2018.03.037. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref487] Gladine C.; Ostermann A. I.; Newman J. W.; Schebb N. H. MS-Based Targeted Metabolomics of Eicosanoids and Other Oxylipins: Analytical and Inter-Individual Variabilities. Free Radic. Biol. Med. 2019, 144, 72–89. 10.1016/j.freeradbiomed.2019.05.012. [DOI] [PubMed] [Google Scholar]

[ref488] O’Donnell V. B.; Milne G. L.; Nogueira M. S.; Giera M.; Helge Schebb N.. Quantitation of Oxylipins in Biological Samples, Focusing on Plasma, and Urine. In Mass Spectrometry for Lipidomics; John Wiley & Sons, 2023; pp 317–350. 10.1002/9783527836512.ch12. [DOI] [Google Scholar]

[ref489] Willenberg I.; Ostermann A. I.; Schebb N. H. Targeted Metabolomics of the Arachidonic Acid Cascade: Current State and Challenges of LC-MS Analysis of Oxylipins. Anal. Bioanal. Chem. 2015, 407 (10), 2675–2683. 10.1007/s00216-014-8369-4. [DOI] [PubMed] [Google Scholar]

[ref490] Giera M.; Galano J. M.. Eicosanoids. In Encyclopedia of Analytical Science, 3rd ed.; Worsfold P., Poole C., Townshend A., Miró M., Eds.; Academic Press: Oxford, 2019; pp 259–263. 10.1016/B978-0-12-409547-2.13984-8. [DOI] [Google Scholar]

[ref491] Ostermann A. I.; Willenberg I.; Schebb N. H. Comparison of Sample Preparation Methods for the Quantitative Analysis of Eicosanoids and Other Oxylipins in Plasma by Means of LC-MS/MS. Anal. Bioanal. Chem. 2015, 407 (5), 1403–1414. 10.1007/s00216-014-8377-4. [DOI] [PubMed] [Google Scholar]

[ref492] Yang W.; Schoeman J. C.; Di X.; Lamont L.; Harms A. C.; Hankemeier T. A Comprehensive UHPLC-MS/MS Method for Metabolomics Profiling of Signaling Lipids: Markers of Oxidative Stress, Immunity and Inflammation. Anal. Chim. Acta 2024, 1297, 342348. 10.1016/j.aca.2024.342348. [DOI] [PubMed] [Google Scholar]

[ref493] O’Donnell V. B.; Murphy R. C.; Watson S. P. Platelet Lipidomics: Modern Day Perspective on Lipid Discovery and Characterization in Platelets. Circ. Res. 2014, 114 (7), 1185–1203. 10.1161/CIRCRESAHA.114.301597. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref494] Aukema H. M.; Ravandi A. Factors Affecting Variability in Free Oxylipins in Mammalian Tissues. Curr. Opin. Clin. Nutr. Metab. Care 2022, 26 (2), 91–98. 10.1097/MCO.0000000000000892. [DOI] [PubMed] [Google Scholar]

[ref495] Deems R.; Buczynski M. W.; Bowers-Gentry R.; Harkewicz R.; Dennis E. A. Detection and Quantitation of Eicosanoids via High Performance Liquid Chromatography-Electrospray Ionization-Mass Spectrometry. Methods Enzymol. 2007, 432, 59–82. 10.1016/S0076-6879(07)32003-X. [DOI] [PubMed] [Google Scholar]

[ref496] Yang J.; Schmelzer K.; Georgi K.; Hammock B. D. Quantitative Profiling Method for Oxylipin Metabolome by Liquid Chromatography Electrospray Ionization Tandem Mass Spectrometry. Anal. Chem. 2009, 81 (19), 8085–8093. 10.1021/ac901282n. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref497] Yuan Z.-X.; Majchrzak-Hong S.; Keyes G. S.; Iadarola M. J.; Mannes A. J.; Ramsden C. E. Lipidomic Profiling of Targeted Oxylipins with Ultra-Performance Liquid Chromatography-Tandem Mass Spectrometry. Anal. Bioanal. Chem. 2018, 410 (23), 6009–6029. 10.1007/s00216-018-1222-4. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref498] Masoodi M.; Eiden M.; Koulman A.; Spaner D.; Volmer D. A. Comprehensive Lipidomics Analysis of Bioactive Lipids in Complex Regulatory Networks. Anal. Chem. 2010, 82 (19), 8176–8185. 10.1021/ac1015563. [DOI] [PubMed] [Google Scholar]

[ref499] Hinz C.; Liggi S.; Mocciaro G.; Jung S.; Induruwa I.; Pereira M.; Bryant C. E.; Meckelmann S. W.; O’Donnell V. B.; Farndale R. W.; Fjeldsted J.; Griffin J. L. A Comprehensive UHPLC Ion Mobility Quadrupole Time-of-Flight Method for Profiling and Quantification of Eicosanoids, Other Oxylipins, and Fatty Acids. Anal. Chem. 2019, 91 (13), 8025–8035. 10.1021/acs.analchem.8b04615. [DOI] [PMC free article] [PubMed] [Google Scholar]

[ref500] Ostermann A. I.; Koch E.; Rund K. M.; Kutzner L.; Mainka M.; Schebb N. H. Targeting Esterified Oxylipins by LC-MS - Effect of Sample Preparation on Oxylipin Pattern. Prostaglandins Other Lipid Mediat. 2020, 146, 106384. 10.1016/j.prostaglandins.2019.106384. [DOI] [PubMed] [Google Scholar]

[ref501] Koch E.; Mainka M.; Dalle C.; Ostermann A. I.; Rund K. M.; Kutzner L.; Froehlich L.-F.; Bertrand-Michel J.; Gladine C.; Schebb N. H. Stability of Oxylipins during Plasma Generation and Long-Term Storage. Talanta 2020, 217, 121074. 10.1016/j.talanta.2020.121074. [DOI] [PubMed] [Google Scholar]

PERMALINK

The Octadecanoids: Synthesis and Bioactivity of 18-Carbon Oxygenated Fatty Acids in Mammals, Bacteria, and Fungi

Johanna Revol-Cavalier

Alessandro Quaranta

John W Newman

Alan R Brash

Mats Hamberg

Craig E Wheelock

Abstract

1. Introduction

1.1. Fatty Acid Biosynthesis

Scheme 1. Formation of PUFAs from the Essential Fatty Acids Linoleic Acid (LA) and α-Linolenic Acid (ALA) via the Action of Desaturases and Elongases.

1.2. Oxylipins

1.3. Octadecanoids

Scheme 2. Examples of 18-Carbon Fatty Acids.

1.4. Scope

2. Proposed Octadecanoid Nomenclature

Scheme 3. Proposed Octadecanoid Nomenclature System.

Scheme 4. Nomenclature for α-Linolenic Acid (ALA)- and γ-Linolenic Acid (GLA)-Derived Octadecanoids.

Scheme 12. Epidermis-Type Lipoxygenase 3 (eLOX3) Catalytic Mechanism100.

Scheme 5. Nomenclature for Triol-Based Octadecanoids.

Scheme 6. Nomenclature for Octadecanoids Possessing Two Functional Groups.

3. Sources of Octadecanoids

3.1. Metabolism Overview

Scheme 7. Biosynthetic Pathways for the Oxidation of α-Linolenic Acid (ALA) in Plants.

3.2. Dietary Octadecanoids

3.2.1. Dietary Intake of C18-PUFAs

Scheme 8. Dietary Nitrate/Nitrite Can Combine with Conjugated Fatty Acids to Form Nitrated Fatty Acids (NO2-FAs) in the Acidic Environments of the Stomach.

3.2.2. Dietary Sources of Octadecanoids

Table 1. Median Concentration Values of Octadecanoids Quantified in Commercial Edible Plant Oils (μM)142.

4. Enzymatic Biosynthesis of Octadecanoids

4.1. Cyclooxygenases

4.1.1. Enzymatic Mechanism

Scheme 9. Cyclooxygenase (COX) Peroxidase Activity.

4.1.2. Cyclooxygenase-Derived Octadecanoids

Scheme 10. Linoleic Acid (LA) and α-Linolenic Acid (ALA)-Derived Octadecanoids Produced by Cyclooxygenases (COX).

4.2. Lipoxygenases

Table 2. Human Lipoxygenase (LOX) Tissue Expression and Substrate Preference31,173.

4.2.1. Enzymatic Mechanism

Scheme 11. Lipoxygenase (LOX) Catalytic Mechanism.

4.2.2. Lipoxygenase-Derived Octadecanoids

Scheme 13. Linoleic Acid (LA)-Derived Octadecanoids Produced by Lipoxygenases (LOX).

Scheme 14. α-Linolenic Acid (ALA)-Derived Octadecanoids Produced by Lipoxygenases (LOX).

Scheme 16. Sebaleic Acid-Derived Octadecanoids Produced by Lipoxygenases (LOX).

Scheme 15. Stearidonic Acid (SDA)-Derived Octadecanoids Produced by Lipoxygenases (LOX).

4.3. Cytochrome P450s

4.3.1. Enzymatic Mechanism

Scheme 17. Cytochrome P450 (CYP) Catalytic Mechanism31,196.

Scheme 18. Cytochrome P450 (CYP) Epoxidation Mechanism196.

4.3.2. Cytochrome P450-Derived Octadecanoids

Scheme 19. Linoleic Acid (LA)-Derived Octadecanoids Produced by Cytochrome P450 (CYP).

Scheme 20. α-Linolenic Acid (ALA)-Derived Octadecanoids Produced by Cytochrome P450 (CYP).

Scheme 21. γ-Linolenic Acid (GLA)- and Oleic Acid (OA)-Derived Octadecanoids Produced by Cytochrome P450 (CYP).

5. Microbial and Fungal Sources

5.1. Gut Microbiota Octadecanoids

Scheme 22. Linoleic Acid (LA)-Derived Octadecanoids Produced by the Gut Microbiota.

Scheme 23. α-Linolenic Acid (ALA)-Derived Octadecanoids Produced by the Gut Microbiota.

Scheme 24. γ-Linolenic Acid (GLA)-Derived Octadecanoids Produced by the Gut Microbiota.

5.2. Other Bacterial and Microbial Metabolites

Scheme 25. Linoleic Acid (LA)-Derived Octadecanoids Produced by Bacteria.

Scheme 26. α-Linolenic Acid (ALA)- and γ-Linolenic Acid (GLA)-Derived Octadecanoids Produced by Bacteria.

Scheme 27. Oleic Acid (OA)-Derived Octadecanoids Produced by Bacteria and Microbes.

5.3. Fungal Metabolites

Scheme 28. Linoleic Acid (LA)-Derived Octadecanoids Produced by Fungi Enzymes.

Scheme 29. Linoleic Acid (LA)-Derived Octadecanoids Produced by Fungi Enzymes.

Scheme 30. α-Linolenic Acid (ALA)-Derived Octadecanoids Produced by Fungal Enzymes.

Scheme 31. Linoleic Acid (LA)-Derived Octadecanoids Produced by Champignon (Agaricus bisporus or Psalliota bispora).

6. Nonenzymatic Biosynthesis of Octadecanoids

Scheme 49. Nonenzymatic Biosynthesis of trans-PhytoP and Hydroperoxide Octadecanoids from α-Linolenic Acid (ALA).

Scheme 32. Specific Octadecanoid Products of Linoleic Acid (LA) and α-Linolenic Acid (ALA) Photosensitized Oxidation.

6.1. Linear Metabolites

6.1.1. Alcohols

Scheme 33. Reduction of Hydroperoxide into Alcohol.

6.1.2. Ketones/Hydroxy-ketones

Scheme 34. Nonenzymatic Formation of Ketones from a Peroxyl Radical.

Scheme 35. Formation of a Ketone and a Hydroperoxide from a Peroxyl and an Alkoxyl Radical.

Scheme 36. Formation of a Ketone from an Alkoxyl Radical.

Scheme 37. Nonenzymatic Production of Octadecanoid Ketone and Hydroxy-Ketone Metabolites of Linoleic Acid (LA) and α-Linolenic Acid (ALA).

6.1.3. Diols

Scheme 38. Principal Mechanism for the Nonenzymatic Formation of Nonvicinal Diols.

Scheme 12. Epidermis-Type Lipoxygenase 3 (eLOX3) Catalytic Mechanism¹⁰⁰.

Scheme 8. Dietary Nitrate/Nitrite Can Combine with Conjugated Fatty Acids to Form Nitrated Fatty Acids (NO₂-FAs) in the Acidic Environments of the Stomach.

Table 1. Median Concentration Values of Octadecanoids Quantified in Commercial Edible Plant Oils (μM)¹⁴².

Table 2. Human Lipoxygenase (LOX) Tissue Expression and Substrate Preference^31,173.

Scheme 17. Cytochrome P450 (CYP) Catalytic Mechanism^31,196.

Scheme 18. Cytochrome P450 (CYP) Epoxidation Mechanism¹⁹⁶.

Scheme 52. Biosynthesis of PhytoFs Proposed by Jahn et al.¹⁰².

Table 3. Comparison of Octadecanoid Levels in Different Human Blood Preparations^a.

Scheme 74. Synthetic Pathway of 9-F₁t-PhytoP 55 and 16-F₁t-PhytoP 56 from Diacetone d-Glucose 53 by the Durand Group.

Scheme 75. Synthetic Pathway of 16-epi-16-E_1t-PhytoP 59 by Spur and Colleagues.

Scheme 76. Synthetic Pathway of ent-9-D_1t-PhytoP 62 by Durand and Colleagues.

Scheme 77. Synthetic Pathway for Formation of 9-J₁-PhytoP 65 and 9-A₁-PhytoP 66 by Zanoni and Vidari⁴⁵⁹.

Scheme 78. Synthetic Pathways for Formation of 9-L₁-PhytoP 69 and 16-B₁-PhytoP 72, and Their Enantiomers ent-69 and ent-72 by Boland and Colleagues⁴⁶².

Scheme 79. Synthetic Pathway for Formation of 16-L₁-PhytoP 79 by Zanoni and Vidari⁴⁶⁸.

Scheme 80. Synthetic Pathway of ent-16(R,S)-13-epi-ST-Δ¹⁴-9-PhytoF 83,⁴⁶⁹ent-16(R,S)-9-epi-ST-Δ¹⁴-10-PhytoF 84,⁴⁷⁰ and ent-9(R,S)-12-epi-ST-Δ¹⁰-13-PhytoF 85(470) by Durand and Colleagues.