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The Journal of Biological Chemistry logoLink to The Journal of Biological Chemistry
. 2008 Jun 6;283(23):15539–15543. doi: 10.1074/jbc.R800001200

Routes to 4-Hydroxynonenal: Fundamental Issues in the Mechanisms of Lipid Peroxidation*,S⃞

Claus Schneider ‡, Ned A Porter §, Alan R Brash ‡,1
PMCID: PMC2414272  PMID: 18285327

Abstract

Although investigation of the toxicological and physiological actions of α/β-unsaturated 4-hydroxyalkenals has made great progress over the last 2 decades, understanding of the chemical mechanism of formation of 4-hydroxynonenal and related aldehydes has advanced much less. The aim of this review is to discuss mechanistic evidence for these non-enzymatic routes, especially of the underappreciated intermolecular pathways that involve dimerized and oligomerized fatty acid derivatives as key intermediates. These cross-molecular reactions of fatty acid peroxyls have also important implications for understanding of the basic initiation and propagation steps during lipid peroxidation and the nature of the products that arise.

4-Hydroxynonenal and Related Aldehydes

HNE2 was first identified as one of several α,β-unsaturated 4-hydroxyaldehydes formed during CCl4-induced lipid peroxidation in rat liver (1). The aldehydes detected varied in chain length from 8 to 11 carbons, but the 9-carbon HNE was the most prominent. HNE was recognized as having the greatest toxicological potential among these aldehyde products and over the years has become a paradigm for studying the pathophysiological effects of the cytotoxic products of lipid peroxidation (e.g. Refs. 2–5). HNE can also trigger signaling events in a physiological context (6, 7), and it is also commonly used as a marker of lipid peroxidation (e.g. Refs. 8 and 9).

The 4-hydroperoxy analog of HNE (4-HPNE) was recognized originally as an enzymatic product of ω6 fatty acid metabolism in plants (10), and more recently in lipid peroxidation, along with the 4-oxo analog, 4-oxononenal (4-oxo-2E-nonenal) (11–13). The 6-carbon analog 4-hydroxyhexenal derived from ω3 fatty acids has been identified as a prominent aldehyde in plant and mammalian tissues (8, 14). The short chain aldehydes have counterparts in the carboxyl-terminal end of the fatty acid molecule, and for esterified fatty acids, the aldehydes remain bound in phospholipids and other cellular esters or complex lipids. We cannot do justice to the many excellent reviews on the biological role of these products; one review appears as part of this minireview series (15).

Classical and Alternative Paradigms of Lipid Peroxidation

The fundamentals of autoxidation or lipid peroxidation were unraveled in the classical studies of Bolland (16), Bateman (17), and colleagues at the British Rubber Producers' Research Association. Their work established the free radical nature of the chain reaction and defined the three well recognized stages of the process as initiation, propagation, and termination (Fig. 1, upper). Notably, in this view of the overall process, the propensity for intermolecular cross-linking achieves a prominence mainly in the later stages as polymerization eventually ensues.

FIGURE 1.

FIGURE 1.

Comparison of classical and alternative paradigms for the initiation and propagation of lipid peroxidation. In the classical paradigm (upper), the most significant reactions involve single fatty acid molecules, until the final stages. In the alternative paradigm (lower), cross-chain peroxyl radical reactions play an important role during initiation and propagation. It is proposed that subsequent breakdown of these dimers and oligomers expands the pool of radicals that drive the autoxidation reaction and also yields aldehydic carbon chain cleavage products, e.g. HNE.

The lower half of Fig. 1 serves to introduce two themes that appear later in this review. In these perspectives of lipid peroxidation, intermolecular reactions of product conjugated diene hydroperoxides are implicated at earlier stages of the process, during initiation and propagation. We refer to these alternative paradigms here because they can have a major impact on understanding the main routes to HNE formation. In particular, there is experimental evidence for the formation of dimers in both early phases of autoxidation. These dimeric fatty acids are especially unstable and fragment to produce new radicals, hence their relevance to initiation, and they also appear to readily fragment to aldehydic cleavage products, hence their relevance to the sources of HNE.

Activation of the Oxygenated Fatty Acid

The cleavage of a fatty acid carbon chain is not a particularly facile chemical reaction, so some special activation is required to promote the ensuing formation of aldehydes. Identifying the basis of this activation has represented a main hurdle in understanding the overall mechanism. An essential part of the activation process involves reaction of the fatty acid with molecular oxygen, but the number of O2 molecules involved and exactly how they react has not been settled.

Enzymatic reactions could provide some clues to the mechanism. In particular, the plant fatty acid hydroperoxide lyase reaction offers a plausible hypothesis for a route to 4-HNE (Fig. 2). Plant 9-hydroperoxide lyase (cytochrome P450 CYP74C) cleaves the linoleic acid 9-hydroperoxide (9-HPODE) to two aldehydes, one being 3Z-nonenal. In observing the facile oxidation of 3Z-nonenal to 4-HPNE, we were prompted to propose a non-enzymatic pathway consisting of autoxidative formation of 9-HPODE from linoleic acid, followed by a Hock-type cleavage to give 3Z-nonenal, which undergoes rapid oxidation into 4-HPNE (12). It was shown that a route via 3Z-nonenal could account for HNE production in plant tissues (18). We surmised that the acid-catalyzed Hock cleavage could be achieved by the fatty acid carboxylate or other acidic moieties present under conditions of lipid peroxidation. (A short synopsis on the Hock rearrangement is given in supplemental Fig. 1.)

FIGURE 2.

FIGURE 2.

Enzymatic and non-enzymatic routes to 4-H(P)NE. Left, in the plant lipoxygenase (LOX)-hydroperoxide lyase (HPL) pathway, the 9-hydroperoxide of linoleic acid is cleaved to 3Z-nonenal via rearrangement into an unstable hemiacetal by CYP74C (47). Facile non-enzymatic oxygenation of 3Z-nonenal gives 4-HPNE. Right, autoxidation studies indicate that both the 9- and 13-hydroperoxides of linoleic acid are precursors of 4-HPNE, although the mechanisms remain controversial (12).

Studies with chiral hydroperoxides established the existence of at least two separate routes to 4-H(P)NE (12). Starting with 9S- and 13S-HPODE as model substrates for autoxidation reactions in a thin lipid film, analysis of the 4-hydroperoxy group showed that 4-HPNE derived from 9S-HPODE was racemic, as expected in the pathway through 3Z-nonenal. In contrast, 4-HPNE derived from 13S-HPODE largely retained the original S-configuration of the starting hydroperoxide. To account for this, we invoked formation of a 10,13-diHPODE intermediate, followed by Hock cleavage between C-9 and C-10 (12, 19). Two groups have synthesized these putative intermediate diHPODEs with a view to testing their intermediacy in HNE production (20, 21). Using these materials, we could show that the 10,13-diHPODE dihydroperoxide does not undergo cleavage to 4-HPNE under autoxidative conditions, thus largely disproving a mechanism involving simple dihydroperoxide intermediates, at least as a major factor under mild conditions (20). Thus, for HPNE formation from 9-HPODE, there is the possibility that carbon chain cleavage is achieved via a Hock rearrangement, whereas from 13-HPODE, an alternative mechanism must apply. The most plausible alternative mechanism, which can be applied equally to 9- or 13-HPODE, involves cross-molecular reactions of peroxyl radicals, as outlined below in this review.

The situation gets even more complicated because in addition to routes via radical-dependent hydroperoxide formation, there is also the possibility of peroxides generated via singlet oxidation of fatty acids. This is physiologically relevant in the retina, where all-trans-retinal can act as a photosensitizer (22). Gu et al. (22) investigated such a pathway and showed that model α-hydroperoxy endoperoxides are cleaved into aldehydes upon treatment with stoichiometric amounts of ferrous iron (Fe2+). The ferrous iron was proposed to reduce the hydroperoxide to an alkoxyl radical, which induces subsequent chain cleavage between the hydroperoxide- and endoperoxide-bearing carbons into two aldehydes. The question as to whether this mechanism of cleavage is also relevant for structurally different hydroperoxy endoperoxides that are formed during radical-dependent lipid peroxidation still needs to be investigated. In general, fatty acid hydroperoxy endoperoxides can readily arise from fatty acids containing at least three double bonds, but not readily from linoleic acid, for example (23, 24).

Also, a novel catalyst, vitamin C, was shown to produce a plethora of HNE-related molecules, including 4,5-epoxy-2E-decenal, 4-oxo-2-nonenal, 4-hydroperoxy-2-nonenal, and 4-hydroxy-2-nonenal, via a proposed pathway involving the hydroperoxide-derived alkoxyl radical from 13S-HPODE (25). In addition to these possibilities, other more hypothetical possibilities have been suggested for routes involving single fatty acid molecules, and at least part of these mechanistic schemes may still have a role in HNE production (26).

New Insights from Novel Products: Evidence for Intermolecular Cross-linking

A revision to our thinking on mechanism stemmed from the structural analysis of a prominent group of products with intermediate polarity formed during autoxidation of 15S-HETE and 15S-HPETE (27). We identified these as epoxyhydroxy and epoxyhydroperoxy derivatives, respectively, in which the original 15S-OH or 15S-OOH of the starting materials remained in place with unaltered chirality, and a new epoxide group replaced one of the double bonds of the conjugated diene system (Fig. 3). This is different in chemistry from synthesis of the group of epoxyhydroxy products that are often termed hepoxilins (28). Mechanistically, the epoxidation of the double bond could be accounted for by cross-molecular reaction of a peroxyl radical with a double bond (27).

FIGURE 3.

FIGURE 3.

Epoxidation of 15S-HETE or 15S-HPETE during thin film autoxidation. The starting hydro(pero)xy group at the ω6 carbon (red ovals) is unchanged in the products. Conversion of 15S-H(P)ETE to 4S-H(P)NE similarly involves retention of the originalω6 hydro(pero)xy group (19). These observations suggested a potential mechanistic connection between formation of the epoxides and carbon chain cleavage; cross-molecular peroxyl radical reactions are implicated (27).

A well documented example for the peroxyl radical-dependent dimerization and polymerization of unsaturated molecules is the copolymerization of styrene and oxygen (Fig. 4) (29, 30). The polymerization is driven by the addition of an initial styrene peroxyl radical to the styrene double bond forming the peroxide bridge between two styrene monomers and an alkyl radical that reacts with oxygen to re-form the peroxyl radical for the next cycle of addition to a double bond (Fig. 4A). “Unzipping” of the copolymer occurs when an alkyl radical reacts back with the peroxide bridge instead of adding a molecule of oxygen (Fig. 4B). In this so-called homolytic displacement reaction, the peroxide is split into two alkoxyl radicals, one of which forms an epoxide, whereas the other induces cleavage between the carbons bearing the alkoxyl radical and the peroxide group to form the two aldehydes benzaldehyde and formaldehyde in a chain reaction. Related chemistry is characterized with other starting materials (31, 32). The overall outcome of carbon chain cleavage of peroxide-linked polymers into aldehydes triggered further interest in the dimerization or oligomerization of fatty acid derivatives during autoxidation.

FIGURE 4.

FIGURE 4.

Styrene-oxygen copolymerization and depolymerization. A, the polymerization reaction; B, the depolymerization (unzipping) forms epoxide, benzaldehyde, and formaldehyde.

Fatty Acid Dimerization and Aldehyde Synthesis

Involvement in the Initiation of Lipid Peroxidation—Beginning with studies in the 1970s, Makio Morita and co-workers (33, 34) sought to identify the components in autoxidizing methyl linoleate that would initiate further peroxidation of lipids. Strikingly, they found that in the absence of added transition metals, fatty acid hydroperoxides showed little activity. Instead, the major active component extracted from partially autoxidized linoleate was identified as a class of peroxide distinct from hydroperoxides; these C-O-O-C-linked products, subsequently identified as fatty acid dimers, were potent inducers of lipid peroxidation. The titles of the articles give the gist of the findings: “Non-hydroperoxide Peroxides as Autocatalysts of Lipid Peroxidation” Monocarbonyls in the Autoxidation of Methyl Linoleate” (34). As indicated in the latter title, in the course of the work, the role of these peroxides in the production of aldehydes was recognized. This aldehyde production was studied with some elegant stable isotope techniques (35, 36). Although H(P)NE was not mentioned, other interesting aldehydes, including hexanal, 2-octenal, and 2,4-decadienal, important in food technologies, received mechanistic attention. The work culminated in identification of the active components as peroxide-linked dimers containing two hydroperoxy groups (37). Their overall conclusion is that radical generation and aldehyde production are two facets of the same process, sparked by the inherent instability of the dimeric peroxyhydroperoxides.

Involvement in Carbon Chain Cleavage to Give HNE-type Aldehydes—The literature on fatty acid cross-chain reactions includes other key studies, the significance of which has been largely unrecognized in the general scientific literature on lipid peroxidation and H(P)NE production. In a series of articles in the 1980s published in Agricultural and Biological Chemistry and Lipids, Miyashita et al. (38–42) studied the polymerization of methyl linoleate during mild conditions of lipid peroxidation, bubbling air through the pure lipid at 30 °C with no additives to accelerate reaction. They showed that as soon as monohydroperoxide products become detectable, peroxide-linked dimers are also present as significant products. Several of their articles described the formation and structural analysis of the dimeric products (39–41). Their studies ended with a report describing the products of decomposition of the dimeric fraction (under similar reaction conditions at 30 °C) as identified by gas chromatography-mass spectrometry (42). Monomeric products that retained the full 18 carbons of the original linoleate included monohydroxy-, epoxyhydroxy-, and trihydroxyoctadecenoates. Chain-shortened products included 8-oxooctanoate, 12-oxo-9-hydro(pero)xydodec-10-enoate, and 4-H(P)NE.

One example of the formation of dimeric peroxides and their involvement in aldehyde production is given in Fig. 5, in this case starting from 15-HPETE. There are parallels to the styrene polymerization/depolymerization scheme in that double bonds reacting with O2 produce multimers that decompose to yield two aldehydes and an epoxide equivalent to the ones we characterized in autoxidation mixtures (27).

FIGURE 5.

FIGURE 5.

Illustrative pathways to H(P)NE and other products via dimerization. Step I involves addition of a peroxyl radical to the 11,12-cis-double bond of 15-HPETE, yielding a carbon-centered allyl radical. If the cross-molecular peroxide bond is split at this stage (the side reaction at step II), the products are the 11,12-epoxy-15-hydroperoxide (Ep-OOH) and an alkoxyl radical. This epoxyhydroperoxide represents the type we have isolated and characterized (27). It is also equivalent to the reaction that leads a G526S mutant of COX-2 to arrest prostaglandin synthesis and instead form diepoxyalcohol products (48). Alternatively, proceeding from step II to step III, the allyl radical is oxygenated to a peroxyl, e.g. at C-12 as shown. This peroxyl can be reduced to a hydroperoxide by picking up a hydrogen, giving the dimer (step IV), or it can attack a third molecule of 15-HPETE at the 11,12-double bond (the side reaction at step III). The dimer formed by hydrogen abstraction is prone to breakage of one of the peroxide bonds, particularly the cross-molecular peroxide. This leads in turn to carbon chain cleavage and production of two aldehydes, 11-oxoundeca-5,8-dienoic acid and 4-HPNE. The C-11 oxo acid is highly prone to oxygenation, giving a second 4-hydroperoxyalkenal, 8-hydroperoxy-11-oxoundeca-5,9-dienoic acid (not shown).

Reactions of More Highly Unsaturated Substrates—In the late 1980s, Edwin Frankel and co-workers (43, 44) extended the investigation of dimerization to the more highly unsaturated linolenate (C18.3ω3). Similar to the Japanese groups (37, 39–42), they succeeded in isolating the dimeric fraction (and higher oligomers) by size exclusion chromatography in organic solvents (43, 44). One issue they encountered was the relative instability of the linolenate-derived dimers, such that further isolation of the individual species proved impractical.

Underexplored and Unexplained Chemistry

Herein we outline a key role for intermolecular cross-linking of peroxyl radicals in fatty acid chain cleavage. Even in the early stages of lipid peroxidation, this appears to be key to the production of reactive free products such as H(P)NE as well as bioactive phospholipid molecules with chain-shortened or oxidized substituents. There is one caveat in stating “in the early stages” of autoxidation. Some primary product formation is required, generating conjugated dienes, before dimerization can become significant. This follows because peroxyl radical addition to a conjugated diene is favored over reaction with a non-conjugated double bond. Once a pool of conjugated dienes is generated, peroxyl radical-initiated cross-linking can better compete with hydrogen abstraction from the unsaturated fatty acid bisallylic methylene. (We note in passing that in the case of conjugated linoleic acid, the chemistry will allow dimerization by reaction of a peroxyl with the primary reactant.) In relation to the competition aspect of peroxyl addition to conjugated diene versus hydrogen abstraction, Bolland (16) noted that “the absence of α-methylenic groups in styrene no doubt contributes to the predominance of the polymeric type of reaction.” In other words, with styrene, there are no α-methylenic hydrogens to compete with radical addition and oligomerization.

Does this dimerization scheme for generation of aldehydic products lead to the cross-linking of phospholipid molecules in membranes and formation of short-lived phospholipid dimers? This remains to be carefully examined. In relation to potential cross-reactions in membranes, we note evidence of another type of molecular cross-reaction shown by Mead and co-workers (45, 46) in the 1970s. They showed that in contrast to bulk-phase autoxidation of linoleate, in which hydroperoxides are the main products, when the fatty acids were absorbed as a unimolecular layer on silica (designed to mimic a biological membrane), the main products were simple linoleic epoxides (9,10-epoxyoctadecenoic and 12,13-epoxyoctadecenoic acids) (45). The same experiment was not conducted with phospholipid ester bilayers. It was demonstrated, however, that this type of epoxide product accumulates in the lungs of rats exposed to traces of the oxidizing gas NO2 (46). The relevance of this reaction to phospholipid oxidation as well as the potential dimerization issues remains to be investigated. Notably, dimerization is comparatively well characterized with methyl linoleate, but there are plenty of opportunities for unraveling this chemistry with more unsaturated fatty acids and their biological esters.

Supplementary Material

Supplemental Data
*

This work was supported, in whole or in part, by National Institutes of Health Grants GM15431 and GM076592. This is the sixth article of seven in the Oxidized Lipids Minireview Series. This minireview will be reprinted in the 2008 Minireview Compendium, which will be available in January, 2009.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. 1 and additional references.

Footnotes

2

The abbreviations used are: HNE, 4-hydroxynonenal (4-hydroxy-2E-nonenal); 4-HPNE, 4-hydroperoxynonenal (4-hydroperoxy-2E-nonenal); HPODE, hydroperoxyoctadecadienoic acid; H(P)ETE, hydro(pero)xyeicosatetraenoic acid.

References

  • 1.Benedetti, A., Comporti, M., and Esterbauer, H. (1980) Biochim. Biophys. Acta 620 281–296 [DOI] [PubMed] [Google Scholar]
  • 2.Liu, Q., Raina, A. K., Smith, M. A., Sayre, L. M., and Perry, G. (2003) Mol. Aspects Med. 24 305–313 [DOI] [PubMed] [Google Scholar]
  • 3.Petersen, D. R., and Doorn, J. A. (2004) Free Radic. Biol. Med. 37 937–945 [DOI] [PubMed] [Google Scholar]
  • 4.Leonarduzzi, G., Chiarpotto, E., Biasi, F., and Poli, G. (2005) Mol. Nutr. Food Res. 49 1044–1049 [DOI] [PubMed] [Google Scholar]
  • 5.West, J. D., and Marnett, L. J. (2006) Chem. Res. Toxicol. 19 173–194 [DOI] [PubMed] [Google Scholar]
  • 6.Nakashima, I., Liu, W., Akhand, A. A., Takeda, K., Kawamoto, Y., Kato, M., and Suzuki, H. (2003) Mol. Aspects Med. 24 231–238 [DOI] [PubMed] [Google Scholar]
  • 7.Dwivedi, S., Sharma, A., Patrick, B., Sharma, R., and Awasthi, Y. C. (2007) Redox Rep. 12 4–10 [DOI] [PubMed] [Google Scholar]
  • 8.Guichardant, M., Bacot, S., Moliere, P., and Lagarde, M. (2006) Prostaglandins Leukot. Essent. Fatty Acids 75 179–182 [DOI] [PubMed] [Google Scholar]
  • 9.Honzatko, A., Brichac, J., and Picklo, M. J. (2007) J. Chromatogr. B Anal. Technol. Biomed. Life Sci. 857 115–122 [DOI] [PubMed] [Google Scholar]
  • 10.Gardner, H. W., and Hamberg, M. (1993) J. Biol. Chem. 268 6971–6977 [PubMed] [Google Scholar]
  • 11.Lee, S. H., and Blair, I. A. (2000) Chem. Res. Toxicol. 13 698–702 [DOI] [PubMed] [Google Scholar]
  • 12.Schneider, C., Tallman, K. A., Porter, N. A., and Brash, A. R. (2001) J. Biol. Chem. 276 20831–20838 [DOI] [PubMed] [Google Scholar]
  • 13.Sayre, L. M., Lin, D., Yuan, Q., Zhu, X., and Tang, X. (2006) Drug Metab. Rev. 38 651–675 [DOI] [PubMed] [Google Scholar]
  • 14.Weichert, H., Kolbe, A., Wasternack, C., and Feussner, I. (2000) Biochem. Soc. Trans. 28 850–851 [PubMed] [Google Scholar]
  • 15.Hazen, S. L. (2008) J. Biol. Chem. 283 15527–15531 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Bolland, J. L. (1949) Quart. Rev. 3 1–21 [Google Scholar]
  • 17.Bateman, L. (1954) Quart. Rev. 8 147–167 [Google Scholar]
  • 18.Noordermeer, M. A., Feussner, I., Kolbe, A., Veldink, G. A., and Vliegenthart, J. F. G. (2000) Biochem. Biophys. Res. Commun. 277 112–116 [DOI] [PubMed] [Google Scholar]
  • 19.Schneider, C., Porter, N. A., and Brash, A. R. (2004) Chem. Res. Toxicol. 17 937–941 [DOI] [PubMed] [Google Scholar]
  • 20.Schneider, C., Boeglin, W. E., Yin, H., Stec, D. F., Hachey, D. L., Porter, N. A., and Brash, A. R. (2005) Lipids 40 1155–1162 [DOI] [PubMed] [Google Scholar]
  • 21.Zhang, W., Sun, M., and Salomon, R. G. (2006) J. Org. Chem. 71 5607–5615 [DOI] [PubMed] [Google Scholar]
  • 22.Gu, X., Zhang, W., and Salomon, R. G. (2007) J. Am. Chem. Soc. 129 6088–6089 [DOI] [PubMed] [Google Scholar]
  • 23.Frankel, E. N. (1987) Chem. Phys. Lipids 44 73–85 [DOI] [PubMed] [Google Scholar]
  • 24.Yin, H., Morrow, J. D., and Porter, N. A. (2004) J. Biol. Chem. 279 3766–3776 [DOI] [PubMed] [Google Scholar]
  • 25.Lee, S. H., Oe, T., and Blair, I. A. (2001) Science 292 2083–2086 [DOI] [PubMed] [Google Scholar]
  • 26.Porter, N. A., and Pryor, W. A. (1990) Free Radic. Biol. Med. 8 541–543 [DOI] [PubMed] [Google Scholar]
  • 27.Schneider, C., Boeglin, W. E., Yin, H., Porter, N. A., and Brash, A. R. (2008) Chem. Res. Toxicol., in press [DOI] [PubMed]
  • 28.Brash, A. R., Yu, Z., Boeglin, W. E., and Schneider, C. (2007) FEBS J. 274 3494–3502 [DOI] [PubMed] [Google Scholar]
  • 29.Mayo, F. R., and Miller, A. A. (1956) J. Am. Chem. Soc. 78 1023–1034 [Google Scholar]
  • 30.Mayo, F. R. (1958) J. Am. Chem. Soc. 80 2465–2480 [Google Scholar]
  • 31.Sugimoto, Y., Taketani, S., Kitamura, T., Uda, D., and Matsumoto, A. (2006) Macromolecules 39 9112–9119 [Google Scholar]
  • 32.Matsumoto, A., and Taketani, S. (2006) J. Am. Chem. Soc. 128 4566–4567 [DOI] [PubMed] [Google Scholar]
  • 33.Morita, M., and Fujimaki, M. (1973) Agric. Biol. Chem. 37 1213–1214 [Google Scholar]
  • 34.Morita, M., and Fujimaki, M. (1973) J. Agric. Food Chem. 21 860–863 [DOI] [PubMed] [Google Scholar]
  • 35.Morita, M., and Tokita, M. (1990) Chem. Phys. Lipids 56 209–215 [Google Scholar]
  • 36.Morita, M., and Tokita, M. (1993) Chem. Phys. Lipids 66 13–22 [Google Scholar]
  • 37.Morita, M., and Tokita, M. (2006) Lipids 41 91–95 [DOI] [PubMed] [Google Scholar]
  • 38.Miyashita, K., Fujimoto, K., and Kaneda, T. (1982) Agric. Biol. Chem. 46 751–755 [Google Scholar]
  • 39.Miyashita, K., Fujimoto, K., and Kaneda, T. (1982) Agric. Biol. Chem. 46 2293–2297 [Google Scholar]
  • 40.Miyashita, K., Fujimoto, K., and Kaneda, T. (1984) Agric. Biol. Chem. 48 2511–2515 [Google Scholar]
  • 41.Miyashita, K., Hara, N., Fujimoto, K., and Kaneda, T. (1985) Lipids 20 578–587 [Google Scholar]
  • 42.Miyashita, K., Hara, N., Fujimoto, K., and Kaneda, T. (1985) Agric. Biol. Chem. 49 2633–2640 [Google Scholar]
  • 43.Neff, W. E., Frankel, E. N., and Fujimoto, K. (1988) J. Am. Oil Chem. Soc. 65 616–623 [Google Scholar]
  • 44.Frankel, E. N., Neff, W. E., Selke, E., and Brooks, D. D. (1988) Lipids 23 295–298 [DOI] [PubMed] [Google Scholar]
  • 45.Wu, G. S., Stein, R. A., and Mead, J. F. (1977) Lipids 12 971–978 [DOI] [PubMed] [Google Scholar]
  • 46.Sevanian, A., Mead, J. F., and Stein, R. A. (1979) Lipids 14 634–643 [DOI] [PubMed] [Google Scholar]
  • 47.Grechkin, A. N., Bruhlmann, F., Mukhtarova, L. S., Gogolev, Y. V., and Hamberg, M. (2006) Biochim. Biophys. Acta 1761 1419–1428 [DOI] [PubMed] [Google Scholar]
  • 48.Schneider, C., Boeglin, W. E., and Brash, A. R. (2004) J. Biol. Chem. 279 4404–4414 [DOI] [PubMed] [Google Scholar]

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