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First published online July 14, 2006; 10.1104/pp.106.082115 Plant Physiology 142:28-39 (2006) © 2006 American Society of Plant Biologists Wounding Stimulates the Accumulation of Glycerolipids Containing Oxophytodienoic Acid and Dinor-Oxophytodienoic Acid in Arabidopsis Leaves1,[W]Division of Biology (C.M.B., P.T., A.A.S., E.J.B., S.M., M.R.R., S.W.E., J.S., R.W.) and Department of Biochemistry (J.Z.), Kansas State University, Manhattan, Kansas 66506; and University of Kansas Mass Spectrometry Laboratory, University of Kansas, Lawrence, Kansas 66045 (S.W.E., T.D.W.)
Although oxylipins can be synthesized from free fatty acids, recent evidence suggests that oxylipins are components of plastid-localized polar complex lipids in Arabidopsis (Arabidopsis thaliana). Using a combination of electrospray ionization (ESI) collisionally induced dissociation time-of-flight mass spectrometry (MS) to identify acyl chains, ESI triple-quadrupole (Q) MS in the precursor mode to identify the nominal masses of complex polar lipids containing each acyl chain, and ESI Q-time-of-flight MS to confirm the identifications of the complex polar lipid species, 17 species of oxylipin-containing phosphatidylglycerols, monogalactosyldiacylglycerols (MGDG), and digalactosyldiacylglycerols (DGDG) were identified. The oxylipins of these polar complex lipid species include oxophytodienoic acid (OPDA), dinor-OPDA (dnOPDA), 18-carbon ketol acids, and 16-carbon ketol acids. Using ESI triple-Q MS in the precursor mode, the accumulation of five OPDA- and/or dnOPDA-containing MGDG and two OPDA-containing DGDG species were monitored as a function of time in mechanically wounded leaves. In unwounded leaves, the levels of these oxylipin-containing complex lipid species were low, between 0.001 and 0.023 nmol/mg dry weight. However, within the first 15 min after wounding, the levels of OPDA-dnOPDA MGDG, OPDA-OPDA MGDG, and OPDA-OPDA DGDG, each containing two oxylipin chains, increased 200- to 1,000-fold. In contrast, levels of OPDA-hexadecatrienoic acid MGDG, linolenic acid (18:3)-dnOPDA MGDG, OPDA-18:3 MGDG, and OPDA-18:3 DGDG, each containing a single oxylipin chain, rose 2- to 9-fold. The rapid accumulation of high levels of galactolipid species containing OPDA-OPDA and OPDA-dnOPDA in wounded leaves is consistent with these lipids being the primary products of plastidic oxylipin biosynthesis.
Oxylipins are cyclic or acyclic oxidation products of fatty acids and are effectors in many biological pathways, including plant development and defense responses (Creelman and Mulpuri, 2002  ; Turner et al., 2002). Jasmonic acid (JA), oxophytodienoic acid (OPDA), and other cyclic oxylipins make up the jasmonate family (Creelman and Mulpuri, 2002; Turner et al., 2002; Weber, 2002). The role of JA in defense response was suggested in 1992 (Farmer and Ryan, 1992), when jasmonate synthesis was discovered to occur in response to wounding. More recently it has been demonstrated that OPDA is also capable of signaling and modulating gene expression (Kutchan, 1993; Blechert et al., 1995; Stintzi et al., 2001; Taki et al., 2005). Current understanding is that each plant or plant tissue produces its own oxylipin signature, or specific mixture of various oxylipins that elicit appropriate responses to particular environmental cues (Weber et al., 1997).
JA synthesis begins in the plastid and is completed in the peroxisome. The conversion of precursor linolenic acid (18:3) to OPDA (Fig. 1
) occurs in the plastid. The pathway involves formation of a hydroperoxide by a lipoxygenase, formation of an epoxide by allene oxide synthase, and formation of a cyclopentenone ring by allene oxide cyclase to produce OPDA. Alternatively, 12,13-epoxyoctadecatrienoic acid can be converted nonenzymatically to a ketol (Hamberg, 1988
In 2001, Stelmach and coworkers determined that 80% to 90% of the OPDA in unstimulated Arabidopsis (Arabidopsis thaliana) shoots is esterified in complex lipid species. They purified and identified a novel OPDA-containing galactolipid, 1-OPDA, 2-16:3 monogalactosyldiacylglycerol (MGDG). More recently, Hisamatsu and colleagues (2003 , 2005) have chemically characterized four additional OPDA-containing galactolipids isolated from Arabidopsis leaves. These compounds are 1-OPDA, 2-dnOPDA MGDG (annotated by these authors as Arabidopside A), OPDA-OPDA MGDG (Arabidopside B), 1-OPDA, 2-dnOPDA digalactosyldiacylglycerol (DGDG; Arabidopside C), and OPDA-OPDA DGDG (Arabidopside D). The relationship between esterified OPDA or dnOPDA and the free fatty acid forms of these compounds has not yet been elucidated. Figure 1 shows two possibilities for this relationship. One potential metabolic route includes the release of trienoic fatty acids (18:3 and 16:3) from galactolipids, followed by the free fatty acids being converted in three steps to OPDA and dnOPDA, which could then be incorporated into galactolipids. The second potential pathway is the conversion of 18:3 and 16:3 to OPDA and dnOPDA, while these acyl groups remain esterified to the galactolipids or phosphatidylglycerol (PG), followed by the release of OPDA and dnOPDA as free fatty acids.
In this work, we take advantage of recent advances in electrospray ionization (ESI) tandem mass spectrometry (MS/MS) methods to identify 13 additional OPDA-, dnOPDA-, and/or related ketol-containing MGDG, DGDG, and PG species, in addition to the one identified by Stelmach et al. (2001) The levels of seven OPDA- and dnOPDA-containing species are examined in Arabidopsis leaves, unstimulated, and during their response to wounding. We show that various oxylipin-containing species are synthesized and degraded at different rates. Identification of the additional oxylipin-containing species, development of a quantitative method for their analysis, and our kinetic observations lay the groundwork for further studies aimed at clarifying the biosynthetic pathway and role of complex lipids containing oxylipins in plants.
Detection of Oxylipins by CID-TOF Analysis
Fatty acyl species present in wounded and unwounded Arabidopsis leaves were identified with an ESI Q-TOF hybrid mass spectrometer. Subjecting a plant-derived lipid extract, ionized by ESI in negative ion mode, to CID in the collision cell of a Q-TOF mass spectrometer without selecting precursor lipid species (i.e. with radio frequency/direct current mass selection in the Q off) produces a spectrum that includes acyl anions of fatty acids derived from both free fatty acids and complex lipids (Figs. 2
and 3
). This experiment is accomplished by increasing the voltage offset on the collision to 35 V while acquiring data in MS1 mode (no precursor selection), thus activating all ions produced at once. We call this analysis CID-TOF here to distinguish it from traditional product ion analysis also performed with the Q-TOF mass spectrometer. Accurate mass data produced by the TOF analyzer can be correlated with chemical formulas to identify the acyl anions present (Table I
). CID-TOF analysis provides a reproducible and excellent semiquantitative view of acyl species present. It can only be considered semiquantitative, mainly because the propensity of an acyl species in a polar diacyl glycerolipid to form an anion depends on its position (Murphy, 1993
The spectra shown in Figure 2 show the total acyl species present from such an analysis before and 5 h after wounding of Arabidopsis leaves. Species that increase in amount after wounding include prominent anions with mass-to-charge ratio (m/z) 263.1656 and 291.1968 and smaller peaks at m/z 281.1733 and 309.2065. The best matches between these masses and chemical formulas for the two largest peaks are C16H23O3 and C18H27O3 (Table I). Whereas typical fatty acid anions contain two oxygen atoms, these anions contain three oxygen atoms, suggesting that these anions are derived from oxylipin species.
Table II
shows oxylipin species described in the literature with anion formulas corresponding to those detected. There are three described oxylipin anions with the formula C18H27O3. Catalytic hydrogenation, which removes only carbon-carbon double bonds, was used to identify the oxylipins present from among these oxylipins. After hydrogenation of the mixture, prominent peaks at nominal m/z 263 and 291 disappeared and new peaks appeared at m/z 267.1970 and 295.2275 (Fig. 3; Table I). As shown in Table II, of the described oxylipins with anion formulas of C18H27O3, only one, OPDA, gains only 4 atomic mass units (amu) upon hydrogenation. Similarly, hydrogenation of the peak at m/z 263 to 267 is consistent with identification of that peak as dnOPDA and not a 16-carbon analog of either ketotrienoic acid or colnelenic acid. The peak at m/z 309.2065 is consistent with a chemical formula of C18H29O4. This formula is consistent with the presence of either or both characterized 18-carbon ketols, 12-oxo-13-hydroxy-9(Z),15(Z)-octadecadienoic acid (12,13-
Complex Lipids Containing dnOPDA, OPDA, and Ketols Precursor scanning with an ESI triple-Q mass spectrometer was used to discover the complex lipid species that contain the oxylipin species found in lipid extracts of wounded Arabidopsis. Figure 4 shows peaks identified in the range m/z 740 to 830; peaks in the range m/z 740 to 1,050 are shown in Supplemental Figure 1. In Figure 4, sections A and B depict precursor ions of head group-derived fragments, while sections C to G depict precursor ions of specific acyl anions. Section A shows a scan specific for PG species. Section B depicts lipid species containing a fragment of m/z 235, which corresponds to the mass of a negative ion of glycero-Gal minus a water molecule. Thus, this section represents MGDG and DGDG species. Due to the presence of ammonium acetate (NH4OAc) in the infusion solvent, both [M – H]– and [M + OAc]– ions are formed, and each MGDG molecular species is represented by two peaks, although some of the MGDG [M + OAc]– peaks are at higher m/z than shown in Figure 4. Sections C and D depict the lipid species containing the normal acyl species 16:3 and 18:3, respectively, while sections E to G depict the lipid species containing dnOPDA, OPDA, and the 18-carbon ketol, respectively. It was not possible to use precursor scanning to elucidate the species containing a 16-carbon ketol because the 16-carbon ketol has the same nominal mass (m/z 281) as the normal chain acyl species oleic acid (18:1). Thus, a scan for precursors of m/z 281 would depict those lipid species containing either a 16-carbon ketol or 18:1. The difference in the scales of sections C and D (in the 108 range) as compared to sections E and F (106 range) and section G (105 range) indicates the greater prominence of the normal chain species (i.e. 16:3 and 18:3; sections C and D) relative to the oxylipin species (dnOPDA, OPDA, and ketols; sections E–G).
Sections E to G, in combination with the previous sections, provide tentative identification for each putative oxylipin molecular species by identifying the nominal masses of its acyl chains. The indicated species account for most, but not all, of the complex lipid species in this m/z range containing the investigated oxylipins. The peak identifications shown in Figure 4, as well as additional identifications indicated in Supplemental Figure 1, were confirmed by Q-TOF (accurate mass) analysis of product (acyl) ions in extract fractions. Before performing the Q-TOF analysis, the extract was fractionated by silicic acid chromatography to produce fraction 2, enriched in MGDG, and fraction 3, enriched in PG. DGDG was enriched in both fractions 2 and 3. Using fractionated material for the product ion analysis reduced the number of lipid species present, allowing more certain identification of acyl pairs. The discovery scans implied that the PG peaks at m/z 755 and 757 (A) contained OPDA (F), and their m/z values indicated that the acyl species paired with OPDA were palmitoleic (16:1) and palmitic (16:0) acids, respectively. Product ion analysis of these species in PG-containing fraction 3 indicated that both OPDA and 16:0 were indeed present in the species at m/z 757 and OPDA and 16:1 were present in the species at m/z 755 (Supplemental Table I). The latter species is depicted in Figure 5 . Similarly, each of the MGDG and DGDG species listed in Table III were identified by product ion mass analysis of the [M – H]– ions in fractions 2 and 3 using Q-TOF MS, and detailed results of these analyses are shown in Supplemental Table I.
Identifications were further substantiated by analysis of unfractionated and fractionated wounded Arabidopsis leaf extracts that were subjected to catalytic hydrogenation. Precursor scanning of a catalytically hydrogenated, unfractionated extract of wounded Arabidopsis (Supplemental Fig. 2) and Q-TOF product ion analysis of the catalytically hydrogenated and fractionated extract (Supplemental Table II) reveal the expected, hydrogenated product of each of the species identified in Table III and Supplemental Table I. These data are consistent with the notion that the oxylipin acyl species in the plastid-localized complex lipids include the same acyl species (OPDA, dnOPDA, and a ketol or ketols) that were found to be increased during wounding by CID-TOF MS scanning of wounded Arabidopsis leaf extracts.
Interestingly, two of the most prominent OPDA-containing MGDG species were the dioxylipin-containing species, OPDA-dnOPDA (Fig. 4, E and F) and OPDA-OPDA (F), while the largest DGDG species was OPDA-OPDA (Supplemental Fig. 1F, peaks o and o'). These are the species denoted by Hisamatsu et al. (2003
Precursor scanning was used to quantify the levels of the major OPDA- and dnOPDA-containing galactolipid molecular species in leaves without wounding and after mechanical wounding of the leaves. Stearoyl (18:0)-18:0 MGDG and 18:0-16:0 MGDG, prepared by catalytic hydrogenation of purified MGDG, served as internal standards for MGDG quantification, and 18:0-18:0 DGDG and 18:0-16:0 DGDG, similarly prepared, served as internal standards for DGDG quantification. In unwounded leaves, the level of each of the major OPDA- and dnOPDA-containing galactolipid molecular species was low, at less than 25 pmol per mg of leaf dry weight (Fig. 6 ). Upon wounding, the levels of these species, in particular the amounts of the species containing two oxylipins, rose sharply in the first 15 min (Fig. 7 ). OPDA-dnOPDA MGDG rose about 200-fold, OPDA-OPDA MGDG rose over 400-fold, and OPDA-OPDA DGDG rose over 1,000-fold in the first 15 min, while OPDA-16:3 MGDG, 18:3-dnOPDA MGDG, OPDA-18:3 MGDG, and OPDA-18:3 DGDG rose 9-, 3-, 2-, and 7-fold, respectively. Between 15 and 45 min after wounding, the levels of OPDA-dnOPDA MGDG and OPDA-OPDA MGDG increased another 1.2- to 1.4-fold each while the levels of OPDA-16:3 MGDG, 18:3-dnOPDA MGDG, OPDA-18:3 MGDG, and OPDA-18:3 DGDG each rose 2- to 5-fold. Between 45 min and 20 h after wounding, levels of all the OPDA- and dnOPDA-containing galactolipid species dropped 0% to 60% with the decrease tending to be more pronounced for the species with two oxylipin chains. Still, at 20 h after wounding, OPDA-dnOPDA MGDG was increased about 100-fold, OPDA-OPDA MGDG was increased 300-fold, and OPDA-OPDA DGDG was increased 600-fold over the level in nonwounded tissues, while OPDA-16:3 MGDG, 18:3-dnOPDA MGDG, OPDA-18:3 MGDG, and OPDA-18:3 DGDG were increased 21-, 5-, 4-, and 18-fold. Figure 8 shows the ratios of measured species containing two oxylipin chains to measured species containing one oxylipin chain at each time point. This view of the data highlights the rapid increase in dioxylipin species, as compared to monooxylipin species, immediately after mechanical wounding.
This work utilizes CID-TOF and traditional product ion spectra from a Q-TOF mass spectrometer, plus discovery scans on a triple-Q mass spectrometer, to identify and quantify numerous plastid complex lipid species containing oxylipins. In all, 18 (17, plus partially characterized Arabidopside C) molecular species of MGDG-, DGDG-, and PG-containing oxylipins were identified, including the five described previously (Stelmach et al., 2001 ; Hisamatsu et al., 2003, 2005). The major species detected were galactolipids containing OPDA and dnOPDA. In addition, ketol acids were found to be increased during wounding and were found to be present in MGDG and DGDG molecular species. This is a powerful combination of MS methods for identification and quantification of species of complex polar lipids.
Each of seven major OPDA- and dnOPDA-containing species was found to increase rapidly upon wounding. Species containing two OPDA or dnOPDA chains rose 200- to 1,400-fold, while species with a single OPDA or dnOPDA chain rose 6- to 30-fold. Compared to previous data on the levels of a single species, OPDA-16:3 MGDG, after wounding (Stelmach et al., 2001
The presence of the ketol acids in the galactolipids (Table I) suggests that a minor portion of the epoxyoctadecatrienoic acid formed during wounding is hydrolyzed nonenzymatically and that the nonenzymatic pathway contributes to the formation of esterified oxylipin species. Nonenzymatic reactions of epoxyoctadecatrienoic acid result in 85% to 90%
The discovery that MGDG and DGDG species containing two OPDA chains or an OPDA and a dnOPDA chain (as depicted in Fig. 5) are the major species formed in the initial response to wounding could imply that lipoxygenase acts directly on plastid-localized lipid species (Fig. 1, right-hand side), rather than on free fatty acids released from these lipids. Although the most abundant lipoxygenase in plastids, LIPOXYGENASE2, appears to be required for JA synthesis (Bell et al., 1995
The very rapid and very large increase in levels of the OPDA-dnOPDA and OPDA-OPDA species lend credence to the assertion that these species are created on the intact galactolipids rather than via the free fatty acid pathway (Fig. 1). OPDA-dnOPDA and OPDA-OPDA galactolipids rose 200- to 1,000-fold in the first 15 min after wounding, while free OPDA has been shown to rise from 3- to 15-fold in the first several hours after wounding (Weber et al., 1997 In a wounded leaf, galactolipid species containing OPDA represent several percent of the total galactolipids (typically totaling approximately 100 nmol/mg dry weight). If OPDA- and dnOPDA-containing polar complex lipid species are precursors of free OPDA and dnOPDA, then these species may serve as a reservoir for the production of free OPDA and dnOPDA at later times. It is also plausible that these lipids, with their folded-back acyl chains, might create a drastic alteration of plastid membrane structure that could affect the function of other membrane components.
A combination of MS methods has been used to discover acyl chains present in complex lipid mixtures, to discover the polar complex lipid species containing each of these chains, to verify the presence of particular complex lipid species, and to quantify the polar complex lipid species in unfractionated lipid extracts. The ability to quantify specific, oxylipin-containing, polar complex lipid species will facilitate our understanding of the physiological roles of these prominent plant lipids.
Plant Material Leaves from approximately 7-week-old Arabidopsis (Arabidopsis thaliana) ecotype Columbia plants were used for identification and kinetic analysis of the lipid species. Seeds were sown in Metro-Mix 360 soil (Scotts Company) and allowed to equilibrate in a 4°C chamber, covered with clear plastic for 72 h. The plants were transferred to a growth chamber at 22°C and 60% humidity with a 12/12 photoperiod at 210 µmol m–2 s–1. The plants were covered with clear plastic for 1 week, were left partially uncovered for 72 h, and then were uncovered for the remainder of the growth period. Two weeks after the seeds were sown, seedlings were thinned so that four or five plants remained, equidistant, in each 3-inch square pot. Miracle-Gro water soluble all purpose plant food (Scotts Company) was applied every 3 weeks, beginning at the time the seeds were sown, according to manufacturer's instructions.
Wounding was performed as described by Laudert and Weiler (1998)
Activated silicic acid (Unisil, Clarkson Chemical) was mixed with chloroform and packed into a column (1.5 cm diameter, 40 mL column volume). Crude extract containing lipid from 144 mg dry weight of Arabidopsis leaves in chloroform was allowed to bind to the column and was then eluted in five fractions: fraction 1, chloroform:acetone (1:1, v/v), 200 mL; fraction 2, acetone, 400 mL; fraction 3, chloroform:methanol (19:1, v/v), 400 mL; fraction 4, chloroform:methanol (4:1, v/v), 400 mL; and fraction 5, chloroform:methanol (1:1, v/v), 800 mL (Christie, 1982
Crude lipid extract was evaporated and dissolved in ethyl acetate/methanol (1:1, v/v) with platinum (IV) oxide at 1% by lipid weight. The suspension was subjected to >1 atm H2 at 25°C for 6 h. Hydrogenated lipid was centrifuged to remove platinum residue, and the supernatant was evaporated and dissolved in chloroform.
An automated ESI-MS/MS approach was used and data acquisition and analysis were performed as described previously (Welti et al., 2002 Samples were introduced using an autosampler (LC Mini PAL, CTC Analytics AG) fitted with the required injection loop for the acquisition time and presented to the ESI needle at 30 mL/min. The collision gas pressure was set at 4 (arbitrary units), the source temperature (heated nebulizer) was 100°C, the interface heater was on, –4.5 kV was applied to the electrospray capillary, the curtain gas was set at 20 (arbitrary units), and the two ion source gases at 45 (arbitrary units). The collision energy was –45 V with nitrogen in the collision cell, declustering potential was –100 V, entrance potential was –10 V, and exit potential was –20 V. The mass analyzers were adjusted to a resolution of 0.7 amu full width at half height. For each spectrum, 36 continuum scans were averaged in multiple channel analyzer mode. The background was subtracted, the data were smoothed, and peak areas integrated using a custom script and Applied Biosystems Analyst software.
The m/z and peak area data were sorted using Microsoft Excel to find peaks within each fatty acyl precursor spectrum corresponding to each target galactolipid molecular species. Peak areas were corrected for precursor ion isotopic distribution (Han and Gross, 2001
ESI-Q-TOF MS/MS and ESI-CID-TOF MS spectra were acquired with a Micromass Q-TOF-2 tandem mass spectrometer (Micromass). The TOF analyzer was tuned for maximum resolution (10,000 resolving power) with argon in the collision cell. Micromass MassLynx software was used as the operating software. All TOF spectra were acquired with daily mass calibration. For Q-TOF analysis, samples containing silicic acid-fractionated lipids were infused directly in chloroform:methanol:300 mM ammonium acetate in water (60:133:7, v/v) at 30 µL/min into the ESI source of the Q-TOF. Negatively charged precursor target peaks, selected by the Q tuned to transmit at 0.8 amu full width at half height, were subjected to product ion scanning in the negative ion mode. The collision energy was 35 V. For CID-TOF MS analysis, radio frequency/direct current mass selection in the Q is turned off, while the collision cell remains at 35 V offset. Unfractionated samples in chloroform:methanol:300 mM ammonium acetate in water (60:133:7, v/v) were infused at 20 µL/min into the ESI source. The unselected precursor ions were subjected to CID at 35 V in the negative mode. Both Q-TOF and CID-TOF mass spectra were mass corrected by locking the m/z to the determined theoretical m/z of the ion formed from a head group fragment with an exact mass of m/z 253.0923 in MGDG (in both Q-TOF and CID-TOF modes) and m/z 397.1346 (in Q-TOF mode) in DGDG. With the locked mass value set, the exact masses of product ions were calculated to ten thousandths of an amu. Chemical formulas for the product ions were determined using the Micromass MassLynx chemical formula tool. Each designated chemical formula match was the best match for a formula containing the indicated elements. Deviations between the detected m/z and theoretical m/z, calculated by the chemical formula tool, of the best-matched chemical formula were determined.
We would like to thank Dr. Giorgis Isaac for critical reading of the manuscript. We would also like to thank the reviewers for their helpful comments. Received April 13, 2006; accepted June 30, 2006.
1 This work was supported by grants from the National Science Foundation (grant nos. MCB 0455318 and DBI 0520140). Support of the Kansas Lipidomics Research Center was from the National Science Foundation's EPSCoR program (grant no. EPS–0236913) with matching support from the State of Kansas through Kansas Technology Enterprise Corporation and Kansas State University, as well as from Core Facility Support from the National Institutes of Health (grant no. P20 RR016475) from the INBRE program of the National Center for Research Resources. Accurate mass analysis was performed at the University of Kansas Mass Spectrometry Laboratory, using a Micromass Q-TOF2, which was purchased with funds from the University of Kansas, the University of Kansas Mass Spectrometry Laboratory, and Kansas National Science Foundation EPSCoR. This is contribution 06–180–J from the Kansas Agricultural Experiment Station. 
2 Present address: The University of Texas Southwestern Medical Center at Dallas, 5323 Harry Hines Boulevard, Dallas, TX 75390.
3 Present address: Children's Nutrition Research Center, Department of Pediatrics, Baylor College of Medicine, Houston, TX 77030. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Ruth Welti (welti@ksu.edu).
[W] The online version of this article contains Web-only data. www.plantphysiol.org/cgi/doi/10.1104/pp.106.082115 * Corresponding author; e-mail welti{at}ksu.edu; fax 785–532–6653.
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Proc Natl Acad Sci USA 94: 10473–10478 Welti R, Li W, Li M, Sang Y, Biesiada H, Zhou HE, Rajashekar CB, Williams TD, Wang X (2002) Profiling membrane lipids in plant stress responses: role of phospholipase D Welti R, Wang X, Williams TD (2003) Electrospray ionization tandem mass spectrometry scan modes for plant chloroplast lipids. Anal Biochem 314: 149–152[CrossRef][Web of Science][Medline] Ziegler J, Stenzel I, Hause B, Maucher H, Hamberg M, Grimm R, Ganal M, Wasternack C (2000) Molecular cloning of allene oxide cyclase: the enzyme establishing the stereochemistry of octadecanoids and jasmonates. J Biol Chem 275: 19132–19138 Zien CA, Wang C, Wang X, Welti R (2001) In vivo substrates and the contribution of the common phospholipase D, PLD This article has been cited by other articles:
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-oxidation result in shortening of the 18-carbon compound to 12-carbon JA. In 1997, Weber et al. identified dinor-OPDA (dnOPDA; [7S,11S]-10-oxy) as a 16-carbon analog of OPDA, formed from hexadecatrienoic acid (16:3) through the same pathway. 
-ketol) and 9-hydroxy-12-oxo-10(E),15(Z)-octadecadienoic acid (
-ketol), which can be formed nonenzymatically from allene oxide (Hamberg, 1988
