Analysis of linoleic acid hydroperoxides generated by biomimetic and enzymatic systems through an integrated methodology

Analysis of linoleic acid hydroperoxides generated by biomimetic and enzymatic systems through an integrated methodology

Industrial Crops and Products 34 (2011) 1474–1481 Contents lists available at ScienceDirect Industrial Crops and Products journal homepage: www.else...

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Industrial Crops and Products 34 (2011) 1474–1481

Contents lists available at ScienceDirect

Industrial Crops and Products journal homepage:

Analysis of linoleic acid hydroperoxides generated by biomimetic and enzymatic systems through an integrated methodology Juan José Villaverde a,∗ , Sónia A.O. Santos a , Mário M.Q. Simões b , Carlos Pascoal Neto a , M. Rosário M. Domingues b , Armando J.D. Silvestre a a b

CICECO and Department of Chemistry, University of Aveiro, 3810-193 Aveiro, Portugal QOPNA and Department of Chemistry, University of Aveiro, 3810-193 Aveiro, Portugal

a r t i c l e

i n f o

Article history: Received 20 January 2011 Received in revised form 27 April 2011 Accepted 4 May 2011 Available online 8 June 2011 Keywords: Oxidation Linoleic acid Linoleic acid hydroperoxides Isomers HPLC–UV HPLC–MS/MS

a b s t r a c t An integrated methodology for the identification and quantification of linoleic acid hydroperoxides (HPODEs) mixtures, obtained by the Fenton’s reaction and by enzymatic oxidation, is reported. Unambiguous identification of the HPODEs formed (13-hydroperoxy-(9Z,11E)-octadecadienoic acid, 13hydroperoxy-(9E,11E)-octadecadienoic acid and 9-hydroperoxyoctadecadienoic acid) was performed by HPLC–MS/MS analysis, while quantification was carried out by HPLC–UV using an external calibration, based on the collection of chromatographic peaks, and measuring of total hydroperoxides by ferrous oxidation-xylenol orange (FOX) method. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Nowadays, the importance of biomass as a source of chemicals, materials, biofuels and energy within the biorefinery concept (Fernando et al., 2006; Kamm et al., 2006) is widely accepted. Although, due to their abundance, carbohydrates are considered as the main raw materials in this scenario (Corma et al., 2007), the integrated exploitation of all biomass fractions should be considered, and, at the current development stage, biorefinery will be in many cases mainly focused on the valorization of by-products from well established agro-forest industries (Fernando et al., 2006; Kamm et al., 2006). Vegetable oils have always been explored as raw materials for the industrial production of biobased chemicals, materials and fuels (Belgacem and Gandini, 2008; Corma et al., 2007; Meier et al., 2007) and, have also attracted an increased interest in this last few years (e.g. the previous references). The most common fatty acids components of vegetable oils have 16 and 18 carbon chains which can be totally aliphatic or bear one to several mid chain unsaturations. In fact, other carbon chain functionalities are only abundantly found in castor oil, from Ricinus communis L., where ricinoleic acid (12-hydroxy-9-cis-octadecenoic

∗ Corresponding author. Tel.: +351 234 370 711; fax: +351 234 370 084. E-mail address: [email protected] (J.J. Villaverde). 0926-6690/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.indcrop.2011.05.001

acid) is the predominant component (Mutlu and Meier, 2010). Additionally, unsaturated or mid-chain epoxidized ␻-hydroxyfatty acids can also be found, among the major components of suberin and cutin type biopolyesters (Gandini et al., 2006). These structural features imply that most vegetable oils/fatty transformation pathways involve either the carboxylic group or the mid-chain double bonds (Corma et al., 2007), and that the development of effective routes for the efficient functionalization of saturated or unsaturated fatty acid chains would be an important contribution to the valorization of this family of biobased raw materials. Tall oil is a by-product of the pulping industry, rich in unsaturated fatty acids (mainly linoleic acid), a commodity (Jen and Mcsweeney, 1985; Magee and Zinkel, 1992) that lost economical relevance in the last few decades. In this perspective, the development of new pathways for the conversion of linoleic acid (or other unsaturated acids) into more valuable components would be an important contribution to the valorization of this by-product and, inherently, to the implementation of the biorefinery concept. Conversion of unsaturated fatty acids into the corresponding hydroperoxides could be an interesting route to meet that goal. In fact, fatty acids hydroperoxides are already key intermediates in the oxypolymerization of fatty acids involved in the crosslinking of drying oils in paint formulations (Gorkum and Bouwman, 2005; Lazzari and Chiantore, 1999). Fatty acids hydroperoxides can

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Fe2+ + H2O2 Fe3+ + OH- + HO· Lipid-H + HO· Lipid · + H2O Lipid· + O2 Lipid-OO· Lipid-OO· + Lipid-H Lipid-OOH + Lipid·

also be versatile intermediates for the preparation of mid-chain hydroxyfatty acids (Gardner, 1989; Kharasch et al., 1950), epoxides (Frankel, 2005; Hamberg and Gotthammar, 1973), alcohols (Frankel, 2005), carbonyls (Frankel, 2005) and diacids (Gardner, 1989; Kharasch et al., 1950). Within a project aiming at discovering new lipoxygenases able to efficiently convert unsaturated fatty acid into the corresponding hydroperoxides, in the present paper we report the development of an integrated methodology for the detection, liquid chromatography separation, identification and quantification of hydroperoxyoctadecadienoic acids (HPODEs). The methodology was optimized using HPODE formed during oxidation of linoleic acid (LA) with Fenton’s system (Fe2+ /H2 O2 ) and quantification of total hydroperoxides was achieved with the ferrous oxidation-xylenol orange (FOX) method (Bou et al., 2008; Morgan et al., 2008), an efficient technique for the direct quantification of hydroperoxides, based on the ability of hydroperoxides to convert ferrous ions into ferric ions which subsequently form a complex with xylenol orange (XO) that is determined through spectrophotometry, with an absorption maximum at 560 nm (Bou et al., 2008; Morgan et al., 2008). The HPODE isomers were then unambiguously identified by high-performance liquid chromatography tandem mass spectrometry (HPLC–MS/MS), and detailed fragmentation pathways observed in the HPLC–MS/MS are discussed. The combination of high-performance liquid chromatography with UV detection (HPLC–UV), with the FOX method, allows carrying out calibrations for the quantification of each HPODE isomer. Finally, the applicability of the analytical methodology to enzymatic systems was validated with a commercial enzyme, what confirms the relevance of the method. 2. Material and methods 2.1. Chemicals Linoleic acid (≥99% purity), ammonium iron (II) sulfate hexahydrate (ReagentPlus® , ≥98%), iron (II) chloride tetrahydrate (ReagentPlus® , 99%) and 2,6-di-tert-butyl-4-methylphenol (BHT) (≥99% purity) were supplied by Sigma Chemicals Co. (Madrid, Spain). Xylenol Orange disodium salt (p.a.), hydrogen peroxide (30% w/w) and formic acid (purity higher than 98%) were purchased from Fluka Chemie (Madrid, Spain). Sulfuric acid (96% purity) was purchased from Acros Organics (Odivelas, Portugal). HPLCgrade methanol, water, acetonitrile and chloroform were obtained from Fisher Scientific Chemicals (Loures, Portugal). Solvents were filtered using a Solvent Filtration Apparatus 58061 (Supelco, Bellefonte, PA, USA). All other reagents were used without further purification. 2.2. Oxidation of linoleic acid Fenton’s reactions were performed in an aqueous solution (final volume = 10 mL) inside a Micro-Reaction Vessel (Supelco), by adding linoleic acid, followed by the addition of aqueous hydrogen peroxide (0.98 M) and FeCl2 (25 mM) to reach 8.5 mM, 0.1 M and 0.1 mM, respectively, in the reaction media. The reactions were favoured by the formation of LA micelles in the aqueous solution.

35 30


Fig. 1. Mechanism for fatty acid hydroperoxides formation by Fenton’s reaction.


25 20 15 10 5 0









Time (hours) Fig. 2. Formation of the LA–OOH during the course of the Fenton’s reaction.

Micelles generation was promoted by dissolving the weighed LA in CHCl3 in the reactor, which was then evaporated with a N2 stream to favour the fixation of the acid to the reactor walls thus increasing its contact surface, followed by the addition of the other reagents and, finally, by sonication of the mixture. The control was prepared by replacing the H2 O2 and FeCl2 volumes by milli-Q water. The mixture was left to react in the dark at 40 ◦ C for different periods of time (3, 6, 9, 12, 24 and 32 h) with occasional sonication to regenerate the micelles. After those periods, the oxidation products were extracted using a modification of the Folch’s method (Folch et al., 1957): 400 ␮L of the reaction mixture, collected under stirring (5.6 × g), were extracted with 1000 ␮L of chloroform/methanol (1:1, v/v), in 1.5 mL Axygen’s microcentrifuge tubes, which were submitted to centrifugation (4105 × g) for 3 min to achieve a good phase separation. Then, the water was carefully removed with a micropipette and the organic phase was introduced into a 1.5 mL amber glass vial, dried with N2 and stored at −20 ◦ C. Enzymatic oxidations were carried out with a commercial lipoxygenase (LOX) from Glycine max (soybean), type I-B, lyophilized powder supplied by Sigma Chemicals Co. (Madrid, Spain). The enzymatic reactions were performed in a MicroReaction Vessel in milli-Q water, by adding linoleic acid to reach a final concentration of 8.5 mM, and using a LOX’s dose of 60 units/mL of reaction mixture. The enzyme was weighed in a microcentrifuge tube, dissolved in 2 mL of milli-Q water, and incubated for 5 min. Simultaneously, dispersions of LA were prepared by micelles generation as described in the Fenton’s system, but only using milli-Q water (8 mL). Finally, the enzyme was added to the LA dispersion to reach a final volume of 10 mL. The mixture was left to react in the dark at 25 ◦ C, under stirring (5.6 × g) for 16 h.

2.3. Direct analysis of the reaction mixtures by electrospray ionization–mass spectrometry (ESI–MS) The ESI–MS and ESI–MS/MS analyses of the reaction mixtures were carried out in two equipments: a QqQ Quattro (Micromass) using MassLynx software (version 4.0) and a linear ion trap (LIT) LXQ (ThermoFinnigan, San Jose, CA, USA) using Xcalibur software (version 2.0.7). All MS spectra were obtained at least in triplicate, in the negative mode and the collision energy used was optimized to reduce the relative abundances of precursor ions of approximately 15%, varying the normalised collision energy between 15 and 40 eV for QqQ and between 18 and 22 (arbitrary units) for LIT.


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Fig. 3. ESI–MS spectra of the reaction mixture before (A) and after (B) Fenton’s oxidation of LA, together with the ESI–MS/MS spectrum of the ion at m/z 311 (C) obtained in the linear ion trap.

2.4. Triple quadrupole QqQ Quattro (Micromass) ESI–MS/MS analyses conditions

mass spectra, ranging from m/z 100 to 1000, and each spectrum, were produced by accumulating data during 1 min.

Samples were prepared for analyses by dissolving the compounds contained in the amber glass vial with 1000 ␮L of CHCl3 . Then 40 ␮L of this sample were diluted with 160 ␮L of methanol and introduced into the electrospray source at a flow rate of 0.6 mL min−1 , setting the needle and cone voltage at 30 V, ion source at 80 ◦ C and desolvation temperature at 150 ◦ C. Tandem mass spectra of the molecular ions were obtained by collision induced dissociation (CID), using argon as collision gas. The gas pressure in the Q2 collision cell was approximately 3.85 × 10−4 mbar. Full scan

2.5. Linear ion trap (LIT) LXQ (ThermoFinnigan) ESI–MS/MS analyses conditions Samples for electrospray analyses were prepared by diluting 2 ␮L of the sample previously dissolved with 1000 ␮L of CHCl3 , in 198 ␮L of methanol. Ion trap mass analyzers main advantages, when compared with the triple quadrupole instrument used, are the higher sensitivity (Lacorte and Fernandez-Alba, 2006), hence the need for further dilution. Electrospray conditions were as

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m/z = 183


· · ·· · ·· ·· ·· ····· ···





m/z = 183


· · ·· · ·· ·· ·· ····· ···




OH Fig. 4. Characteristic fragmentation pathways of oxidized products from LA: (A) hydroperoxides and (B) diols.

follows: nitrogen sheath gas was set at 30 psi along with spray voltage of 4.7 kV, heated capillary temperature of 275 ◦ C, capillary voltage of 35 V, and tube lens voltage of 125 V. MS2 experiments were performed on mass selected precursor ions using standard isolation and excitation procedures (activation q value of 0.25 and activation time of 30 ms). Full scan mass spectra, ranging from m/z 100 to 1500, and each spectrum, were produced by accumulating data during 1 min. 2.6. HPLC–UV and HPLC–MS/MS analysis of the reaction mixtures Analyses were carried in a Hewlett-Packard (HP) 1050 liquid chromatograph (Agilent Technologies, Waldbronn, Germany) equipped with a Rheodyne injector with a 10 ␮L loop, a quaternary pumping system and a UV detector. The optimal chromatographic conditions for LA oxidation products separation, after dissolving them in acetonitrile, were achieved using a C-18 column (15 cm × 2.1 mm × 5 ␮m supplied by Supelco), and a flow rate of 0.3 mL min−1 . The composition of the eluent was the following: Eluent A: 0.1% of HCOOH in 90.8% of water and 9.1% of acetonitrile/Eluent B: 0.1% of HCOOH in acetonitrile. Elution started with 100% A and 0% B for 5 min, and the eluent composition was ramped to 40% B in 5 min, and finally to 90% B in 30 min. UV detection was carried out at 234 nm. The HPLC system was also coupled to the triple quadrupole mass spectrometer described above, operating in negative ion mode. The cone and capillary voltages were set at 30.0 V and 2.9 kV, respectively. The source temperature was 143 ◦ C and the desolvation temperature was 350 ◦ C. MS/MS spectra were obtained using argon as collision gas with the collision energy set at 25 V. The detection was carried out considering a mass range of 50–1000 m/z. 2.7. Detection and quantification of HPODE by ferrous oxidation-xylenol orange method and HPLC–FOX FOX reagent solution (100 mL) containing 250 ␮M ammonium ferrous sulfate, 100 ␮M xylenol orange (XO), 4 mM 2,6-di-tertbutyl-4-methylphenol (BHT), 25 mM H2 SO4 and 90% methanol (HPLC grade), being the rest of the volume completed with H2 O milli-Q, was prepared (Bou et al., 2008). For the preparation of this solution it should be taken into account that Fe+2 rapidly converts to Fe+3 at pH above 7; so, the ammonium ferrous sulfate has to be weight and directly dissolved in H2 O milli-Q acidified with H2 SO4 , and then dissolve the XO in this solution. Also, it is advisable to dissolve apart the BHT, with some of the MeOH, due to its difficulty to be dissolved.

Fig. 5. Typical HPLC–UV chromatogram of the reaction mixtures issued from Fenton’s oxidation of linoleic acid, and intensity of the FOX measurement of the samples collected from the HPLC effluent. *Artifact from the chromatographic system.

The quantification of total HPODE, by the FOX method, was carried out by adding 950 ␮L of FOX reagent to 50 ␮L of the reaction mixture, obtained after extraction with the modified Folch’s extraction method referred above, but prior to the drying step. The mixtures were vortexed and incubated in the dark at room temperature for 30 min, and then the absorbance was measured at 560 nm. Quantification was carried after calibration of the FOX method using hydrogen peroxide as standard, in the range of 0–20 ␮M. The following calibration curve was obtained for HPODE quantification: Abs = 0.0310 × [HPODE]/␮M + 0.0079 (r2 : 0.9992). The FOX method was also used in combination with the HPLC–UV fractionation of HPODE, in order to obtain a more accurate quantification of individual hydroperoxides. The HPLC fractionation was carried out under the chromatographic conditions described above, dissolving the stored sample in the amber glass vial with 50 ␮L of acetonitrile, and collecting fractions of 0.15 mL (0.5 min elution). The hydroperoxide contents of these samples were then measured by the FOX method, as described above. To carry out the external calibration with the HPLC–UV using the FOX results, the starting sample was dissolved with different volumes of acetonitrile (between 250 and 2000 ␮L), and the FOX quantification was carried out by adding 650 ␮L of FOX reagent to 350 ␮L of the fraction collected between 24.4 and 26.4 min, where the hydroperoxides eluted after HPLC separation and UV detection. The mixtures were again vortexed, incubated and measured as described above. This allowed obtaining a correlation between the global area of unsaturated fatty acid hydroperoxides (UFA–OOH) in the HPLC–UV chromatogram and their abundance. All quantitative analysis by FOX and HPLC were performed at least in triplicate and accepted when the variability between analyses was lower than 5%. Quantitative results presented here are the average of the triplicate analysis of two distinct samples for each concentration level.

3. Results and discussion Iron is well known to be an important component of biological free radical oxidation systems, and the mechanism that has been generally accepted for Fe2+ role in the Fenton’s reaction leading to hydroperoxides is shown in Fig. 1 (Frankel, 1984; Qian and Buettner, 1999). In the present study, Fenton’s system was used to obtain mixtures of linoleic acid hydroperoxides that could be used to implement suitable HPLC–MS conditions for their efficient chromatographic separation and identification/quantification.


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Fig. 6. HPLC–MS (A), RIC m/z 295 (B) and RIC m/z 311 (C) of the Fenton’s reaction mixture.

The analysis of the reaction mixtures with the FOX reagent allowed to observe an increase in the amount of HPODE formed until 12 h of reaction. After that, the total amount of HPODE remained constant around 2.55 mM, which corresponds to approximately 30% of the initial total amount of LA (Fig. 2). Therefore, in order to maximise the amount of HPODE for ESI–MS and HPLC–MS analysis, these were carried out with samples obtained after 16 h of reaction. The formation of HPODE was further confirmed by direct analysis of the reaction mixtures by ESI–MS/MS. The ESI–MS spectra of the starting reaction mixture (Fig. 3A) only showed the LA [M−H]− ion at m/z 279, while after 12 h, apart from that ion, another abundant ion at m/z 311, 32 mass units higher than LA, and compatible with the addition of two oxygen atoms (formation of an –OOH group or a diol) was observed (Fig. 3B). Despite the fact that the

presence of HPODE was confirmed by the FOX essay, the MS/MS spectra of the ion at m/z 311 (Fig. 3C) allows to observe a product ion at m/z 183 resulting from C7–C8 cleavage, which however is compatible with both hydroperoxide and diol LA derived structures (Fig. 4). It was therefore necessary to analyse the reaction mixtures by HPLC–UV and HPLC–ESI–MS/MS in order to qualitatively and quantitatively analyse the reaction mixtures, particularly for the presence of HPODE. The HPLC–UV analysis (Fig. 5) of the reaction mixtures showed the presence of four intense peaks, of which, based on the FOX analysis of the fractions collected from the HPLC, the last three should correspond to hydroperoxides (Fig. 5). To achieve unambiguous identification of the detected HPODE isomers, it was necessary to carry out the individual MS/MS analysis of each chromatographic peak. The total ion chromatogram

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Fig. 7. HPLC–ESI–MS/MS spectra of the ions at m/z 311 observed in the Fenton’s reaction mixture from LA and corresponding with the HPLC–UV peaks (Fig. 5) at 24.7 (A: 13-Z,E-HPODE), 25.2 (B: 13-E,E-HPODE) and 25.7 (C: 9-HPODE) min.


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Table 1 Quantitative analysis of HPODE isomers obtained upon biomimetic and enzymatic oxidation of linoleic acid (LA).

y = 18673.4641x + 11623.9389




r = 0.9971













μM HPODE Fig. 8. Calibration curve for the HPLC–UV quantification of HPODEs, based on the FOX measurement of samples collected between 24.5 and 26.5 min.

obtained after HPLC–ESI–MS of the reaction mixtures (Fig. 6A), and the corresponding RICs of the ions m/z 295 [M−H + 16]− (Fig. 6B) and m/z 311 [M−H + 32]− (Fig. 6C), reconfirmed the presence of oxidized LA structures. Specifically, the first HPLC–UV peak at 23.6 min (Fig. 5) corresponds to a hydroxyoctadecadienoic acid (HODE) while the peaks at 24.7, 25.2 and 25.7 min correspond to HPODE. The ions detected at m/z 315 and 333 suggest the presence of sodium ion in the dehydrated precursor ions [M+Na−H2 O–2H]− and the precursor ions [M+Na–2H]− . The unambiguous distinction of the three HPODE isomers was achieved by HPLC–ESI–MS/MS (Fig. 7A–C). Thus, the chromatographic peaks observed at 24.7 and 25.2 min (Fig. 5) are the 13hydroperoxy-(9Z,11E)-octadecadienoic acid (13-Z,E-HPODE) and 13-hydroperoxy-(9E,11E)-octadecadienoic acid (13-E,E-HPODE), respectively. The ESI–MS/MS [M−H]− fragmentation profile of these compounds present the most abundant product ion at m/z 113 after loss of water and cleavage of the allylic double bond to the hydroperoxide moiety. Other characteristic product ions at m/z 179 and 195 come from losses of CO2 and CO (Dufour and Loonis, 2005; MacMillan and Murphy, 1995; Tyurin et al., 2009). However, the relative abundance of the product ion at m/z 223, which results from C13-C14 cleavage, being higher in the 13-E,E-HPODE than in the 13-Z,E-HPODE, is the key to distinguish the two isomers (Dufour and Loonis, 2005; MacMillan and Murphy, 1995; Tyurin et al., 2009) (Fig. 7A and B). The HPLC–UV peak at 25.7 min (Fig. 5) was identified as the 9-hydroperoxyoctadecadienoic acid (9-HPODE) due to its characteristic product ions at m/z 125, 141, 185 and 197 (Fig. 7C) (Dufour and Loonis, 2005; MacMillan and Murphy, 1995; Tyurin et al., 2009). Decarboxylation and decarbonylation give the two first ions, respectively, while loss of water and cleavage of the allylic double bond to the hydroperoxide moiety lead to the formation of a typical ion at m/z 185. The ion at m/z 171, resulting from the C9–C10 scission, followed by the elimination of water, was also observed (Fig. 7C). However, it is impossible to distinguish, based on their characteristic fragment ions in terms of m/z and intensity (Dufour and Loonis, 2005), if this peak (at 25.7 min) corresponds to the 10E,12Z or 10E,12E 9-hydroperoxyoctadecadienoic acid isomers (i.e. 9-E,Z-HPODE and 9-E,E-HPODE, respectively), or if it is a mixture of both. After the unambiguous identification of isomeric HPODE, it was necessary to accomplish their individual quantification. For this, it was necessary to take into account the difficulties arising from the differences existing in the molar extinction coefficients of the different UFA–OOH in the UV spectrum, and from the extreme difficulty in fully drying these UFA–OOH without degrading them,





Abundance (mM) Relative %

1.5 54.9

0.6 23.0

0.6 22.1


Abundance (mM) Relative %

0.8 82.5

0.1 9.2

0.1 8.3

making difficult the preparation of standard solutions (Haefliger and Sulzer, 2007). So, in order to obtain a more reliable quantification methodology, an external calibration of the HPLC–UV was carried out, measuring the total amount of HPODE in the HPLC effluent, collected between 24.5 and 26.5 min. The HPODE contents measured by the FOX method revealed a linear relation with the total area of the three chromatographic peaks (Fig. 8). Therefore, this experimental procedure allows carrying out the individual quantification of each HPODE isomer and avoids the use of standards which, in addition to the problems already mentioned above, are expensive. The quantitative results obtained are shown in Table 1, showing that 13-Z,E-HPODE, 13-E,EHPODE and 9-HPODE account for 1.5, 0.6 and 0.6 mM, respectively, in the Fenton’s reaction media after 16 h. Furthermore, 13-Z,EHPODE is the predominant isomer representing ∼55% of the formed hydroperoxides. Finally, samples from the enzymatic oxidation of linoleic acid with a commercial lipoxygenase from G. max (soybean) were analyzed through this methodology. The chromatographic profile obtained for the enzymatic essay (Fig. 9) is qualitatively similar to that obtained with the biomimetic Fenton’s system, in terms of the hydroperoxides formed and even the HODE eluting at 23.6 min. The HPLC analysis showed that 13-Z,E-HPODE is also the predominant isomer (83% of the HPODEs formed). Finally, the quantification of the individual HPODEs using the proposed FOX calibration of the HPLC system showed that 13-Z,E-HPODE, 13-E,EHPODE and 9-HPODE account for 0.8, 0.1 and 0.1 mM, respectively, in the enzymatic reaction media after 16 h. Furthermore, the proportion of the positional isomers 13-HPODE:9-HPODE (91.7:8.3) and geometrical isomers 13-Z,E-HPODE:13-E,E-HPODE (90:10) are similar to that observed by Nikolaev et al. (1990), confirming definitely the validity of the method. Concluding, this study demonstrates a simple and integrated methodology for the identification and quantification of linoleic acid hydroperoxides that can be applied for biomimetic and enzymatic systems. This methodology can now be applied to

Fig. 9. Typical HPLC–UV chromatogram of the reaction mixtures issued from the enzymatic oxidation (Soybean LOX) of linoleic acid. *Artifact from the chromatographic system.

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adequately study different enzymatic systems and also experimentally extrapolated to the quantification of other unsaturated fatty acid hydroperoxides and, therefore, to contribute to the valorization of unsaturated fatty acids through hydroperoxide intermediates. Acknowledgments FCT and EU are acknowledged for financial support of the ERANOEL Project (Novel enzyme tools for production of functional oleochemicals from unsaturated lipids – ERA-IB/BIO/0001/2008), which also funds J.J. Villaverde Postdoctoral Grant. References Belgacem, M.N., Gandini, A., 2008. Materials from vegetable oils: major sources, properties and applications. In: Belgacem, M.N., Gandini, A. (Eds.), Monomers, Polymers and Composites from Renewable Resources. Elsevier, pp. 39–66. Bou, R., Codony, R., Tres, A., Decker, E.A., Guardiola, F., 2008. Determination of hydroperoxides in foods and biological samples by the ferrous oxidation–xylenol orange method: a review of the factors that influence the method’s performance. Anal. Biochem. 377, 1–15. Corma, A., Iborra, S., Velty, A., 2007. Chemical routes for the transformation of biomass into chemicals. Chem. Rev. 107, 2411–2502. Dufour, C., Loonis, M., 2005. Regio- and stereoselective oxidation of linoleic acid bound to serum albumin: identification by ESI–mass spectrometry and NMR of the oxidation products. Chem. Phys. Lipids 138, 60–68. Fernando, S., Adhikari, S., Chandrapal, C., Murali, N., 2006. Biorefineries: current status, challenges, and future direction. Energy Fuels 20, 1727–1737. Folch, J., Lees, M., Stanley, G.H.S., 1957. A simple method for the isolation and purification of total lipides from animal tissues. J. Biol. Chem. 226, 497–509. Frankel, E.N., 1984. Lipid oxidation: mechanisms, products and biological significance. J. Am. Oil Chem. Soc. 61, 1908–1917. Frankel, E.N., 2005. Lipid Oxidation, second ed. The Oily Press. Gandini, A., Pascoal Neto, C., Silvestre, A.J.D., 2006. Suberin: a promising renewable resource for novel macromolecular materials. Prog. Polym. Sci. 31, 878–892. Gardner, H.W., 1989. Oxygen radical chemistry of polyunsaturated fatty acids. Free Radic. Biol. Med. 7, 65–86. Gorkum, R., Bouwman, E., 2005. The oxidative drying of alkyd paint catalysed by metal complexes. Coord. Chem. Rev. 249, 1709–1728.


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