Quantitation of Acyl Chain Oxidation Products Formed upon Thermo

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Quantitation of Acyl Chain Oxidation Products Formed upon Thermo-Oxidation of Phytosteryl/-stanyl Oleates and Linoleates Stefan Wocheslander, Franziska Groß, Birgit Scholz, and Karl-Heinz Engel J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.7b00424 • Publication Date (Web): 03 Mar 2017 Downloaded from http://pubs.acs.org on March 5, 2017

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Journal of Agricultural and Food Chemistry

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Quantitation of Acyl Chain Oxidation Products Formed upon Thermo-

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Oxidation of Phytosteryl/-stanyl Oleates and Linoleates

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Stefan Wocheslander, Franziska Groß, Birgit Scholz and Karl-Heinz Engel*

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Lehrstuhl für Allgemeine Lebensmitteltechnologie, Technische Universität München,

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Maximus-von-Imhof-Forum 2, D-85354 Freising, Germany

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*Corresponding Author: Phone +49 (0)8161 71 4250. Fax +49 (0)8161 71 4259. E-mail:

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[email protected]

1 ACS Paragon Plus Environment


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ABSTRACT

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A GC-based approach involving pre-separation via solid phase extraction was established for

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the quantitation of acyl chain oxidation products (ACOPs) formed upon thermo-oxidation

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(180 °C, 40 min) of oleates and linoleates of phytostanols and phytosterols. The

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concentrations of ACOPs resulting from initially formed 9-hydroperoxides (octanoates, 8-

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hydroxyoctanoates, 9-oxononanoates) were higher than those from 8-hydroperoxides

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(heptanoates, 7-hydroxy- and 7-oxoheptanoates, 8-oxooctanoates) in both oleates and

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linoleates. Significantly higher amounts of ACOPs were found in heat-treated linoleates

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compared to oleates. However, despite lower thermally induced losses of stanyl oleates and

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linoleates compared to the respective steryl esters higher concentrations of ACOPs (approx.

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9% and 10% of the ester losses, respectively) were observed in the heat-treated stanyl esters.

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In contrast, in the heated steryl oleates and linoleates the contribution of the ACOPs to the

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ester losses was lower (approx. 3% and 5%, respectively), and there was a more pronounced

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formation of oxidation products of the sterol moieties (approx. 26% and 18% of the ester

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losses, respectively).

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Keywords: Phytosteryl ester oxidation, core aldehydes, lipid autoxidation


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INTRODUCTION

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Due to their cholesterol-lowering properties, phytosterols and phytostanols are used as

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functional ingredients for the enrichment of various foods.1,2 For technological reasons, such

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as the improved solubility in lipid matrices, the fatty acid esters rather than the free

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phytosterols/-stanols are added to lipid-based foods.3 Mixtures of saturated and unsaturated

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fatty acids derived from edible oil sources are employed for the esterification step. Similarly

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to cholesterol and its fatty acid esters, phytosteryl/-stanyl fatty acid esters are prone to

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oxidation reactions.4 Previous studies mainly focused on investigations of oxidation products

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of the phytosterol moieties. There is rather detailed knowledge on the formation of the

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principal derivatives, i.e. 7-keto, 7-hydroxy and 5,6-epoxy compounds, generally termed as

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“phytosterol oxidation products” (POPs). Comprehensive data on the formation of POPs upon

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heat treatment of enriched foods5 and recent reviews on daily intakes and biological effects

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are available.6,7

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However, phytosteryl/-stanyl esters particularly of unsaturated fatty acids are also susceptible

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to oxidations in the fatty acid moieties. The knowledge on these types of oxidative

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modifications has been rather limited. The formation and the degradation of stigmasteryl ester

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hydroperoxides have been investigated.8 In another study the formation of sitosteryl and 7-

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ketositosteryl 9,10-dihydroxystearate upon heating of a phytosteryl ester-enriched margarine

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has been reported.9 Only recently, other acyl chain oxidation products (ACOPs) resulting

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from thermo-oxidation of a mixture of phytosteryl/-stanyl linoleates have been identified. The

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oxidation products covered by the employed GC-based methodology comprised three classes:

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phytosteryl/-stanyl

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hydroxyheptanoates,

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oxooctanoates, 9-oxononanoates).10 Representative structures are shown in Figure 1. The

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study provided an analytical platform to identify ACOPs of phytosteryl-stanyl fatty acid esters

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without the need for a hydrolysis or transesterification step resulting in a loss of structural

alkanoates

(heptanoates,

8-hydroxyoctanoates)

and

octanoates),

hydroxyalkoanoates

oxoalkanoates

(7-oxoheptanoates,

(78-



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information. This significantly extended the knowledge on these types of oxidation products;

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however, quantitative data and thus an estimation of their contribution to the decreases of

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phytosteryl/-stanyl fatty acid esters were still missing. Therefore, the objective of the present

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study was to develop an approach for the quantitation of these ACOPs and to determine their

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amounts upon thermal treatment of phytosteryl/-stanyl esters of unsaturated fatty acids

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contained in mixtures commonly used to enrich foods.

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MATERIALS AND METHODS

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Chemicals and Reagents

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A mixture of phytosterols/-stanols, named “β-sitosterol” and consisting of 75% sitosterol,

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12% sitostanol, 10% campesterol, 2% campestanol and 1% others, was purchased from Acros

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Organics (Morris Plains, NJ, USA). A mixture of phytostanols, named “stigmastanol” and

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consisting of 98 % sitostanol and 2 % campestanol, was purchased from Santa Cruz

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Biotechnology (Dallas, TX, USA). Linoleic acid (≥ 99%), oleic acid (≥ 99%), pyridine

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(anhydrous, ≥ 99.8%), cholesterol (95%), cholesteryl palmitate (≥ 97%), 5,6β-

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epoxycholesterol (≥ 95%), 7-ketocholesterol (≥ 90%), lactic acid (≥ 90%) and acetic

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anhydride (≥ 99%) were purchased from Sigma-Aldrich (Steinheim, Germany). The silylation

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reagent BSTFA/TMCS (99:1) was purchased from Supelco (Bellefonte, PA, USA). Methyl

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tert-butyl ether (MTBE) and cyclohexane (analytical grade) were supplied by Evonik

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Industries AG (Essen, Germany). n-Hexane (HiPerSolv Chromanorm) and chloroform

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(AnalaR Normapur) were purchased from VWR International (Darmstadt, Germany).

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Synthesis of Phytosteryl/-stanyl Oleates and Linoleates

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The synthesis of phytosteryl/-stanyl oleates and linoleates was carried out as previously

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described.10,11 Briefly, 100 mg of the phytosterol/-stanol sample “β-sitosterol” and of the

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phytostanol sample “stigmastanol”, respectively, were treated as follows: 170 mg oleic or 4 ACS Paragon Plus Environment

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linoleic acid were placed in a sealed, nitrogen-flushed (in order to avoid oxidation) 2 mL glass

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vial and heated for 25 h at 180 °C. The excess of acid was removed with 2.5 mL potassium

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hydroxide (1 M), and the esters were extracted with 3 × 2.5 mL hexane/MTBE (3:2, v/v).

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Final purification on a 500 mg Supelclean silica SPE column from Supelco (Bellefonte, PA,

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USA) with cyclohexane as an eluent resulted in 107 mg phytosteryl/-stanyl oleates (yield =

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65%), 94 mg phytostanyl oleates (yield = 58%), 79 mg phytosteryl/-stanyl linoleates (yield =

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48%) and 72 mg phytostanyl linoleates (yield = 44%). The phytosterol/-stanol distributions

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corresponded to those of the phytosterol/-stanol samples used for esterification. The purity of

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the esters was determined by 1H NMR and GC-FID and was ≥ 98% for all ester mixtures.

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Synthesis of Oxidized Cholesteryl Esters as Internal Standards

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Cholesteryl octanoate and cholesteryl 9-oxononanoate were synthesized via esterification with

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the respective acid, and cholesteryl 9-hydroxynonanoate was obtained by reduction of

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cholesteryl 9-oxononanoate with sodium borohydride as described in a previous publication

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for the corresponding sitosteryl esters.10 The NMR spectra of cholesteryl octanoate and

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cholesteryl 9-oxononanoate corresponded to previously published data.12,13 The identity of

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cholesteryl 9-hydroxynonanoate was confirmed by two-dimensional NMR of the fatty acid

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part (13C/1H): δ 173.29 (FA-1), 63.06/3.66 (FA-9, 2H, t), 34.70/2.29 (FA-2, 2H, t), 32.71/1.58

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(FA-8, 2H, m), 29.22/1.35 (FA-4/5/6, 2H, m), 29.22/1.35 (FA-4/5/6, 2H, m), 29.03/1.35 (FA-

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4/5/6, 2H, m), 25.66/1.64 (FA-7, 2H, m), 25.02/1.65 (FA-3, 2H, m). No impurities were

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observed for cholesteryl octanoate (GC-FID and 1H NMR); cholesteryl 9-oxononanoate and

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cholesteryl 9-hydroxynonanoate had a purity of ≥ 92%.

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Thermal Treatment of Phytosteryl/-stanyl Oleates and Linoleates

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Each ester mixture (11 ± 0.2 mg) was weighed into a 2 mL brown glass vial without a lid and

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oxidized in a heating block (VLM Metal, Bielefeld, Germany) at 180 °C for 40 min. After 5 ACS Paragon Plus Environment


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cooling to room temperature, the sample was dissolved in n-hexane/MTBE (3:2, v/v) and

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transferred to a graduated flask with a final volume of 5 mL. For each ester mixture, the

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heating experiment was performed in triplicate (n=3).

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Quantitation of Nonpolar ACOPs and Nonoxidized Phytosteryl/-stanyl Oleates and

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Linoleates

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An aliquot (0.5 mL) of the sample (5 mL) obtained after heat treatment was transferred into a

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2 mL brown glass vial, and 20 µL cholesteryl octanoate (0.78 mg/mL) and 100 µL cholesteryl

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palmitate (2.5 mg/mL) were added as internal standards. After removal of the solvent under a

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gentle stream of nitrogen, the residue was dissolved in 250 µL hexane/MTBE (3:2, v/v). This

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sample was directly subjected to GC-FID/MS analysis for the determination of nonpolar

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oxidation products. Their identification was based on retention times and mass spectra as

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previously described,10 quantitation was performed relative to the internal standard cholesteryl

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octanoate using a response factor (Rf) of 1.0 as default value.

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For the quantitation of nonoxidized oleates and linoleates the samples were further diluted

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(between 1:3 and 1:8), depending on the amounts of nonoxidized esters remaining after

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thermal treatment. Subsequently, the samples were analyzed according to a previously

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established method based on GC-FID/MS,11 and quantitated using cholesteryl palmitate as

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internal standard.

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Quantitation of Polar ACOPs

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An aliquot (4 ml for phytostanyl ester mixtures and 2 mL for phytosteryl/-stanyl ester

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mixtures) of the sample (5 mL) obtained after heat treatment was transferred into a 12 mL

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glass vial. Subsequently, the following volumes of stock solutions of the internal standards

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cholesteryl 9-hydroxynonanoate (2.17 mg/mL) and cholesteryl 9-oxononanoate (1.03 mg/mL)

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were added: phytostanyl oleates (10 µL and 10 µL); phytostanyl linoleates (40 µL and 20 6 ACS Paragon Plus Environment

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µL); phytosteryl/-stanyl oleates (5 µL and 5 µL); phytosteryl/-stanyl linoleates (10 µL and 10

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µL). After removal of the solvent under a gentle stream of nitrogen, the polar oxidation

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products were separated from nonoxidized esters by silica solid-phase extraction as described

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recently.10 Briefly, nonoxidized esters and nonpolar oxidation products were eluted from a

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silica SPE-cartridge with cyclohexane (fraction 1), followed by the elution of the polar

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oxidation products with MTBE (fraction 2). The residue of the polar MTBE fraction was

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silylated with 100 µL pyridine and 100 µL BSTFA/TMCS for 20 min at 80 °C. The silylation

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reagent was evaporated under a gentle nitrogen stream, the residue was dissolved in 100-400

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µL hexane/MTBE (3:2, v/v) and subjected to GC-FID/MS analysis. Identification of polar

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oxidation products was based on retention times and mass spectra,10 quantitation was done

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relative to the internal standards using a response factor (Rf) of 1.0 as default value.

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Method Validation

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The limits of detection (LOD) and quantification (LOQ) were determined according to the

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method of Vogelgesang and Hädrich14 for sitosteryl octanoate, sitosteryl 9-hydroxynonanoate

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(silylated) and sitosteryl 9-oxononanoate, as representatives of the three classes of ACOPs

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covered by the employed methodology.10

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Linearity in the working range was validated by establishing five point calibration curves in

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the range from 3-73 µg/ml for cholesteryl octanoate and sitosteryl octanoate, 11-217 µg/ml

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for cholesteryl 9-hydroxynonanoate (silylated) and sitosteryl 9-hydroxynonanoate (silylated),

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and 10-191 µg/ml for cholesteryl 9-oxononanoate and sitosteryl 9-oxononanoate. The

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equations of the calibration curves were determined by linear regression analysis of the peak

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area versus the concentration of the analyte.

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For the recovery of polar oxidation products, known amounts of sitosteryl 9-

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hydroxynonanoate in the range of 15-82 µg and of sitosteryl 9-oxononanoate in the range of



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23-115 µg were spiked to 1.5 mg sitosteryl linoleate (non-heated) in triplicate. Subsequently,

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the samples were taken through the SPE-procedure as described above.

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The reproducibility of the method including the thermo-oxidation and sample preparation step

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was tested by repeated analysis of heated phytostanyl linoleates on another day by a second

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operator in triplicate (n=3).

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Quantitation of Phytosterol Oxidation Products

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An aliquot (2 mL) of a sample (5 mL) obtained after heat treatment of β-sitosteryl oleate and

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linoleate sample, respectively, was transferred into a 12 mL glass vial. After adding 50 µL

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5,6ß-epoxycholesterol (0.61 mg/mL) and 50 µL 7-ketocholesterol (0.5 mg/mL) as internal

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standards, the solvent was removed under a gentle stream of nitrogen. Transesterification,

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extraction of phytosterol oxidation products and subsequent acetylation were performed as

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previously described.15 The residue was dissolved in 0.3 mL of n-hexane/MTBE/isopropanol

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(80:20:0.3, v/v/v) and subjected to on-line LC-GC analysis.15 The on-line LC-GC system

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consisted of a 1220 Infinity LC, which was coupled to a 7890A GC equipped with a flame

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ionization (FID) via a 1200 Infinity series 2-position/6-port switching valve (Agilent

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Technologies, Böblingen, Germany). Evaporation of the solvent was performed via the

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temperature programmable multimode inlet (MMI) in the programmable temperature

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vaporizer (PTV) solvent vent mode. Identification was based on relative retention times and

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mass spectra of acetylated phytosterol oxidation products, quantification was done using

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response factors relative to the respective internal standard (1.0 for 5,6-epoxy- and 7-

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ketophytosterols and 1.2 for 7-hydroxyphytosterols).

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According to the procedures described above, the synthesized phytosteryl/-stanyl oleates and

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linoleates were also analyzed regarding the presence of ACOPs and POPs before the thermal

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treatments.

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GC-FID Analysis

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Analysis was performed using a 6890N GC equipped with an FID (Agilent Technologies,

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Böblingen, Germany). The separations were carried out on a 30 m × 0.25 mm i.d., 0.1 µm

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film, Rtx-200MS fused silica capillary column (Restek, Bad Homburg, Germany). The

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temperature of the injector was set to 280 °C, and hydrogen was used as carrier gas with a

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constant flow rate of 1.5 ml/min. The split flow was set to 11 ml/min, resulting in a split ratio

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of 1:7.5. The oven temperature was programmed as follows: Initial temperature, 100 °C;

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programmed at 15 °C/min to 310 °C (2 min), then 1.5 °C/min to 340 °C (3 min). The FID

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temperature was set to 340 °C and nitrogen was used as a makeup gas with a flow rate of 25

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ml/min. The injection volume was set to 1 µL. Data acquisition was performed by

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ChemStation B.04.03.

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GC-MS Analysis

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Mass spectra were recorded with a Finnigan Trace GC ultra coupled with a Finnigan Trace

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DSQ mass spectrometer (Thermo Electro Corp., Austin, Texas, USA). Mass spectra were

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obtained by positive electron impact ionization at 70 eV in the scan mode at unit resolution

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from 40 to 750 Da. The interface was heated to 330 °C and the ion source to 250 °C. Helium

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was used as carrier gas with a constant flow rate of 1.0 ml/min. The other GC conditions were

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the same as described for GC-FID analysis. Data acquisition was perfomed by Xcalibur 3.063

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software.

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NMR Spectroscopy

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The compounds under study were dissolved in 0.5 mL deuterated chloroform. 1H NMR and

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13

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500 spectrometers operating at 27 °C. 1H-Detected experiments including two-dimensional

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COSY, NOESY, HSQC and HMBC were measured with an inverse 1H/13C probehead, direct

C NMR spectra were recorded at 500 MHz and 126 MHz, respectively with Avance-HD



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13

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experiments were done in full automation using standard parameter sets of the TOPSPIN 3.2

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software package (Bruker, Billerica, MA, USA).

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decoupled mode. Data processing was done with the MestreNova software.

C-measurements were performed with a QNP

13

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C/31P/29Si/19F/1H cryoprobe. All

C NMR spectra were recorded in proton-

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Statistical Analysis

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The software RStudio (version 0.99.903) was used for statistical analyses. Statistical

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differences were evaluated by one-way ANOVA and unpaired Student’s t-test, respectively.

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Levels of significance: p < 0.001, highly significant (***); p < 0.01, very significant (**); p

Quantitation of Acyl Chain Oxidation Products Formed upon Thermo

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