Identification of Acyl Chain Oxidation Products upon Thermal Treatment of a Mixture of Phytosteryl/-stanyl Linoleates

Article abstract of DOI:10.1021/acs.jafc.6b04326

A mixture of phytosterols/-stanols, consisting of 75% β-sitosterol, 12% sitostanol, 10% campesterol, 2% campestanol, and 1% others, was esterified with linoleic acid. The resulting mixture of phytosteryl/-stanyl linoleates was subjected to thermal oxidation at 180 °C for 40 min. A silica solid-phase extraction was applied to separate a fraction containing the nonoxidized linoleates and nonpolar degradation products (heptanoates, octanoates) from polar oxidation products (oxo- and hydroxyalkanoates). In total, 15 sitosteryl, sitostanyl, and campesteryl esters, resulting from oxidation of the acyl chain, could be identified by GC-FID/MS. Synthetic routes were described for authentic reference compounds of phytosteryl/-stanyl 7-hydroxyheptanoates, 8-hydroxyoctanoates, 7-oxoheptanoates, 8-oxooctanoates, and 9-oxononanoates, which were characterized by GC-MS and two-dimensional NMR spectroscopy. The study provides data on the formation and identities of previously unreported classes of acyl chain oxidation products upon thermal treatment of phytosteryl/-stanyl fatty acid esters.

Full text of DOI:10.1021/acs.jafc.6b04326

Article
pubs.acs.org/JAFC
Identification of Acyl Chain Oxidation Products upon Thermal
Treatment of a Mixture of Phytosteryl/-stanyl Linoleates
Stefan Wocheslander, Wolfgang Eisenreich, Birgit Scholz, Vera Lander, and Karl-Heinz Engel*
†
‡
†
§
,†
†
Lehrstuhl fu
̈ ̈
̈
r Allgemeine Lebensmitteltechnologie, Technische Universitat Munchen, Maximus-von-Imhof-Forum 2, D-85354
Freising, Germany
‡
Lehrstuhl fu
§
̈
r Biochemie, Technische Universitat
̈
Mu
̈
nchen, Lichtenbergstrasse 4, D-85748 Garching, Germany
Bayerisches Landesamt fur Gesundheit und Lebensmittelsicherheit, Veterinar
̈
̈
strasse 2, D-85764 Oberschleissheim, Germany
ABSTRACT: A mixture of phytosterols/-stanols, consisting of 75% β-sitosterol, 12% sitostanol, 10% campesterol, 2% cam-
pestanol, and 1% others, was esterified with linoleic acid. The resulting mixture of phytosteryl/-stanyl linoleates was subjected
to thermal oxidation at 180 °C for 40 min. A silica solid-phase extraction was applied to separate a fraction containing the
nonoxidized linoleates and nonpolar degradation products (heptanoates, octanoates) from polar oxidation products (oxo- and
hydroxyalkanoates). In total, 15 sitosteryl, sitostanyl, and campesteryl esters, resulting from oxidation of the acyl chain, could
be identified by GC-FID/MS. Synthetic routes were described for authentic reference compounds of phytosteryl/-stanyl
7
-hydroxyheptanoates, 8-hydroxyoctanoates, 7-oxoheptanoates, 8-oxooctanoates, and 9-oxononanoates, which were characterized
by GC-MS and two-dimensional NMR spectroscopy. The study provides data on the formation and identities of previously
unreported classes of acyl chain oxidation products upon thermal treatment of phytosteryl/-stanyl fatty acid esters.
KEYWORDS: phytosteryl ester oxidation, solid-phase extraction, NMR spectroscopy, core aldehydes, fatty acid cleavage
INTRODUCTION
the decreased esters. This indicated that oxidative modifications
in the fatty acid moieties may account for part of the degraded
esters, particularly for those of unsaturated fatty acids.
■
Since the authorization of a yellow fat spread enriched with
phytosteryl fatty acid esters as a novel food in the European
1
The oxidation behavior of cholesteryl esters has been
Union in 2000, these functional ingredients have been added
11,12
thoroughly investigated.
In vitro studies on the oxidation
to a broad spectrum of foods due to their cholesterol-lowering
2
,3
of low- and high-density lipoprotein and isolated cholesteryl
esters with copper or tert-butyl hydroperoxide resulted in
cholesteryl ester hydroperoxides as well as so-called cholesteryl
ester core aldehydes that consist of sterols and oxysterols ester-
ified to scission products of oxidized fatty acids with terminal
properties. Fat-based foods are enriched with fatty acid
esters of phytosterols/-stanols due to their enhanced solubility
4
in lipid media. The esters are obtained by esterification of
phytosterols/-stanols with saturated as well as unsaturated fatty
acids of plant oil origin. Similarly to the structurally related
cholesterol and its fatty acid esters, phytosterols as well as their
fatty acid esters are susceptible to oxidation in the steroid ring
due to their unsaturation between C-5 and C-6. Oxidation reac-
tions of phytosterols are catalyzed by light, transition metals,
water, heat, and oxygen. Previous studies almost exclusively
focused on the analysis of secondary oxidation products, mainly
-keto, 7-hydroxy, and 5,6-epoxy derivatives of the phytosterol
moieties, which are commonly referred to as “phytosterol oxi-
5
13−15
carbonyl groups.
The most abundant cholesteryl ester core
aldehydes resulting from cholesteryl linoleate oxidation were
cholesteryl 9-oxononanoate and cholesteryl 8-oxooctanoate.
These types of oxidized fatty acid moieties, typically analyzed as
fatty acid methyl esters after transesterification, are also well-
known products formed upon thermo-oxidation and frying of
6
16−18
fats, oils, and unsaturated fatty acid methyl esters.
7
On the other hand, knowledge of the compounds result-
ing from oxidative modifications in the fatty acid parts of
phytosteryl/-stanyl esters is very scarce. There are investiga-
tions on the formation and decomposition of phytosteryl ester
6,7
dation products“ (POPs). The occurrence of POPs in enriched
foods, estimations of their daily intake, and their biological effects
have recently been reviewed. Comprehensive data sets on
8
19
hydroperoxides. Another study following the formation of
sitosteryl oleate oxidation products upon heating of phytosteryl
ester-enriched margarine reported only the presence of
the formation of POPs in foods during heating conditions
representing typical ways of preparing foods in the home are
9
available. However, the commonly used fatty acid ester mix-
20
sitosteryl 9,10-dihydroxystearate. The fact that no other acyl
chain oxidation products of phytosteryl/-stanyl fatty acid esters
have been described is probably also due to the unavailability of
the respective reference compounds.
tures of phytosterols/-stanols are prone to oxidation not only
in the steroid moieties but also in the fatty acid moieties.
In a recent study the decreases of a spectrum of individual
phytosteryl/-stanyl esters upon heating of phytosteryl/-stanyl
fatty acid ester-enriched margarines and the formation of POPs
were simultaneously followed. In the phytosteryl ester-
enriched margarine only approximately 20% of the ester losses
could be explained by the formation of POPs; in the phytostanyl
ester-enriched margarine the POPs accounted for even <1% of
10
Received: September 28, 2016
Revised: November 19, 2016
Accepted: November 20, 2016
Published: November 20, 2016
©
XXXX American Chemical Society
A
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
Therefore, the aim of the present study was to identify acyl
chain oxidation products upon heat treatment of a mixture of
phytosteryl/-stanyl linoleates, taking the existing body of
knowledge on oxidations of cholesteryl fatty acid esters and
of fats and oils into account. Crucial steps of the investigations
were (i) the development of a method suitable to separate the
nonoxidized esters from the polar oxidation products, (ii) the
synthesis of authentic reference compounds of steryl/-stanyl
esters with oxidized fatty acid moieties, and (iii) the iden-
tification of the oxidation products in the heat-treated mixture
by means of capillary gas chromatography−mass spectrometry.
MATERIALS AND METHODS
■
Chemicals and Reagents. A mixture of phytosterols/-stanols,
named “β-sitosterol” and consisting of 75% sitosterol, 12% sitostanol,
Figure 1. General structures and atom numbering of acyl chain
oxidation products of sitosteryl linoleate.
10% campesterol, 2% campestanol, and 1% others, was purchased from
Acros Organics (Morris Plains, NJ, USA). Linoleic acid (≥99%),
octanoic acid (≥98%), heptanoic acid (≥96%), pyridine (anhydrous,
(St-2, 2H, m), 27.82/1.57;1.88 (St-16, 2H, m), 26.03/1.19 (St-23, 2H,
m), 24.31/1.04;1.60 (St-15, 2H, m), 23.05/0.90 (St-28, 2H, m),
21.03/1.01;1.51 (St-11, 2H, m), 19.85/0.89 (St-26/27, 3H, d), 19.34/
1.04 (St-19, 3H, s), 19.04/0.89 (St-26/27, 3H, d), 18.79/0.94 (St-21,
3H, d), 12.00/0.89 (St-29, 3H, t), 11.87/0.70 (St-18, 3H, s). The
NMR signals of the fatty acid parts were as follows ( C/ H): sitosteryl
heptanoate δ 173.37 (FA-1), 34.74/2.29 (FA-2, 2H, t), 31.49/1.32
(FA-5, 2H, m), 28.81/1.32 (FA-4, 2H, m), 25.05/1.64 (FA-3, 2H, m),
22.51/1.31 (FA-6, 2H, m), 14.07/0.88 (FA-7, 3H,t); sitosteryl
octanoate δ 173.35 (FA-1), 34.76/2.29 (FA-2, 2H, t), 31.69/1.3
(FA-6, 2H, m), 29.1/1.32 (FA-4/5, 2H, m), 28.96/1.32 (FA-4/5, 2H,
m), 25.08/1.63 (FA-3, 2H, m), 22.62/1.3 (FA-7, 2H, m), 14.1/0.9
(FA-8, 3H, t).
≥
99.8%), trimethylsilyldiazomethane (2 M in hexane), 3-chloroper-
benzoic acid (m-CPBA, ≥77%), cyclooctanone (≥98%), cyclohepta-
none (≥98%), 2-iodoxybenzoic acid (45 wt %), sodium borohydride
(
≥96%), 4-toluenesulfonic acid monohydrate (4-TsOH, ≥98%) and
13
1
pyridinium chlorochromate (≥98%) were purchased from Sigma-
Aldrich (Steinheim, Germany). The silylation reagent BSTFA/TMCS
(
99:1) was purchased from Supelco (Bellefonte, PA, USA).
Dicyclohexylcarbodiimide (DCC, 1 M in dichloromethane, > 99%)
and 4-(dimethylamino)pyridine (DMAP, puriss.) were ordered from
Fluka Analytical (Steinheim, Germany). Aleuritic acid (9,10,16-
trihydroxypalmitic acid, ≥94%) was bought from Santa Cruz
Biotechnology (Heidelberg, Germany). Methyl tert-butyl ether
(
MTBE) was supplied by Evonik Industries AG (Essen, Germany)
Synthesis of 9-Oxononanoic Acid. 9-Oxononanoic acid was
and hexane (HiPerSolv Chromanorm) was purchased from VWR
International (Darmstadt, Germany).
synthesized according to a previously described method via diol cleav-
2
2
age of aleuritic acid (9,10,16-trihydroxypalmitic acid). Briefly, 6 g of
potassium periodate in 300 mL of sulfuric acid (1 M) was quickly
added to 8 g of aleuritic acid in methanol/water (200 + 200 mL) at
40 °C. After 10 min of stirring, the mixture was cooled to 15 °C in a
methanol/ice bath and extracted immediately with 2 × 400 mL of
diethyl ether. The combined organic layers were extracted with 2 ×
100 mL saturated sodium hydrogen carbonate solution, and the
combined aqueous layers were acidified with concentrated hydro-
chloric acid. The aqueous solution was extracted with 2 × 100 mL of
diethyl ether, and the combined ether layers were washed with 2 ×
100 mL of an aqueous sodium chloride solution (10%) and dried over
magnesium sulfate. After removal of the solvent under a nitrogen
stream, 3.2 g of a colorless oil was obtained, which was used for the
subsequent esterification step.
Synthesis of Phytosteryl/-stanyl Linoleates. The synthesis of
the mixture of phytosteryl/-stanyl linoleates was carried out according
21
to a previously described protocol. Briefly, 100 mg of the mixture of
phytosterols/-stanols (“β-sitosterol”) and 170 mg of linoleic acid were
heated in a sealed, nitrogen-flushed 2 mL glass vial for 25 h at 180 °C.
The excess of acid was removed with 2.5 mL of potassium hydroxide
(
(
(
1 M), and the esters were extracted with 3 × 2.5 mL hexane/MTBE
3:2, v/v). Final purification on a 500 mg silica solid-phase extraction
SPE) column (Supelco) with cyclohexane as an eluent resulted in
4
7 mg of a mixture of phytosteryl/-stanyl linoleates (yield = 29%) as a
colorless oil. The phytosterol/phytostanol distribution of the mixture
corresponded to that of the “β-sitosterol” sample used for ester-
ification. The purity of the linoleates determined by H NMR and
1
GC-FID was ≥98%.
Synthesis of Phytosteryl/-stanyl 9-Oxononanoates. The
Synthesis of Phytosteryl/-stanyl Octanoates and Hepta-
noates. Phytosteryl/-stanyl octanoates and heptanoates were synthe-
sized according to the same procedure as described for the linoleates.
The reaction of the phytosterol/-stanyl mixture “β-sitosterol” (100 mg)
and octanoic acid/heptanoic acid (90 mg) resulted in 50 mg of
phytosteryl/-stanyl octanoates (yield = 38%) and 61 mg of
phytosteryl/-stanyl heptanoates (yield = 48%), respectively, as white
solids. The sterol/stanol distributions corresponded to that of the
esterification was carried out as previously described for the synthesis
2
3
of cholesteryl 9-oxononanoate. 9-Oxononanoic acid (200 mg),
DMAP (5.6 mg), and the phytosterol/-stanyl mixture “β-sitosterol”
(195 mg) were dissolved in 4 mL of dichloromethane, 1 mL of DCC
solution was added, and the mixture was stirred for 24 h at room
temperature. The mixture was filtered in the dark and dissolved in
10 mL of dichloromethane, and the solution was subsequently washed
with 25 mL of hydrochloric acid (0.5 M) and 25 mL of saturated
sodium hydrogen carbonate solution. The organic layer was dried over
magnesium sulfate and filtered. For final purification, the esterified
product was dissolved in 5 mL of cyclohexane. The sample was loaded
onto a 500 mg silica SPE column, preconditioned with 6 mL of
cyclohexane. The esters were eluted with 15 mL of heptane/MTBE
(98:2, v/v). After evaporation of the solvent, 203 mg of a mixture of
phytosteryl/-stanyl 9-oxononanoates was obtained as a white solid.
The phytosterol/-stanol distribution of the mixture corresponded to
that of the “β-sitosterol” sample used for esterification. The purity of
“
β-sitosterol” sample used for esterification. No impurities were
1
observed in H NMR and GC-FID analyses. The mass spectra of
synthesized octanoates were in agreement with previously published
spectra. Full NMR signal assignments could be achieved for the
quantitatively dominating sitosteryl esters (for atom numbering, see
Figure 1). Both esters exhibited the same NMR signals for the steryl
21
13
1
moiety ( C/ H): δ 139.70 (St-5), 122.59/5.39 (St-6, 1H, d), 73.67/
.63 (St-3, 1H, m), 56.68/1.00 (St-14, 1H, m), 56.02/1.10 (St-17, 1H,
m), 50.01/0.98 (St-9, 1H, m), 45.81/0.93 (St-24, 1H, m), 42.31
St-13), 39.72/1.17;2.03 (St-12, 2H, m), 38.16/2.33 (St-4, 2H, d),
7.01/1.13;1.86 (St-1, 2H, m), 36.60 (St-10), 36.17/1.37 (St-20, 1H,
m), 33.92/1.32 (St-22, 2H, m), 31.91/1.96;2.03 (St-7, 2H, m), 31.86/
.31 (St-8, 1H, m), 29.12/1.32 (St-25, 1H, m), 28.27/1.59;1.87
4
1
(
3
the oxoesters was determined to be ≥92% by H NMR and GC-FID.
13
1
NMR signals of the fatty acid moiety ( C/ H): δ 202.87/9.79 (FA-9,
1H, t), 173.22 (FA-1), 43.89/2.44 (FA-8, 2H, d of t), 34.63/2.29 (FA-2,
2H, t), 29.02/1.35 (FA-4/5/6, 2H, m), 28.97/1.35 (FA-4/5/6, 2H, m),
1
B
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
2
8.89/1.35 (FA-4/5/6, 2H, m), 27.82/1.61 (FA-7, 2H, m), 24.95/1.63
were ≥82% for 8-hydroxyoctanoates and ≥80% for 7-hydroxyheptan-
13
1
(FA-3, 2H, m).
oate. NMR signals of the fatty acid moieties ( C/ H): sitosteryl
7-hydroxyheptanoate, δ 173.25 (FA-1), 62.94/3.67 (FA-7, 2H, t),
34.6/2.32 (FA-2, 2H, t), 32.55/1.6 (FA-6, 2H, m), 28.9/1.39 (FA-4/5,
2H, m), 28.86/1.31 (FA-4/5, 2H, m), 25.4/1.66 (FA-3, 2H, m);
sitosteryl 8-hydroxyoctanoate, δ 173.29 (FA-1), 63.06/3.66 (FA-8, 2H, t),
34.64/2.29 (FA-2, 2H, t), 32.71/1.58 (FA-7, 2H, m), 29.21/1.37
(FA-4/5, 2H, m), 29.02/1.29 (FA-4/5, 2H, m), 25.66/1.37 (FA-6,
2H, m), 24.95/1.66 (FA-3, 2H, m).
Synthesis of 8-Oxooctanoic Acid and 7-Oxoheptanoic Acid.
8
-Oxooctanoic acid and 7-oxoheptanoic acid were synthesized on the
24
basis of a previously published method via Baeyer−Villiger reaction
of the corresponding cycloalkanones, lactone opening, and subsequent
oxidation of the hydroxyacids. Cyclooctanone and cyclohepta-
none (2.079 and 1.392 g, respectively) were dissolved in 10 mL of
dichloromethane, and m-CPBA (2.152 and 1.392 g, respectively) was
added. After 6 days of stirring at room temperature, the reaction was
quenched by the addition of 500 μL of saturated sodium thiosulfate
solution. The organic layer was washed with 3 × 20 mL saturated
sodium hydrogen carbonate solution and 2 × 20 mL of an aqueous
sodium chloride solution (10%), dried over magnesium sulfate, and
filtered. After evaporation of the solvent under nitrogen, the following
colorless liquids (purities determined by GC) were obtained: 2-oxo-
nanone (723 mg; 20% lactone and 80% educt) and 2-oxocanone
Synthesis of Stigmasta-3,5-dien-7-one. “β-Sitosterol” (740 mg)
was acetylated with 3 mL of acetic acid anhydride in 30 mL of pyridine
at room temperature for 12 h. The solvent was removed under
reduced pressure, and sitosteryl acetate was used for the synthesis of
2
6
7-ketositosteryl acetate as described by Geoffroy et al. Briefly, 7.4 g
of pyridinium chlorochromate was added to a suspension of 18 g of
Celite in 90 mL of benzene, followed by the addition of sitosteryl
acetate. The reaction mixture was refluxed for 24 h. After cooling to
room temperature, the reaction mixture was filtered on a fritted glass
funnel, and the filtrate was carefully washed with 3 × 15 mL of ethyl
acetate. After removal of the solvents under reduced pressure,
sitosteryl acetate (623 mg) was removed from the reaction mixture
on a silica gel column (30 g) with ethyl acetate/hexane (5:95, v/v),
followed by the elution of 7-ketositosteryl acetate (90 mg) with ethyl
acetate/hexane (10:90, v/v). Then, an aliquot of 23 mg of
7-ketositosteryl acetate and 52 mg of p-toluenesulfonic acid, dissolved
in 5 mL of toluene, was refluxed for 2 h according to the method of
(
563 mg; 21% lactone and 79% educt). For lactone opening, the
reaction products were dissolved in 2 mL of dioxane, treated with
5 mL of sodium hydroxide (3 M), and stirred overnight at room
1
temperature. The mixtures were washed with 15 mL of ethyl acetate,
and the pH was adjusted to 3.5 with hydrochloric acid (25%). The
mixtures were extracted with 2 × 20 mL of ethyl acetate, washed with
2
× 20 mL of saturated sodium hydrogen carbonate solution, dried
over magnesium sulfate, filtered, and evaporated under nitrogen
stream. 8-Hydroxyoctanoic acid (168 mg) and 7-hydroxyheptanoic
acid (130 mg) were obtained as white solids. After silylation, GC
purities of ≥90% were determined for both acids. 8-Hydroxyoctanoic
acid and 7-hydroxyheptanoic acid were dissolved in dimethyl sulfoxide
2
7
Abramovitch and Micetich The mixture was dissolved in diethyl
ether and washed with an aqueous sodium carbonate solution and
distilled water. After evaporation to dryness, 20 mg of stigmasta-
3,5-dien-7-one was obtained as a yellow oil (purity determined by
GC = 75%).
(3.7 and 3.2 mL, respectively), before 2-iodoxybenzoic acid (463 and
4
40 mg, respectively) was added. The mixtures were stirred at room
temperature for 4 h, the reactions were quenched with distilled water,
and the mixtures were filtered. After extraction with 2 × 20 mL of
ethyl acetate, drying over magnesium sulfate, filtering, and evaporation
under nitrogen stream, 165 mg of 8-oxooctanoic acid and 125 mg of
Thermal Treatment of the Mixture of Phytosteryl/-stanyl
Linoleates. The mixture of phytosteryl/-stanyl linoleates was weighed
into a 2 mL brown glass vial without a lid (12 ± 0.2 mg) and oxidized
in a heating block (VLM Metal, Bielefeld, Germany) at 180 °C for
40 min. After thermal treatment, the samples were cooled to room
temperature before undergoing further sample preparation.
Solid-Phase Extraction. The heated linoleates were dissolved in
3 mL of cyclohexane, and the solution was loaded onto a Supelclean
500 mg silica SPE column (Supelco), which had been preconditioned
with 6 mL of hexane. The nonoxidized linoleates and nonpolar
oxidation products were eluted from the SPE column, attached to a
vacuum chamber, with 21 mL of cyclohexane at a flow rate of approx-
imately 1 mL/min (fraction 1). Subsequently, the polar oxidation
products were eluted with 6 mL of MTBE (fraction 2). The solvents of
both fractions were evaporated under a gentle nitrogen stream. The
residue of fraction 1 was dissolved in 1 mL of hexane/MTBE (3:2,
v/v) and subjected to GC-FID/GC-MS analysis. The residue of
fraction 2 was silylated with 100 μL of pyridine and 100 μL of BSTFA/
TMCS for 20 min at 80 °C. The silylation reagent was evaporated
under a gentle nitrogen stream, and the residue was dissolved in
500 μL of hexane/MTBE (3:2, v/v) and subjected to GC-MS/FID
analysis.
7-oxoheptanoic acid were obtained as white solids.
Synthesis of Phytosteryl/-stanyl 8-Oxooctanoates and
-Oxoheptanoates. Esterifications of the phytosterol/-stanyl mix-
7
ture “β-sitosterol” with 8-oxooctanoic acid and 7-oxoheptanoic acid
were performed as described for the 9-oxononanoates. After esterifica-
tion, the esters were purified by silica SPE as described for the
9-oxononanoates, but the esters were eluted with MTBE. After
purification, 43.5 mg of a mixture of phytosteryl/-stanyl 8-oxoocta-
noates and 26 mg of a mixture of phytosteryl/-stanyl 7-oxoheptanoates
were obtained as colorless oils. The phytosterol/-stanol distribution of
the mixture corresponded to that of the “β-sitosterol” sample used for
1
esterification. The purities of the oxoesters determined by H NMR
and GC-FID were ≥80% for 8-oxooctanoates and ≥70% for 7-oxo-
13
1
heptanoates. NMR signals of the fatty acid moieties ( C/ H):
sitosteryl 7-oxoheptanoate, δ 202.59/9.79 (FA-7, 1H, t), 173 (FA-1),
4
(
3.68/2.46 (FA-6, 2H, d of t), 34.57/2.31 (FA-2, 2H, t), 28.57/1.28
FA-4/5, 2H, m), 27.81/1.59 (FA-4/5, 2H, m), 24.73/1.62 (FA-3, 2H,
m); sitosteryl 8-oxooctanoate, δ 203/9.78 (FA-8, 1H, t), 173 (FA-1),
3.82/2.45 (FA-7, 2H, d of t), 34.56/2.29 (FA-2, 2H, t), 28.82/1.36
FA-4/5, 2H, m), 28.8/1.36 (FA-4/5, 2H, m), 27.82/1.66 (FA-6, 2H,
4
(
GC-FID Analysis. The analysis of the SPE fractions (1 μL injection
volume) was performed using a 6890N GC equipped with an FID
(Agilent Technologies, Boblingen, Germany). The separations were
̈
m), 24.8/1.65 (FA-3, 2H, m).
Reduction of Oxoesters to the Corresponding Hydroxyest-
carried out on a 30 m × 0.25 mm i.d., 0.1 μm film, Rtx-200MS fused
silica capillary column (Restek, Bad Homburg, Germany). The tem-
perature of the injector was set to 280 °C, and hydrogen was used as
carrier gas with a constant flow rate of 1.5 mL/min. The split flow
was set to 11 mL/min, resulting in a split ratio of 1:7.5. The oven
temperature was programmed as follows: initial temperature, 100 °C;
programmed at 15 °C/min to 310 °C (2 min) and then at 1.5 °C/min
to 340 °C (3 min). The FID temperature was set to 340 °C, and
nitrogen was used as makeup gas with a flow rate of 25 mL/min. Data
acquisition was performed by ChemStation B.04.03.
ers. The oxoesters were reduced to the hydroxyesters according to
2
5
a procedure described for the reduction of 9-oxononanoic acid. The
oxoesters (5−10 mg) were dissolved in 200 μL of dioxane and cooled
in an ice bath. Sodium borohydride (5 mg) was added to the solution
and stirred for 10 min at room temperature. Crushed ice was added to
quench the reaction, and the hydroxides were extracted with 2 × 3 mL
hexane/MTBE (3:2, v/v). After drying over magnesium sulfate and
solvent evaporation under a nitrogen stream, 8.4 mg of a mixture of
phytosteryl/-stanyl 8-hydroxyoctanoates was obtained as a white solid
and 4.5 mg of a mixture of 7-hydroxyheptanoates as a colorless oil.
The phytosterol/-stanol distributions of the mixtures corresponded
to those of the oxoesters used for reduction. The purities of the
GC-MS Analysis. Mass spectra were recorded with a Finnigan
Trace GC ultra coupled with a Finnigan Trace DSQ mass spectrom-
eter (Thermo Electro Corp., Austin, TX, USA). Mass spectra were
obtained by positive electron-impact ionization at 70 eV in the scan
1
hydroxyesters determined by H NMR and GC-FID (after silylation)
C
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
Figure 2. Capillary gas chromatographic separations of (A) the unpolar and (B) the polar fractions obtained via SPE from the heated phytosteryl/-
stanyl linoleate mixture. 1, sitosteryl heptanoate; 2, sitostanyl heptanoate; 3, campesteryl octanoate; 4, sitosteryl octanoate; 5, sitostanyl octanoate; 6,
campesteryl linoleate; 7, campestanyl linoleate; 8, sitosteryl linoleate; 9, sitostanyl linoleate; 10, linoleic acid; 11, sitosterol; 12, stigmasta-3,5-dien-7-
one; 13, sitosteryl 7-hydroxyheptanoate; 14, campesteryl 8-hydroxyoctanoate; 15, sitosteryl 8-hydroxyoctanoate; 16, sitostanyl 8-hydroxyoctanoate;
1
7, sitosteryl 7-oxoheptanoate; 18, sitosteryl 8-oxooctanoate; 19, sitostanyl 8-oxooctanoate; 20, campesteryl 9-oxononanoate; 21, sitosteryl
9-oxononanoate; 22, sitostanyl 9-oxononanoate; 23/24, α/β-7-hydroxysitosteryl linoleate (tentative); 25, α/β-5,6-epoxysitosteryl linoleate (tentative).
Table 1. Characteristic GC-MS Fragment Ions of Steryl/Stanyl Ester Oxidation Products
a
b
molecular ion [M]+
acyl chain oxidation product
RT (min)
characteristic fragments [m/z]
1
2
3
4
5
1
1
1
1
1
1
1
2
2
2
, sitosteryl heptanoate
, sitostanyl heptanoate
, campesteryl octanoate
, sitosteryl octanoate
, sitostanyl octanoate
14.565
14.636
14.662
14.972
15.049
16.606
16.740
17.277
17.398
17.736
18.590
18.761
18.856
19.559
19.722
526 (−)
528 (6)
526 (−)
540 (−)
542 (4)
614 (−)
614 (−)
628 (−)
630 (0.2)
540 (−)
554 (−)
556 (0.9)
554 (−)
568 (−)
570 (1.3)
397 (33), 396 (100), 381 (27), 275 (15), 255 (22), 213 (16)
513 (0.3), 399 (10), 398 (41), 383 (35), 344 (10), 257 (18), 215 (100)
383 (36), 382 (100), 367 (33), 261 (22), 255 (26), 213 (20)
397 (49), 396 (100), 381 (46), 275 (34), 255 (40), 213 (34)
527 (1), 399 (21), 398 (53), 383 (39), 344 (11), 257 (18), 215 (100)
397 (47), 396 (100), 381 (24), 275 (14), 255 (17), 213 (14), 524 (0.3)
383 (39), 382 (100), 367 (23), 261 (13), 255 (15), 213 (26)
397 (43), 396 (100), 381 (18), 275 (12), 255 (17), 213 (14), 538 (0.2)
399 (17), 398 (42), 383 (32), 344 (5), 257 (17), 215 (100), 540 (0.3)
525 (0.1), 397 (35), 396 (100), 381 (28), 275 (14), 255 (20), 213(19)
397 (50), 396 (100), 381 (41) 275 (20), 255 (24), 213 (18)
541 (0.7), 399 (14), 398 (50), 383 (36), 344 (4), 257 (14), 215 (100)
383 (30), 382 (100), 367 (27), 261 (12), 255 (14), 213 (13),
397 (32), 396 (100), 381 (25), 275 (13), 255 (15), 213 (12),
c
3, sitosteryl 7-hydroxyheptanoate
c
4, campesteryl 8-hydroxyoctanoate
c
5, sitosteryl 8-hydroxyoctanoate
c
6, sitostanyl 8-hydroxyoctanoate
7, sitosteryl 7-oxoheptanoate
8, sitosteryl 8-oxooctanoate
9, sitostanyl 8-oxooctanoate
0, campesteryl 9-oxononanoate
1, sitosteryl 9-oxononanoate
2, sitostanyl 9-oxononanoate
555 (0.3), 399 (21), 398 (58), 383 (36), 344 (7), 257 (16), 215 (100)
a
b
c
Numbers correspond to those in the chromatogram shown in Figure 2. Retention times on Rtx-200MS. As trimethylsilyl ether.
mode at unit resolution from 40 to 750 Da. The interface was heated
to 330 °C and the ion source to 250 °C. Helium was used as carrier
gas with a constant flow rate of 1.0 mL/min. The other GC conditions
were the same as described for GC-FID analysis. Data acquisition was
perfomed by Xcalibur 3.063 software.
13C NMR spectra were recorded in proton-decoupled mode. Data
processing was typically done with the MestreNova software.
RESULTS AND DISCUSSION
■
Thermal Treatment and Sample Preparation. Phytos-
teryl/-stanyl linoleates prepared via esterification of a commer-
cially available mixture of sterols (75% β-sitosterol, 10% cam-
pesterol) and stanols (12% sitostanol, 2% campestanol) were
used as model substrates for the thermal treatment. Such esters
account for a large proportion of the phytosteryl/-stanyl fatty
NMR Spectroscopy. The compounds were dissolved in 0.5 mL of
1
13
deuterated chloroform. H NMR and C NMR spectra were recorded
at 500 and 126 MHz, respectively, with Avance-HD 500 spectrometers
1
(
Bruker, Billerica, MA, USA) operating at 27 °C. H-detected experi-
ments including two-dimensional COSY, NOESY, HSQC, and HMBC
1
13
13
were measured with an inverse H/ C probe head; direct C mea-
10
surements were performed with a QNP 13C/ P/ Si/ F/ H cryo-
probe. All experiments were done in full automation using standard
parameter sets of the TOPSPIN 3.2 software package (Bruker).
31
29
19
1
acid ester mixtures being added to foods.
Nonoxidized phytosteryl/-stanyl linoleates were anticipated to
be still quantitatively dominating in the thermally treated sample.
D
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
+
Figure 3. Electron-impact ionization spectra of (A) sitosteryl 9-oxononanoate and (B) sitostanyl 9-oxononanoate. [M] , molecular ion; [FA], fatty
acid; [SC], sterol/stanol side chain.
Therefore, as a first step, an SPE method was developed
to preconcentrate the acyl chain oxidation products by removal
of the nonoxidized esters. The oxidized esters were expected
to be more polar than the starting esters. Accordingly, the
strategy was based on an elution of the nonoxidized esters from
a silica SPE column with a nonpolar eluent in a first fraction,
followed by a second eluting step for the polar oxidation
products. In previous studies, SPE methods have also been
described for the preconcentration of cholesteryl ester
hydroperoxides formed upon heating of cholesteryl esters and
for the separation of oxidized fatty acid methyl esters into
elution powers were tested to find an eluent just capable of
eluting the nonoxidized linoleates from the silica column
without causing the elution of polar oxidation products. The
compounds retained on the column were subsequently eluted
with MTBE, and both fractions were checked for nonoxidized
linoleates by GC-FID/MS. When using heptane or hexane as
elution solvents for the first fraction, remainders of nonoxidized
esters were still detectable in the MTBE fraction. Using cyclo-
hexane, nonoxidized linoleates could be effectively removed
from the silica column in the first elution step (Figure 2A), and
no nonoxidized linoleates were detected in the second fraction
after elution with MTBE (Figure 2B).
17,19,28
nonpolar and polar fractions.
In the course of the method
development several nonpolar solvents with increasing
E
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
1
Figure 4. NMR spectra of synthesized sitosteryl 9-oxononanoate. (A) H NMR; characteristic proton signals of the sterol moiety and the fatty acid
1
3
moiety: δ 9.79 (FA-9, 1H, t), 5.39 (St-6, 1H, m), 4.63 (St-3, 1H, m), 2.29 (FA-2, 2H, t). The asterisks represents an impurity. (B) C NMR;
characteristic carbon signals of the sterol moiety and the fatty acid moiety: δ 202.87 (FA-9), 173.22 (FA-1), 139.7 (St-5), 122.59 (St-6), 73.67 (St-3).
Identification of Oxidation Products. The capillary gas
chromatographic analysis of fraction 1 revealed that in addition
to the remaining nonoxidized linoleates several minor peaks
were detectable (Figure 2A). Peaks 1−5 were identified via
GC-MS as heptanoates and octanoates of sitosterol, campester-
ol, and sitostanol (Table 1).
reported as a major oxidation product resulting from the
30,31
thermal treatment of sitosteryl esters.
The respective
7-ketositosteryl linoleate was not detected in the present
study; stigmasta-3,5-dien-7-one might be explained, for
example, as degradation product formed from 7-ketositosteryl
linoleate via liberation of linoleic acid. On the other hand, the
analogous oxidation product stigmasta-3,5,22-trien-7-one has
also been identified upon thermo-oxidation of free stigmaster-
In fraction 2, three major peaks were detected (Figure 2B).
Peaks 10 and 11 were identified as trimethylsilyl derivatives of
linoleic acid and β-sitosterol, respectively. According to GC and
32
ol, indicating that compound 12 might also be the deg-
radation product of intermediately formed β-sitosterol.
1
H NMR analysis, both were not contained as impurities in the
In addition to these three quantitatively dominating com-
pounds, GC-MS analysis of fraction 2 revealed the presence of
a series of minor peaks exhibiting fragmentation patterns
synthesized mixture of phytosteryl/-stanyl linoleates used for
the heating experiment. Their occurrence might be explained
by a potential formation via hydrolysis during the heating
procedure. On the other hand, the presence of linoleic acid may
also be related to the formation of compound 12. On the basis
of comparison of the retention time and the mass spectrum to
those of a synthesized reference compound, peak 12 was assigned
as stigmasta-3,5-dien-7-one. This compound has previously been
similar to those described for nonoxidized phytosteryl/-stanyl
21,33,34
esters.
The respective part of the chromatogram is
enlarged in Figure 2B, and the MS data are listed in Table 1.
The base peaks m/z 396 observed for compounds 13, 15, 17,
1
8, and 21 and m/z 382 for compounds 14 and 20 correspond
to those described to result from the cleavage of the fatty acid
from sitosteryl and campesteryl esters, respectively.
27,29
33,34
identified in heartwood and in the smoke of incense.
In
The
previous investigations involving the cleavage of the ester bond
as part of the analytical procedure, 7-ketositosterol has been
base peak m/z 215 observed for compounds 16, 19, and 22 has
been described as being generated by cleavage of the fatty acid
F
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
Figure 5. Classes of identified oxidation products in the fatty acid part upon heating of sitosteryl, campesteryl, and sitostanyl esters, exemplarily
shown for sitosteryl linoleate. Class I (alkanoates): 1, sitosteryl heptanoate; 4, sitosteryl octanoate. Class II (hydroxyalkanoates): 13, sitosteryl
7-hydroxyheptanoate; 15, sitosteryl 8-hydroxyoctanoate. Class III (oxoalkanoates): 17, sitosteryl 7-oxoheptanoate; 18, sitosteryl 8-oxooctanoate; 21,
sitosteryl 9-oxononanoate. The numbering corresponds to that of Figure 2.
and further fragmentation involving the sterol side chain of
phytosteryl/-stanyl mixture as used for the esterification of lin-
oleic acid. The corresponding hydroxyalkanoates were obtained
by reduction of 7-oxoheptanoates and 8-oxooctanoates with
sodium borohydride.
21
short-chain sitostanyl esters. This enabled the differentiation
of steryl and stanyl esters; exemplary mass spectra are shown in
Figure 3.
21
In a previous study in which the same mass spectrometric
conditions were used, weak but characteristic molecular ions
could be detected for sitostanyl fatty acid esters. Therefore, on
the basis of the oxidative modifications in the acyl part known
The identities of the synthesized reference compounds were
confirmed via NMR. With a proportion of 75%, sitosterol was
the major phytosterol in the synthesized phytosteryl/-stanyl
1
13
ester mixtures; therefore, H/ C NMR signals of sitosteryl
esters could be interpreted. By making use of a repertoire of
two-dimensional techniques, it was possible to assign almost
every carbon and proton signal to their respective atom number
14
from peroxidation of cholesteryl linoleate and high-temper-
ature degradation of methyl linoleate and triacylglycer-
1
8,35,36
ides,
molecular masses of potential acyl chain oxidation
13
products of sitostanyl linoleate, that is, oxo- and hydroxyalka-
noates, were calculated and compared with the mass spectra
determined in fraction 2. This screening gave first indica-
tions that the peaks 16, 19, and 22 corresponded to sitostanyl
(Figure 1). By comparison of the C spectrum of unesterified
13
sitosterol with C spectra of the sitosteryl ester reference com-
pounds, the carbon signals of the acyl chains could be assigned.
Each synthesized compound exhibited the same proton and
carbon signals in the sterol moiety, thus only differing in their
signals in the fatty acid part. Equally characteristic for each
sitosteryl ester was the carbon atom FA-1 involved in the ester
bond, showing a chemical shift of 173 ppm, and the carbon
atom FA-2, with a chemical shift of 34.6 ppm and a well-
resolved triplet proton signal pattern at 2.3 ppm. The signals of
terminal carbon and proton atoms of the fatty acid part enabled
discrimination between the synthesized reference classes. The
terminal carbon atom of sitosteryl alkanoates exhibited a signal
at 14.1 ppm, and the corresponding proton showed a triplet
proton signal pattern at 0.9 ppm, whereas the terminal alde-
hyde function of core aldehyde esters had a chemical shift at
8
-hydroxyoctanote, 8-oxooctanoate, and 9-oxononanoate,
respectively.
These preliminary assignments initiated the next phase of the
study, namely, the synthesis and characterization of authentic
reference compounds. First, potential candidates of oxidized
fatty acid moieties were synthesized via established synthetic
routes: 9-oxononanoic acid was obtained by oxidative diol
22
cleavage of aleuritic acid; 8-oxooctanoic acid and 7-oxo-
heptanoic acid were obtained by Baeyer−Villiger reaction of the
corresponding cycloalkanones and subsequent oxidation with
24
iodoxybenzoic acid. These synthesized acids as well as hep-
tanoic and octanoic acid were esterified with the same
G
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
Figure 6. Formation pathways of steryl/stanyl ester oxidation products in the acyl chain by homolytic β-scission of primarily formed allylic
3
7
hydroperoxides on positions (A) 8 and (B) 9. Adapted from Frankel. Numbering corresponds to that in Figures 2 and 5. Only pathways leading to
the compounds identified in this study are illustrated.
2
03 ppm and an intense triplet pattern was observed for the
hydroperoxides on positions 8 and 9, respectively, are degraded
by homolytic β-scission according to route I (Figure 6). After
interaction of the resulting alkyl radical with a hydrogen radical,
heptanoates and octanoates, respectively, are obtained. Alter-
natively, 7-hydroxyheptanoates and 8-hydroxyoctanoates are
formed by reaction of the alkyl radical with a hydroxyl radical.
Repeated oxidation of the alkyl radical formed from the
degradation of the 8-hydroperoxide with oxygen leads to the
formation of 7-hydroperoxides, which can decompose to
7-oxoheptanoates via an alkoxyl radical. On the other hand,
8-oxooctanoates and 9-oxononanoates emerge directly upon
β-scission of allylic hydroperoxides on positions 8 and 9 fol-
lowing route II. In addition, 9-oxononanoates may be formed
from the allylic hydroperoxide in position 10 following route I
and subsequent reaction with a hydroxyl radical.
aldehyde proton at 9.8 ppm (Figure 4). The acquired signal pat-
tern and peak assignments correlated with published H and C
1
13
23
data of cholesteryl 9-oxononanoate. The terminal carbon atom
of sitosteryl hydroxyalkanoates showed a signal at 63 ppm and a
well-resolved triplet pattern at 3.66 ppm of the terminal protons.
Final peak identifications were carried out by comparing
retention times and mass spectra of sample peaks with those of
the synthesized reference compounds (Table 1). Due to their
slightly higher polarity compared to the respective linoleates,
sitosteryl octanoate 4 and sitostanyl octanoate 5 were also
present in fraction 2. In total, seven types of oxidation products
of the fatty acid moiety belonging to three different classes, that
is, alkanoates, hydroxyalkanoates, and oxoalkanoates, could be
identified upon heating of phytosteryl/phytostanyl linoleates
(
Figure 5). These classes of acyl chain oxidation products were
In addition to the described acyl chain oxidation prod-
ucts, three representatives of linoleates of oxidized sterols
were detected in fraction 2. Peaks 23 and 24 (Figure 2B) were
assumed to be silylated α/β-7-hydroxysitosteryl linoleates,
because their mass spectra (Table 2) exhibited a characteristic
in line with those previously reported for cholesteryl esters,
fatty acid methyl esters, and triacylglycerides.
14,17,18
The formation of these acyl chain oxidation products may be
explained analogously to the routes described for the oxidation
of fatty acid methyl esters.
18,37,38
+
The primarily formed allylic
base peak of m/z 484 corresponding to [M − FA] , which was
H
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
Table 2. Characteristic GC-MS Fragment Ions of Tentatively
Assigned Sitosteryl Linoleates Oxidized in the Sterol Moiety
acid monohydrate; DCC,, dicyclohexylcarbodiimide; DMAP,,
4-(dimethylamino)pyridine; MTBE, methyl tert-butyl ether
b
sitosteryl linoleate
RT
molecular
ion [M]+
characteristic fragments
a
oxidation product
(min)
[m/z]
REFERENCES
■
2
2
3, 7α/β-hydroxy
20.955
765 (−)
486 (12), 485 (55),
484 (100), 483 (33)
(1) Commission Decision of 24 July 2000 on authorising the placing
on the market of “yellow fat spreads with added phytosterol esters” as
a novel food or novel food ingredient under regulation (EC) No. 258/
97 of the European Parliament and of the Council. Off. J. Eur.
Communities 2000, L200, 59−60.
,
cd
sitosteryl linoleate
4, 7α/β-
23.038
765 (−)
486 (16), 485 (46),
484 (100), 483 (56)
hydroxysitosteryl
,
cd
linoleate
2
5, 5,6α/β-
25.845
693 (−)
413 (44), 395 (100)
(
2) Demonty, I.; Ras, R. T.; van der Knaap, H. C. M.; Duchateau, G.
epoxysitosteryl
d
linoleate
S. M. J. E.; Meijer, L.; Zock, P. L.; Geleijnse, J. M.; Trautwein, E. A.
Continuous dose-response relationship of the LDL-cholesterol-low-
ering effect of phytosterol intake. J. Nutr. 2009, 139, 271−284.
(3) AbuMweis, S. S.; Barake, R.; Jones, P. J. H. Plant sterols/stanols
as cholesterol lowering agents: a meta-analysis of randomized
controlled trials. Food Nutr. Res. 2008, 52, 1811.
a
Numbers correspond to those in the chromatogram shown in
b
c
Figure 2B. Retention times on Rtx-200MS. As trimethylsilyl ether.
d
α/β-Stereoisomers could not be distinguished.
also described as the base peak of double-silylated 7-hydroxy-
(4) Moreau, R. A.; Whitaker, B. D.; Hicks, K. B. Phytosterols,
phytostanols, and their conjugates in foods: structural diversity,
quantitative analysis, and health-promoting uses. Prog. Lipid Res. 2002,
39
sitosterols in a previous study. Peak 25 was tentatively assigned
as α- or β-5,6-epoxysitosteryl linoleate, exhibiting characteristic
fragment ions similar to those reported for silylated α/β-5,6-
epoxysitosterols, with m/z 395 potentially corresponding to
4
(
1, 457−500.
5) SCF (Scientific Committee on Food). Opinion of the scientific
+
+
40
commitee on food on a request for the safety assessment of the use of
phytosterol esters in yellow fat spreads; http://ec.europa.eu/food/fs/
sc/scf/out56_en.pdf (accessed Sept 14, 2016).
[
M − FA − 18] and m/z 413 resulting from [M − FA] .
As no reference compounds were available for these esters,
their identifications are considered only tentative.
(6) Garcia-Llatas, G.; Rodriguez-Estrada, M. T. Current and new
There are other representatives of potential acyl chain oxi-
dation products that have not been detected in this study, for
example, those deriving from the 13-hydroperoxides or dihy-
droxyacids, comparable to the previously reported 9,10-dihy-
insights on phytosterol oxides in plant sterol-enriched food. Chem.
Phys. Lipids 2011, 164, 607−624.
(7) Otaegui-Arrazola, A.; Menen
Astiasaran
48, 3289−3303.
8) Scholz, B.; Guth, S.; Engel, K.-H.; Steinberg, P. Phytosterol
́
̃
dez-Carreno, M.; Ansorena, D.;
, I. Oxysterols: a world to explore. Food Chem. Toxicol. 2010,
́
20
droxystearates of sitosterol or of 7-ketositosterol. It remains
to be investigated whether they are not covered by the analytical
procedure or whether they are not formed under the employed
conditions.
(
oxidation products in enriched foods: occurrence, exposure, and
biological effects. Mol. Nutr. Food Res. 2015, 59, 1339−1352.
(9) Lin, Y.; Knol, D.; Menendez-Carreno, M.; Blom, W. A. M.;
In conclusion, the employed methodology, that is, a precon-
centration of polar oxidation products via SPE, the establish-
ment of capillary gas chromatographic conditions enabling the
separation of various oxidation products, and the synthesis of
authentic reference compounds, provided data on the for-
mation and identities of previously unreported acyl chain
oxidation products resulting from the thermal treatment of
phytosteryl/-stanyl fatty acid esters. The observed classes of
products correspond to those expected from the thermo-
oxidation of cholesteryl fatty acid esters, unsaturated fatty acids,
and triglycerides. However, the fact that the approach is not
based on the formation of methyl esters via transesterification
enabled for the first time the analysis of a spectrum of indi-
vidual phytosteryl/-stanyl fatty acid esters oxidized in the acyl
moiety. This will be a crucial cornerstone of subsequent quan-
titative studies on the contribution of these oxidation products
to the decreases of phytosteryl/-stanyl fatty acid esters upon
thermal treatment.
Matthee, J.; Janssen, H.-G.; Trautwein, E. A. Formation of plant sterol
oxidation products in foods during baking and cooking using
margarine without and with added plant sterol esters. J. Agric. Food
Chem. 2016, 64, 653−662.
(10) Scholz, B.; Menzel, N.; Lander, V.; Engel, K.-H. Heating two
types of enriched margarine: complementary analysis of phytosteryl/
phytostanyl fatty acid esters and phytosterol/phytostanol oxidation
products. J. Agric. Food Chem. 2016, 64, 2699−2708.
(
11) Leitinger, N. Cholesteryl ester oxidation products in
atherosclerosis. Mol. Aspects Med. 2003, 24, 239−250.
12) Miyoshi, N.; Iwasaki, N.; Tomono, S.; Higashi, T.; Ohshima, H.
(
Occurrence of cytotoxic 9-oxononanoyl secosterol aldehydes in human
low-density lipoprotein. Free Radical Biol. Med. 2013, 60, 73−79.
(13) Kamido, H.; Kuksis, A.; Marai, L.; Myher, J. J. Lipid ester-bound
aldehydes among copper-catalyzed peroxidation products of human
plasma lipoproteins. J. Lipid Res. 1995, 36, 1876−1886.
(14) Kamido, H.; Kuksis, A.; Marai, L.; Myher, J. J. Identification of
core aldehydes among in vitro peroxidation products of cholesteryl
esters. Lipids 1993, 28, 331−336.
(15) Kamido, H.; Kuksis, A.; Marai, L.; Myher, J. J. Identification of
AUTHOR INFORMATION
Corresponding Author
(K.-H.E.) Phone: +49 (0)8161 71 4250. Fax: +49 (0)8161 71
259. E-mail: k.h.engel@wzw.tum.de.
cholesterol-bound aldehydes in copper-oxidized low density lip-
■
oprotein. FEBS Lett. 1992, 304, 269−272.
(16) Velasco, J.; Marmesat, S.; Marquez-Ruiz, G.; Dobarganes, M. C.
*
Formation of short-chain glycerol-bound oxidation products and
oxidized monomeric triacylglycerols during deep-frying and occur-
rence in used frying fats. Eur. J. Lipid Sci. Technol. 2004, 106, 728−735.
4
ORCID
Karl-Heinz Engel: 0000-0001-8932-5175
Notes
The authors declare no competing financial interest.
́
(17) Dobarganes, M. C.; Marquez-Ruiz, G. 6 − Formation and
analysis of oxidized monomeric, dimeric, and higher oligomeric
triglycerides A2. In Deep Frying, 2nd ed.; Erickson, M. D., Ed.; AOCS
Press: 2007; pp 87−110.
(
́ ́ ́ ́
18) Berdeaux, O.; Fontagne, S.; Semon, E.; Velasco, J.; Sebedio, J.
ABBREVIATIONS USED
POP, phytosterol oxidation products; SPE, solid-phase extraction;
m-CPBA, 3-chloroperbenzoic acid; 4-TsOH, 4-toluenesulfonic
■
L.; Dobarganes, C. A detailed identification study on high-temperature
degradation products of oleic and linoleic acid methyl esters by GC−
MS and GC−FTIR. Chem. Phys. Lipids 2012, 165, 338−347.
I
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Journal of Agricultural and Food Chemistry
Article
(
19) Lehtonen, M.; Lampi, A.-M.; Ollilainen, V.; Struijs, K.; Piironen,
(40) Zhang, X.; Julien-David, D.; Miesch, M.; Geoffroy, P.; Raul, F.;
V. The role of acyl moiety in the formation and reactions of steryl ester
́
Roussi, S.; Aoude-Werner, D.; Marchioni, E. Identification and
quantitative analysis of β-sitosterol oxides in vegetable oils by capillary
gas chromatography−mass spectrometry. Steroids 2005, 70, 896−906.
hydroperoxides. Eur. Food Res. Technol. 2011, 233, 51−61.
(
20) Julien-David, D.; Zhao, M.; Geoffroy, P.; Miesch, M.; Raul, F.;
Aoude-Werner, D.; Ennahar, S.; Marchioni, E. Analysis of sitosteryl
oleate esters in phytosterols esters enriched foods by HPLC-ESI-MS2.
Steroids 2014, 84, 84−91.
(
21) Barnsteiner, A.; Esche, R.; di Gianvito, A.; Chiavaro, E.; Schmid,
W.; Engel, K.-H. Capillary gas chromatographic analysis of complex
phytosteryl/-stanyl ester mixtures in enriched skimmed milk-drinking
yoghurts. Food Control 2012, 27, 275−283.
(
22) Reuter, J. M.; Salomon, R. G. Aleuritic acid, an abundant source
of prostanoid synthons. J. Org. Chem. 1978, 43, 4247−4248.
23) Boechzelt, H.; Karten, B.; Abuja, P. M.; Sattler, W.; Mittelbach,
M. Synthesis of 9-oxononanoyl cholesterol by ozonization. J. Lipid Res.
998, 39, 1503−1507.
24) Rajabi, M.; Lanfranchi, M.; Campo, F.; Panza, L. Synthesis of a
(
1
(
series of hydroxycarboxylic acids as standards for oxidation of
nonanoic acid. Synth. Commun. 2014, 44, 1149−1154.
(
25) Kallury, R. K.; Krull, U. J.; Thompson, M. Synthesis of
phospholipids suitable for covalent binding to surfaces. J. Org. Chem.
987, 52, 5478−5480.
26) Geoffroy, P.; Julien-David, D.; Marchioni, E.; Raul, F.; Aoude-
Werner, D.; Miesch, M. Synthesis of highly pure oxyphytosterols and
oxy)phytosterol esters. Steroids 2008, 73, 702−707.
27) Abramovitch, R. A.; Micetich, R. G. Extractives from Populus
1
(
(
(
tremuloides heartwood-structure and synthesis of tremulone. Can. J.
Chem. 1962, 40, 2017−2022.
(
28) Lehtonen, M.; Lampi, A.-M.; Riuttamaki, M.-A.; Piironen, V.
Oxidation reactions of steryl esters in a saturated lipid matrix. Food
Chem. 2012, 134, 2030−2039.
(
29) Dalibalta, S.; Elsayed, Y.; Alqtaishat, F.; Gomes, I.; Fernandes,
N. A health risk assessment of Arabian incense (Bakhour) smoke in
the United Arab Emirates. Sci. Total Environ. 2015, 511, 684−691.
(
30) Soupas, L.; Huikko, L.; Lampi, A.-m.; Piironen, V. Pan-frying
may induce phytosterol oxidation. Food Chem. 2007, 101, 286−297.
(
31) Scholz, B.; Wocheslander, S.; Lander, V.; Engel, K.-H. On-line
liquid chromatography−gas chromatography: a novel approach for the
analysis of phytosterol oxidation products in enriched foods. J.
Chromatogr. A 2015, 1396, 98−108.
(
̃
́ ́
32) Menendez-Carreno, M.; Ansorena, D.; Astiasaran, I.; Piironen,
V.; Lampi, A.-M. Determination of non-polar and mid-polar
monomeric oxidation products of stigmasterol during thermo-
oxidation. Food Chem. 2010, 122, 277−284.
(
33) Esche, R.; Barnsteiner, A.; Scholz, B.; Engel, K.-H. Simultaneous
analysis of free phytosterols/phytostanols and intact phytosteryl/
phytostanyl fatty acid and phenolic acid esters in cereals. J. Agric. Food
Chem. 2012, 60, 5330−5339.
(
34) Barnsteiner, A.; Lubinus, T.; di Gianvito, A.; Schmid, W.; Engel,
K.-H. GC-based analysis of plant stanyl fatty acid esters in enriched
foods. J. Agric. Food Chem. 2011, 59, 5204−5214.
(
35) Berdeaux, O.; Marquez-Ruiz, G.; Dobarganes, C. Selection of
methylation procedures for quantitation of short-chain glycerol-bound
compounds formed during thermoxidation. J. Chromatogr. A 1999,
8
(
63, 171−181.
36) Velasco, J.; Marmesat, S.; Berdeaux, O.; Marquez-Ruiz, G.;
Dobarganes, C. Quantitation of short-chain glycerol-bound com-
pounds in thermoxidized and used frying oils. A monitoring study
during thermoxidation of olive and sunflower oils. J. Agric. Food Chem.
2
(
005, 53, 4006−4011.
37) Frankel, E. N. Lipid Oxidation, 2nd ed.; The Oily Press:
Bridgewater, UK, 2005; Vol. 18, pp 67−96.
38) Frankel, E. N.; Neff, W. E.; Selke, E. Analysis of autoxidized fats
(
by gas chromatography-mass spectrometry. IX. Homolytic vs.
heterolytic cleavage of primary and secondary oxidation products.
Lipids 1984, 19, 790−800.
(
39) Apprich, S.; Ulberth, F. Gas chromatographic properties of
common cholesterol and phytosterol oxidation products. J. Chroma-
togr. A 2004, 1055, 169−176.
J
DOI: 10.1021/acs.jafc.6b04326
J. Agric. Food Chem. XXXX, XXX, XXX−XXX