Lipid Structure Influences the Digestion and Oxidation Behavior of
Docosahexaenoic and Eicosapentaenoic Acids in the Simulated
Digestion System
Gabriele Beltrame, Eija Ahonen, Annelie Damerau, Haraldur G. Gudmundsson,
Gudmundur G. Haraldsson, and Kaisa M. Linderborg*
Cite This: J. Agric. Food Chem. 2023, 71, 10087−10096 Read Online
ACCESS Metrics & More Article Recommendations *sı Supporting Information
ABSTRACT: Omega-3 fatty acids such as eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) are essential for human
health but prone to oxidation. While esterification location is known to influence the stability of omega-3 in triacylglycerols (TAGs)
in oxidation trials, their oxidative behavior in the gastrointestinal tract is unknown. Synthesized ABA- and AAB-type TAGs
containing DHA and EPA were submitted to static in vitro digestion for the first time. Tridocosahexaenoin and DHA as ethyl esters
were similarly digested. Digesta were analyzed by gas chromatography, liquid chromatography−mass spectrometry, and nuclear
magnetic resonance spectroscopy. Besides the formation of di- and monoacylglycerols, degradation of hydroperoxides was detected
in ABA- and AAB-type TAGs, whereas oxygenated species increased in tridocosahexaenoin. Ethyl esters were mainly unaffected.
EPA was expectedly less susceptible to oxidation prior to and during the digestion process, particularly in sn-2. These results are
relevant for the production of tailored omega-3 structures to be used as supplements or ingredients.
KEYWORDS: DHA, EPA, TAG enantiomers, ethyl ester, in vitro digestion, oxidation
1. INTRODUCTION
Polyunsaturated omega-3 fatty acids (n-3 PUFAs), such as
eicosapentaenoic acid (EPA) and docosahexaenoic acid
(DHA), are essential for human health and development.
They are considered crucial in all growth stages, from
conception to aging, and are associated with visual develop-
ment, cognitive health, and protection from cardiovascular
diseases.1 As components of cell membranes, they affect cell
signal transduction, gene expression, cell growth, and apoptosis
by changing the fluidity and structure of membrane protein
domains.2 Moreover, n-3 PUFAs act as a cell regulator at the
extracellular level with derivatives such as prostaglandins,
which act as competitors in the arachidonic acid cascade
involved in inflammation.3 n-3 PUFAs are mainly produced in
the marine environment, and their main dietary source is
seafood. New sources are increasingly sought to counter the
inability of fisheries to sustainably supply the minimum
required dose of PUFAs to the world population.4
The development of products including n-3 PUFAs is
challenged by their proneness to oxidation. Light, heat, or
metal ions can catalyze the abstraction of hydrogen adjacent to
double bonds with a lower bond dissociation energy compared
to mono- or unsaturated fatty acids. The generated lipid radical
reacts with atmospheric oxygen to produce peroxyl radical,
which in turn is able to abstract another hydrogen to generate
lipid hydroperoxide and another lipid radical. Hydroperoxides
homolytically decompose to alkoxy radicals and hydroxy
radicals, which, depending on reaction conditions and lipid
structure, rearrange or decompose to a plethora of compounds,
including aldehydes.5 These compounds are responsible for the
rancidity and off-flavors, but they are also antinutritional.
Triacylglycerol (TAG) hydroperoxides might contribute to
atherosclerosis,6 while unsaturated aldehydes are considered to
be toxic and with mutagenic potential.7,8 Protection of PUFAs
from oxidation is therefore of utmost importance.
Industrially, PUFA concentrates are commonly produced by
chemical/enzymatic transesterification of the natural TAG
structure into ethyl esters (EEs) for upper concentration.
However, the higher bioavailability of TAG form compared to
EE form,9 especially from low-fat diets, makes the TAG
structure more appealing for product development. The
position of PUFAs in the glycerol backbone influences their
oxidative stability, with position sn-2 being the most
protective.10,11 Our group also observed that oxidative stability
is affected by the lipid structure (TAG or EE).12 Therefore,
one strategy for the protection of PUFAs from oxidation could
be their inclusion in the most stable chemical form, possibly in
synthesized TAGs with predetermined distribution of fatty
acids in the glycerol backbone.13 While most oxidation studies
utilized TAGs with randomized PUFA positions or with
known sn-2 but undistinguished sn-1,3 positions,14 regio- and
even enantiomerically pure TAGs (i.e., TAGs with defined
fatty acid positions) are nowadays available.15,16 Very recent
Received: April 5, 2023
Revised: June 1, 2023
Accepted: June 2, 2023
Published: June 20, 2023
Articlepubs.acs.org/JAFC
© 2023 The Authors. Published by
American Chemical Society
10087
https://doi.org/10.1021/acs.jafc.3c02207
J. Agric. Food Chem. 2023, 71, 10087−10096
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results from our group11 showed that regio- and enantiomeri-
cally pure DHA esterified in the sn-2 position was the most
resistant to oxidation, with or without antioxidants. Moreover,
as recently reviewed,17 there is evidence pointing to better
absorption of DHA by intestinal mucosa when located in
position sn-2. The same was also observed by our group in a rat
feeding trial with regio- and enantiomerically pure TAGs.18
Most PUFA oxidation studies focus on production and
storage conditions. More recently, attention has also been
given to the gastrointestinal tract, which has been characterized
to possess pro-oxidant conditions,19 due to the oxygen
incorporated during mastication, the low pH of the stomach,
and the presence of metal ions in food.20 In the gastrointestinal
tract, gastric lipase starts TAG hydrolysis in the stomach,
producing mainly DAGs. As these molecules are amphipathic,
they facilitate dispersions of lipids. In the small intestine, lipids
are emulsified by bile acids to form micelles, which facilitate
hydrolysis to 2-MAGs and free fatty acids. At physiological
conditions, micelles are absorbed by the enterocytes, in which
TAGs are resynthesized and enter lymphatic circulation.21 The
hydrolytic activity of lipase during digestion has been
correlated to oxidative degradation of oil, although the
complexity of the system has made general statements
difficult.22 Due to ethical reasons and costs, studies on lipid
oxidation during digestion generally utilize in vitro models.
Multiple models are available in the literature, with
INFOGEST 2.023 being considered the one with the widest
international consensus.
The present study aimed to investigate the oxidation
behavior of PUFAs with defined positions on the glycerol
backbone in a simulated digestion system. The main aim was
to compare structured lipids with DHA and EPA in positions
sn-1, sn-2, or sn-3. The study also aimed to compare regio- and
enantiomerically pure TAGs with pure DHA in the form of
TAG (tridocosahexaenoin) or EE, which are commonly used
as n-3 PUFA supplements.24 Oxidation behavior and digestion
products were analyzed together with undigested samples
utilizing gas chromatography coupled with flame-ionization
detection (GC-FID), liquid chromatography coupled with
mass spectrometry (LC−MS), and nuclear magnetic resonance
(NMR) spectroscopy.
2. MATERIALS AND METHODS
2.1. Chemicals, Samples, and Reagents. All chemical solvents
were supplied by Sigma-Aldrich (Saint Louis, MO, USA), unless
specified otherwise. All salts used for the simulated digestion were
supplied by VWR Chemicals (Leuven, Belgium), unless specified
otherwise. Regiopure ABA-type DHA- and EPA-TAGs (sn-2) and
regio- and enantiopure AAB-type DHA- and EPA-TAGs (sn-1, sn-3)
were synthesized (see Section 2.2). Tridocosahexaenoin (purity >
99%) and ethyl docosahexaenoate (DHA-EE, purity > 99%) were
purchased from Larodan (Solna, Sweden). These compounds, once
dissolved in hexane, were added with α-tocopherol (0.05%
tocopherol/DHA ratio) from Sigma-Aldrich (Buchs, Switzerland)
and stored at −80 °C until experiments were performed. For
simulated digestion, rabbit gastric lipase (RGE15-500) was obtained
from Lipolytic (Marseille, France), while amylase, porcine pepsin,
porcine pancreatin, and bile salts were purchased from Sigma-Aldrich
(Saint Luis, USA). Potato flour, whey protein concentrate, and wheat
bran were supplied by Finnamyl Oy (Kokemak̈i, Finland), HSNG AB
(Solna, Sweden), and Raisio Oy (Raisio, Finland), respectively.
2.2. Synthesis of Regiopure and Enantiopure TAGs.
Regiopure symmetrically structured ABA-type and regio- and
enantiopure asymmetrically structured AAB-type DHA-TAGs and
EPA-TAGs were synthesized. The TAGs were synthesized from pure
palmitic acid (16:0) and DHA or EPA, all supplied by Larodan
(Solna, Sweden). The ABA-type TAGs were synthesized from
glycerol by first obtaining 1,3-DAG 16:0/16:0 with lipase CAL-B
(Novozymes, Gladsaxe, Denmark), followed by incorporation of
DHA or EPA in position sn-2 with coupling agent.15 On the other
hand, the asymmetric AAB-type TAGs were synthesized from R- or S-
solketal (2,2-dimethyl-1,3-dioxolane-4-methanol), after benzyl ether
protection and subsequent deprotection of the isopropylidene group
and incorporation of palmitic acid into the resulting hydroxyl groups
by CAL-B. After deprotection of the benzyl group, DHA or EPA were
incorporated with a coupling agent to the remaining hydroxyl group.25
The presence of BHT in the final product was verified by NP-HPLC
using a Shimadzu Nexera XR LC-30 HPLC instrument (Shimadzu,
Kyoto, Japan) equipped with a LiChroCART Superspher Si 60
column (250 × 4 mm; Merck KGaA, Darmstadt, Germany) using
hexane with an isocratic flow rate of 0.5 mL/min and UV detection at
287 nm. Concentrations of BHT were calculated by injecting standard
solutions of 2.5−100 μg/mL of BHT (Sigma-Aldrich) in hexane.
Traces of BHT in the ABA- and AAB-type DHA-TAGs were
equalized to 0.018% w/w. No BHT could be detected in the ABA-
and AAB-type EPA-TAGs.
2.3. Simulated Digestion. 2.3.1. Simulated Meal. A simulated
meal was prepared for the static in vitro digestion experiments. Potato
flour, whey protein concentrate, and wheat bran were used as
carbohydrate, protein, and fiber sources, respectively. The carbohy-
drate, protein, and fiber contents were calculated from the
composition information provided by the manufacturers. Energy
intake was used to calculate the composition of the simulated meal:
52.5% of the energy was derived from carbohydrates, 15% from
proteins, and 32.5% from lipids. Fiber was added with the ratio 1 g/
79.7 kcal of meal. Potato flour, whey protein, and wheat bran were
defatted with a chloroform/methanol 2:1 v/v (although dichloro-
methane should be preferred) extraction for 48 h. The simulated meal
was prepared by mixing the ingredients in adequate ratios and heating
at 100 °C for 20 min after the addition of deionized water. Simulated
meal, and the lipid component were added separately.26 The lipid
components were DHA-EE, DHA-TAG, and enantio- and regiopure
TAGs containing DHA and EPA in sn-1, sn-2, or sn-3 positions.
2.3.2. Static In Vitro Digestion. The simulated in vitro digestion
was performed according to the INFOGEST 2.0 protocol23 with
modifications. The whole procedure was carried out in dim light,
while digestion itself was carried out in darkness. The simulated saliva
fluid, simulated gastric fluid, and the simulated intestinal fluid were
prepared according to the protocol. Enzyme solutions in their
respective simulated fluids were prepared the day of the experiments.
The required aliquots of CaCl2·2H2O 0.3 M were added to the
simulated fluids only prior to enzyme addition. Solutions were
preheated to 37 °C before use. The pH was adjusted with HCl and
NaOH 1 M solutions. The required aliquots of HCl and NaOH
solutions were verified prior to digestions. Simulated oral phase: 0.90 g
of simulated meal and 30 mg of lipid component were added to 1 mL
of simulated saliva fluid, containing alfa-amylase 75 U/mL. The pH of
the mixture was adjusted to 7. Chewing process was simulated with a
clean glass rod by randomly striking the mixture 32 times.
Subsequently, the mixture was incubated for 2 min at 37 °C.
Chewing time was excluded from the incubation time. Simulated
gastric phase. The simulated oral bolus was added to 1 mL of
simulated gastric fluid containing pepsin and rabbit gastric lipase to
reach enzymatic activity of 2000 U/mL and 60 U/mL, respectively
(final chyme volume). After pH adjustment to 3, the mixture was
incubated for 2 h at 37 °C. Simulated intestinal phase. The simulated
gastric chyme was added to 2 mL of simulated intestinal fluid
containing pancreatin and bile salts to reach concentrations in the
digesta of 6 g/L27 and 10 mM, respectively. After pH adjustment to 7,
the mixture was incubated for 2 h at 37 °C.
2.3.3. Lipid Extraction. Lipids were extracted immediately after
digestion using hexane/isopropanol (2:1, v/v). BHT was added to the
extraction solvent at a concentration of 0.05% in order to prevent
further oxidations.28 Throughout the procedure, samples were kept on
ice. The solvent was added to the digestates in a ratio of 1:1 v/v, and
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tubes were vortexed for 10 s and centrifuged at 1000 rpm for 3 min.
The extraction was performed twice. The upper phases were
combined and evaporated to dryness with N2 at 37 °C. Samples
were recovered with 3 mL of chloroform/methanol (1:1, v/v) and
stored at −80 °C.
2.4. Fatty Acid Analysis. The fatty acids in the samples were
methylated for GC analysis. A methanolic hydrochloric acid mixture
was prepared by adding cold acetyl chloride (Sigma-Aldrich St. Louis,
MO, USA) to cold methanol (1:10, v/v) in ice bath. Heptadecanoic
acid (C17:0, Sigma-Aldrich) was used as an internal standard.
Aliquots of digestates, after addition of an internal standard, were
dried, added to 2 mL of methanolysis mixture, and incubated
overnight at 50 °C. Analysis was carried out with a Shimadzu GC-
2030 equipped with an autoinjector and an FID detector. Analytical
details have been reported elsewhere.24
2.5. Non-volatile Oxidation Product Analysis. Non-volatile
oxidation products were analyzed by Elute UHPLC and Bruker
Impact II quadrupole time-of-flight (QTOF) instruments from Bruker
Daltonic (Bremen, Germany) and a Phenomenex (Torrance, CA,
USA) Kinetex 2.6 μm PS C18 column (100 × 2.1 mm). The method
was modified from previous work.12 Temperatures for the column
oven and autosampler cooler were 30 and 10 °C, respectively.
Samples were diluted to 0.01 mg/mL in MeOH/chloroform (1:1, v/
v), and 1 μL was injected. A binary solvent system was applied for
analyte separation. Solvent A consisted of 95% water (ultrapure from
the Purelab Chorus instrument, Elga Veolia, High Wycombe, UK),
5% methanol (Honeywell/Riedel de Haen̈, Seelze, Germany), and 10
mM ammonium formate (Sigma-Aldrich, Steinheim, Germany).
Solvent B comprised of 70% 2-propanol (Honeywell/Riedel de
Haen̈, Seelze, Germany), 30% methanol, 0.1% water, and 10 mM
ammonium formate. For the TAG samples, the LC gradient program
was as follows: B increased from 60% to 90% in 3 min, to 96.5% in 8
min, to 100% in 0.5 min, held for 2 min, decreased to 60% in 0.5 min,
and held for 3 min. For the ethyl ester samples, the following
modifications were applied: B increased from 60% to 78% in 3 min, to
85% in 8 min, to 100% in 0.5 min, held for 2.5 min, decreased to 60%
in 0.5 min, and held for 4.0 min. The total run time was 18.5 min with
a flow rate of 0.3 mL/min. Electrospray ionization (ESI) was applied
in the positive mode. The capillary voltage was set to 4.5 kV, and the
end plate was offset set to 500 V. Nebulizer gas pressure, drying gas
flow rate, and drying gas temperature were 1.5 bar, 4 L/min, and 350
°C, respectively. Auto MS/MS scanning mode from 60 to 1200 m/z
was applied. Internal calibration was performed with sodium formate.
The concentration of the samples was equalized before analysis. Peak
area values were obtained with Bruker Compass DataAnalysis (Bruker
Daltonic, Bremen, Germany).
2.6. NMR Spectroscopy. 1H NMR spectra were collected from
undigested lipid components and lipids extracted from digesta. For
the analysis, 500 μL of sample was dried with N2 flow and recollected
to 200 μL of CDCl3/DMSO-d6 (5:1, v/v, previously dried with 4 Å
molecular sieves), of which 180 μL was pipetted into 3 mm NMR
tubes. Samples were prepared the previous day before the analysis and
stored overnight at −20 °C in desiccators. 1H NMR spectra were
collected at 298 K with a Bruker Avance 600 MHz (Bruker Biospin,
Switzerland) equipped with a Prodigy TCI CryoProbe and SampleJet
sample changer. Proton spectra were collected with 32 scans, an
acquisition time of 4 s, and a relaxation time of 5 s. A selective
gradient excitation pulse program (selgpse) was applied for region-
specific excitation of hydroperoxide (11.5−10.5 ppm) and aldehyde
(10−9 ppm) protons. The program had 4 dummy scans and 128
scans, with an acquisition time of 2.7 s and a relaxation time of 5 s.
The 180-degree shaped pulse had a length of 1566.15 s.12 NMR data
were processed with TopSpin 4.0.7 (Bruker, MA, USA).
2.7. Statistical Analysis. The statistical analysis was performed
with RStudio.29 Shapiro−Wilk and Bartlett tests (stats package) were
used to assess the normality of the data and the homogeneity of the
data variance. A t-test was performed with the function t_test (rstatix
package). Analysis of variance was performed with the functions aov
and TukeyHSD (stats package) for the ANOVA test and Tukey’s
posthoc test or tamhaneT2Test (PMCMRplus package) for Tamha-
neT2 posthoc test. The statistical significance of differences among
samples was considered at a confidence level of 95% (p < 0.05).
Principal component analysis of the HPLC-qTOF data was performed
with the PCA function (FactoMineR package) of RStudio. The most
important loadings of the obtained model were selected according to
their contribution to each computed principal component, using as
cutoff the reverse number of compounds used to compute the PCA
model.
3. RESULTS AND DISCUSSION
3.1. Fatty Acid Ratio and Fatty Acid Composition. As
PUFAs are consumed during the oxidation process, the
decrease in the PUFA:16:0 ratio can be utilized as an
oxidation indicator for regio- and enantiopure lipids. The
ratios were calculated utilizing the fatty acid composition of
digestates obtained from the gas-chromatographic analysis
(Figure 1). The fatty acid composition of undigested and
digested samples is reported in Supporting Information Figure
S1. Prior to digestion, the PUFA:16:0 ratio was about 0.60 for
all regio- and enantiopure lipids. After digestion, there was a
statistically significant (p < 0.05) decrease in ratio for DHAsn-1
(0.58), DHAsn-3 (0.58), and EPAsn-1 (0.56) (Figure 1). In
the case of DHAsn-2, the average ratio was higher after
digestion (0.67 against 0.59, p < 0.05), and the same was
observed for EPAsn-2, although with a lack of statistical
significance (0.64 against 0.60, p > 0.05). Finally, EPAsn-3 had
a decrease in ratio after digestion but with a high deviation
(0.58 against 0.59, p > 0.05). Auto-oxidation experiments have
assigned position sn-2 a protective effect against oxidation.11,30
In addition to this, the rabbit gastric lipase utilized in the
gastric phase is stereoselective for position sn-3,31 therefore
determining a release of 16:0 from this position in case of
DHAsn-2 and EPAsn-2. Also, this enzyme has lower affinity for
long-chain and unsaturated fatty acids.32 The high deviation
for EPAsn-3 could be explained by the hydrolytic effect of
Figure 1. PUFA (DHA and EPA) to 16:0 weight ratio of synthesized
ABA- and AAB-type TAGs containing DHA and EPA in either sn-1,
sn-2, or sn-3 positions and 16:0 in the remaining positions prior to and
after digestion. The letter (d) marks the digestates. Values are average
± standard deviation (n = 2). The symbol marks statistically
significant difference (p < 0.05).
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pancreatic lipase, which is less selective for position sn-3 and
also effects position sn-1. On the other hand, the difference in
the PUFA:16:0 ratio in these two digestates could also be
explained by the higher resistance to oxidation by EPA
compared to DHA in similar chemical structures.31,33 The gas
chromatographic analysis of DHA-TAG and DHA-EE
digestates showed a decrease in DHA compared to the
standard compound of 8.9 ± 0.2 and 9.2 ± 0.7%, respectively.
It can be deduced that the PUFA amounts in digestates would
be influenced by different factors, such as oxidation (and
protection from) and lipase activity.
3.2. Non-volatile Digestion and Oxidation Com-
pounds. 3.2.1. Identification of Undigested Compounds
and Digestion Products. Compound peak area data was
obtained from the extracted ion chromatograms of undigested
standards and digestate extracts. Compounds were identified as
ammonium and sodium adducts. Table 1 reports mass values
(m/z) for the identified compounds. Ammonium adducts were
the most abundant for unoxidized TAGs and DHA-EE,
whereas oxidation products, DAGs (except DAG 16:0/22:6)
and MAGs, had the sodium adduct as the most abundant. The
integration results are reported in Figures 2 and 3. Details for
the identification of undigested lipids, digestion compounds,
and primary oxidation products are reported in Supporting
Information Text S1.
3.2.2. Influence of Lipid Structure on the Formation of
Digestion Products. At our experimental conditions, no
statistically significant decrease in the area of AAB- and
ABA-type TAGs was observed, whereas, in the case of DHA-
TAG and DHA-EE, the area of undigested molecules
increased. This could be attributed to a higher ionizability of
the molecules, likely due to residual salts in the digestate
extracts. In terms of DAGs, the comparison of AAB- and ABA-
type TAGs showed that DHAsn-2 produced more (p = 0.003)
DAG 16:0/22:6 than DHAsn-3. Despite the higher affinity of
lipase for sn-3 than sn-1, production of DAG 16:0/22:6 from
DHAsn-1 and DHAsn-3 had no difference. EPAsn-1 produced
DAG 16:0/20:5 significantly more than DHA-containing
TAGs (p < 0.05), but no more than EPAsn-3. The peak area
values of DAG 16:0/PUFA and DAG 16:0/16:0 (Figure 2)
were generally higher for EPA-containing TAG compared to
DHA, indicating higher lipolysis. This could be explained by
the higher steric hindrance of 22:6 compared to 20:5,34
especially regarding the sn-3 position, which is preferentially
hydrolyzed, and by the decrease in lipolytic activity of rabbit
gastric lipase with the increase in carbon chain length.32 DAG
Table 1. Compounds and Their Ammonium [M + 18]+ and Sodium [M + 23]+ Adduct Masses (m/z) Identified in HPLC-
qTOF Chromatograms, together with Their Retention Times and Fragments Used for Identificationa
m/z [M + 18]+ [M + 23]+
r.t.
(min) main fragments m/z [M + 18]+ [M + 23]+
r.t.
(min)
main
fragments
DHA
regio-/enantiopure
EPA
regio-/enantiopure
16:0/16:0/22:6 878.7 896.7 901.7 9.5 551.5, 623.5,
313.3, 311.3,
239.2
16:0/16:0/20:5 852.7 870.7 875.7 8.8 551.5, 597.5,
313.3
16:0/16:0/22:6 +
1O (16)
894.7 912.7 917.7 7.8 367.2, 661.5,
551.5
16:0/16:0/20:5 +
1O (16)
868.7 886.7 891.7 7.6 617.5, 341.2,
551.5
16:0/16:0/22:6 +
1O (14)
892.7 910.7 915.7 7.2 365.2, 659.5,
551.5
16:0/16:0/20:5 +
1O (14)
866.7 884.7 889.7 7.0
16:0/16:0/22:6 +
2O
910.7 928.7 933.7 6.8 887.7, 806.6,
847.7, 383.2
16:0/16:0/20:5 +
2O
884.7 902.7 907.7 6.6 781.5, 357.2,
551.5
16:0/16:0/22:6 +
4O
942.7 960.7 965.7 5.9 16:0/16:0/20:5 +
4O
916.7 934.7 939.7 5.7
16:0/16:0 568.5 586.5 591.5 5.3 313.3, 551.5,
339.3
16:0/16:0 568.5 586.6 591.5 5.3 313.3, 551.5,
339.3
16:0/22:6 640.5 658.5 663.5 5.1 313.3, 385.3,
641.5
16:0/20:5 614.5 632.5 637.5 4.9 313.3, 359.3,
285.2
16:0/22:6 + 1O
(16)
656.5 674.5 679.5 4.6 367.2, 661.5,
313.3
16:0/20:5 + 1O
(16)
630.5 648.5 653.5 4.5
16:0/22:6 + 1O
(14)
654.5 672.5 677.5 4.4
16:0/22:6 + 2O 672.5 690.5 695.5 4.3 16:0/20:5 + 2O 646.5 664.5 669.5 4.2
MAG 16:0 330.3 348.3 353.3 2.9 335.3, 226.2,
263.2
MAG 16:0 330.3 348.3 353.3 2.9 335.3, 226.2,
263.2
MAG 22:6 402.3 420.3 425.3 2.7 311.2, 385.3,
403.3, 293.2
MAG 20:5 376.3 394.3 399.3 2.3 267.2, 285.2,
303.2
DHA-TAG DHA-EE
22:6/22:6/22:6 1022.7 1040.7 1045.7 8.2 695.5, 385.3 DHA-EE 356.3 374.3 379.3 4.9 311.2, 293.2
22:6/22:6/22:6 +
1O (16)
1038.7 1056.7 1061.7 7
22:6/22:6/22:6 +
2O
1054.7 1072.7 1077.7 6.2 383.2, 950.6,
991.6
M + 2O 388.3 406.3 411.3 3.2 365.2, 285.1,
245.1,
393.2
22:6/22:6/22:6 +
4O
1086.7 1104.7 1109.7 5.4 M + 4O 420.3 438.3 443.3 1.2
22:6/22:6 712.5 730.5 735.5 4.9 385.3, 311.2,
293.2
22:6/22:6 + 2O 744.5 762.5 767.5 4.2
MAG 22:6 402.3 420.3 425.3 2.7 239.2, 308.8
aMass values in bold represent ion currents used for identification.
Journal of Agricultural and Food Chemistry pubs.acs.org/JAFC Article
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16:0/16:0 (Figure 2) could be either a reminder of TAG
synthesis or a hydrolysis product of PUFAsn-3 (by both
lipases) and PUFAsn-1 (by pancreatic lipase). The observed
increase of DAG 16:0/16:0 after digestion of DHAsn-2 and
EPAsn-2 had no statistical significance (p > 0.1 for both). At
the same time, there was no significant difference between the
areas of DAG 16:0/16:0 after digestion of sn-1 and sn-3 TAGs.
A similar trend was observed with MAG 16:0, found solely in
digestates as a hydrolytic product of either DAG 16:0/16:0 or
DAG 16:0/PUFA, possibly indicating that the steric hindrance
of DAGs generated by PUFAs was lower than in TAGs. The
presence of MAG 16:0 in sn-2 digestates can be explained as
the hydrolytic product of remainder DAG 16:0/16:0 since
neither lipase has affinity for position sn-2. Moreover, its
presence could be explained by the formation, due to acyl
migration, of 1,3-DAGs from 1,2-DAGs and their consequent
hydrolysis to MAG. However, the absence of MAG 22:6 and
MAG 20:5 in the respective sn-1 and sn-3 digestates indicated
acyl migration was a marginal phenomenon under our
experimental conditions. The production of MAG PUFA has
biological relevance, as 2-MAGs are well absorbed by intestinal
mucosa and grant higher bioavailability of PUFAs compared to
1- or 3-MAGs.17
Digestion of tridocosahexaenoin produced, as expected,
DAG 22:6/22:6 and MAG 22:6 (Figure 2). The area of the
latter was statistically similar to the areas of MAG 22:6 and
MAG 20:5 originating from DHAsn-2 and EPAsn-2 (p > 0.05).
Therefore, despite the steric hindrance of DHA, hydrolysis of
position sn-3 was overall similar between DHA-TAG and AAB-
and ABA-type EPA-TAGs. This is possibly due to the absence
of 16:0 in DHA-TAG. At the same time, the area of DAG
22:6/22:6 differed only from DHAsn-1 (p < 0.05) and DHAsn-
3 (p < 0.01), suggesting DAGs were less hindered than TAGs.
Our analytical conditions could not unambiguously assign free
Figure 2. Peak area values (in logarithmic scale) of digestion products
identified in HPLC-qTOF chromatograms of digested docosahex-
aenoin (DHA-TAG) and AAB- and ABA-type TAGs containing DHA
and EPA. For their respective classes of compounds, DAG PUFA
indicates DAG 16:0/22:6 and DAG 16:0/20:5. Values are average ±
standard deviation (n = 3 for DHA-EE and DHA-TAG; for others, n =
2).
Figure 3. Peak area values (in the logarithmic scale) of oxidated compounds identified in HPLC-qTOF chromatograms of undigested and digested
AAB- and ABA-type TAGs, docosahexaenoin (DHA-TAG), and ethyl docosahexaenoate (DHA-EE). Digestates are marked with (d). Values are
average ± standard deviation (n = 3 for DHA-EE and DHA-TAG; for others, n = 2).
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fatty acids, and no digestion products of DHA-EE were
observed.
3.2.3. Influence of Lipid Structure on Oxidized Com-
pounds during Digestion. According to our results, simulated
digestion under our experimental conditions determined a
reduction in oxygenated AAB- and ABA-type structures and
the appearance of oxygenated digestion products. On the other
hand, an increase in oxygenated species was observed for
DHA-TAG and DHA-EE (Figure 3). Compared to the other
samples, EPAsn-2 had the lowest signals for these molecules,
indicating the protective effect of position sn-2 and higher
stability of this PUFA, as could be expected. Also, only for
EPAsn-2, the decreases in TAG + 16 (1O), TAG + 14 (1O),
and TAG + 32 (2O) lacked statistical significance (0.08 < p <
0.9). Conversely, the decrease in TAG + 64 lacked statistical
significance for DHAsn-2, DHAsn-3, and EPAsn-2. The area
ratio of oxygenated TAGs after and prior to digestion is
reported in Figure 4. For EPAsn-2, the area of most oxygenated
species decreased only about 25% (ratio 0.75), except TAG +
32, which decreased to a ratio of 0.42. On the other hand,
DHAsn-2 had the highest decreases (ratios 0.16, 0.13, and 0.19
for TAG + 16, TAG + 14, and TAG + 32, respectively) except
for TAG + 64, whose decrease had no significant difference (p
> 0.05) with other decreases, as observed in Figure 3. A ratio of
0.5 (a decrease of 50%) was observed for TAG + 64 species of
about all AAB- and ABA-type structures except EPAsn-3.
Overall, EPAsn-2 had the lowest decrease in oxygenated
species, while DHAsn-2 had the highest, without taking into
account the seemingly scantly affected TAG + 64 (Figure 4).
In the case of EPAsn-2, DAG 16:0/20:5 + 32 had about 1.75
times the area of the respective TAG + 32 (Supporting
Information Figure S3). In this case, it can be deduced that the
loss of 16:0 decreased the protective effect of sn-2 and that new
oxidized species were formed. According to our results, there
was no correspondence between the loss of oxygenated TAG
and the area of oxygenated DAG of regio- and enantiopure
compounds, as the areas of the latter corresponded to only 3−
14% of the losses. Therefore, most of the losses of TAG + 16,
TAG + 14 (for DHA), and TAG + 32 compounds were
ascribable to further degradation. Marquez-Ruiz et al.
described hydroxy and keto fatty acids as relatively stable in
digestion conditions, but their experiments were conducted
with a saturated fatty acid. Conversely, their degradation of
hydroperoxidated and epoxidated methyl linoleate was, despite
the difference in chemical structure, in general agreement with
our results.35
Differently from regio- and enantiopure TAGs, oxygenated
DHA-TAG increased after digestion (Figure 3). Moreover, the
addition of one oxygen was observed only in digestates, but
noticeably only in TAG species. The ratio between DAG 22:6/
22:6 + 32 and TAG 22:6/22:6/22:6 + 32 and the ratio
between DAG 22:6/22:6 and TAG 22:6/22:6/22:6 had no
significant difference (p > 0.05). As pancreatic lipase is proven
to act also on oxidized TAGs,36 it could be concluded that,
while DAG 16:0/PUFA + 32 was mainly the oxidation product
of DAG 16:0/PUFA, DAG 22:6/22:6 + 32 arose from lipolysis
of the oxidized TAG. It is believed that TAG lipolysis rate is
not affected by peroxidation under gastrointestinal condi-
Figure 4. Area ratio of oxygenated structural lipids after and prior to
digestion. Values are average ± standard deviation (n = 3 for DHA-
TAG; for others, n = 2). Different letters mark statistically significant
difference (p < 0.05).
Figure 5. Principal component analysis of HPLC-qTOF data of structured lipids. Model computed with all the unoxygenated and oxygenated
species in common between DHA- and EPA-structured lipids (a). Model computed with all the species in common, excluding MAG PUFA (b).
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tions37 and changes in lipase activity are usually observed after
oxidation at cooking temperatures38 or prolonged oxidation39
experiments. Different from TAGs, DHA-EE was the sole lipid
unaffected by digestion, since the area increase was identical
for all the species (Figure 3) and therefore ascribable solely to
a higher ionizability. This finding is in agreement with previous
results from Song and co-workers40 and our group,12 which
have shown, although under different oxidative conditions, the
higher stability of DHA-EE compared to DHA-TAG when
oxidized in the presence of α-tocopherol. However, Sullivan
and co-workers have reported higher oxidation rates for EE of
DHA and EPA in comparison with their TAG forms at 30 °C.
Further studies monitoring primary and secondary oxidation
products could address this discrepancy.41
3.2.4. PCA of HPLC-qTOF Data. Scores and loading plots of
the principal component analysis (PCA) of the HPLC-qTOF
data of structured lipids are reported in Figure 5. The first
principal component separated digestates from undigested
samples and represented more than 75% of the data variance.
In the first model (Figure 5a), digestates of EPAsn-2 and
DHAsn-2 grouped together in the lower right quadrant of the
score plot, while digestates from sn-1 and sn-3 grouped
together in the upper right quadrant. Undigested samples were
all grouped together except for EPAsn-2. Therefore, the second
principal component discriminated sn-2-structured lipids from
the others (14% of the variance). The compounds more
responsible for the discrimination (contribution higher than
10%) were DAG PUFA, MAG 16:0, DAG PUFA + 16, and
DAG PUFA + 32 for PC1 and MAG PUFA for PC2. As MAG
PUFA is found only in sn-2 digestates (and hence its relevance
for discrimination), a PCA model was computed without this
compound (Figure 5b). In the second model, digestates were
discriminated again according to DAG PUFA, MAG 16:0,
DAG PUFA + 16, and DAG PUFA + 32. On the other hand,
the discrimination on the second principal component was
determined by oxidized TAGs (TAG + 16, TAG + 32, and
TAG + 64). Score values on the second principal component
are highlighted in Supporting Information Figure S6. As
position on PC2 was determined by oxidation status prior to
and after digestion, EPAsn-2 was positioned farthest from the
other samples. However, while prior to digestion, EPAsn-1 was
the second-farthest sample, EPAsn-2 had lower scores after
digestion, while the score of EPAsn-1 increased. This
positioning hinted at a correlation trend between EPA position
and amounts of oxidized TAGs after digestion: sn-2 < sn-3 <
sn-1. The protective effect of position sn-2 has already been
reported and has been attributed to steric hindrance.11,30
Moreover, this protective effect has also been observed in
emulsion systems.14 Differences between sn-1 and sn-3 have
been scarcely addressed in the literature, as these compounds
are difficult to obtain. In our group, slightly higher stability of
sn-3 compared to sn-1 was observed during auto-oxidation of
DHA-containing regio- and enantiopure TAGs in the absence
of antioxidants, while the opposite was observed in their
presence.11 The lower correlation of EPAsn-3 with oxidized
TAGs after digestion indicated a higher reduction in their
amounts compared to EPAsn-1 as their positioning on PC2
switched. Therefore, the PCA hinted at a higher stability at
digestion conditions of oxygenated PUFA in position sn-1,
meaning that EPAsn-1 hydroperoxides could be more stable
than in EPAsn-3. However, it is also possible that EPAsn-1
hydroperoxides degraded and new species formed at a higher
rate than EPAsn-3, which would indicate higher stability of
EPAsn-3 compared to EPAsn-1. This hypothesis would be
supported by Larsson et al., who reported little change in
hydroperoxide amounts in cod oil after simulated digestion but
a net increase in secondary oxidation products.42 Nevertheless,
further experiments are being undertaken by our group to
verify the stability of EPA-containing regio- and enantiopure
TAGs during auto-oxidation. More studies on the detailed
kinetic profile of primary oxidation products at simulated
digestion are also required.
3.3. 1H NMR Spectroscopy. The collected 1H NMR
spectra are reported in Supporting Information Figures S4 and
S5. All the TAG samples showed the typical signal pattern,
with the sn-1/3 and sn-2 position signals around 4.14, 4.29, and
5.24 ppm, respectively. Conversely, the DHA-EE spectrum had
the methylene signal of the ethyl chain at 4.12 ppm. In ABA
and AAB structures, the terminal methyl signal of 16:0 was
observed at 0.88 ppm, while the terminal methyl signal of the
ethyl chain of DHA-EE was at 1.25 ppm. In DHA-TAG, DHA-
EE, and structured lipids, the terminal methyl signal of PUFA
was at 0.97 ppm. After digestion of DHA-EE, a small but
significant decrease in the ratio between ethyl −CH2− and
terminal −CH3 of DHA was observed (12.2%, p = 0.007),
which can be attributed to the hydrolytic activity of lipase.
Lower hydrolytic rates of ethyl esters, compared to TAGs, have
also been observed by others43 and have been considered an
explanation for the lower bioavailability of PUFAs in this
chemical form.9 On the other hand, after digestion of TAGs,
DAG and MAG signals were observed. The signals were
ascribed to 1,2-DAG (5.07, 4.35, and 3.64 ppm) and 2-MAG
(4.88 and 3.70 ppm). In digestates, the quartet at 4.14 ppm
was overlapping with another signal, which could either be due
to 1,2-DAG or 1,3-DAG. The latter, if present, could not be
clearly assigned as it typically produces a broad multiplet at
4.04−4.21 ppm.15
The 1H NMR spectra obtained after selective gradient
excitation of the hydroperoxide region are reported in
Supporting Information Figure S7. Regio- and enantiopure
TAGs with DHA and DHA-TAG had similar peroxide proton
signals, which were observed at 10.93, 11.04, and 11.07 ppm.
The first signal can be assigned to a linear structure, the other
two to cyclic forms, or to hydroperoxides positioned farther
from the terminus of the fatty acid chain.44 Regio- and
enantiopure TAGs with EPA had an additional proton peak at
11.01 ppm, which could belong either to a linear or cyclical
structure.44 Among all samples, EPAsn-2 and DHA-EE were
the sole proton spectra completely lacking hydroperoxide
signals, confirming the lower degree of oxidation observed with
HPLC-qTOF (Section 3.2 and Figure 2b). After digestion,
hydroperoxide proton signals disappeared from the spectra.
For AAB- and ABA-type structures, this confirmed the
reduction in oxygenated species observed with HPLC-qTOF
(Figure 2b). The disappearance of hydroperoxide protons in
DHA-TAG digestates could indicate that hydroperoxides are
further degraded to hydroxides or epoxides. Such a pathway
would be confirmed by the appearance of TAG + 16 after
digestion of DHA-TAG (Figure 2b). Peroxidated bisallylic
systems are considered to produce a hydroxy moiety after
radical scission of the O−O bond and quenching of the
generated alkoxy radical. Alternatively, the radical would react
with the proximal unsaturated carbon, generating an
epoxide.45,46 Scission radicals further continue the oxidation
pathway to secondary products such as aldehydes, which are
reactive with proteins and DNA. The most toxic ones are
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malondialdehyde and 4-hydroxy-2-alkenals.7 The earlier
originates from cyclic hydroperoxides, the latter from further
oxidation of hydroxy-PUFA.5 Different factors, such as micelle
inclusion and steric hindrance, might have impeded the radical
quenching activity of α-tocopherol in DHA-TAG and rather
favored oxidation, while DHA-EE was more stable and lacked
hydroperoxide protons even prior to digestion. In the case of
ABA and AAB-type TAGs, no antioxidant was added (besides
BHT trace normalization) and, therefore, oxidative degrada-
tion was possibly ruled by steric hindrance (i.e., position of the
PUFA). At the same time, the ratio between PUFA and 16:0
(Figure 1) indicated a lower presence of abstractable protons
in the structure, compared to DHA-TAG.
3.4. Considerations for Oxidation in the Gastro-
intestinal Tract. According to the recent work of Takahashi
et al., when partially hydroperoxidized or hydroxylated TAGs
were introduced into the rat stomach, these compounds were
also detected in the lymph, while these compounds were
absent when no oxidized molecule was introduced. At the same
time, deuterium labeling indicated no absorption of hydro-
peroxidized or hydroxylated TAGs. Therefore, oxidation
damage propagated in the gastrointestinal tract. The same
authors reported the reduction of hydroperoxidized fatty acids
to hydroxy species not only in vivo but also ex vivo with rat
small intestine mucosa homogenate.47 This reduction has also
been observed in Caco-2 cell lines by other authors.48
While there is general agreement on the pro-oxidant
conditions of the gastrointestinal tract, the effect of lipolysis
on lipid oxidation is less understood. Larsson et al. were among
the first ones to observe an increase, although little, in primary
and secondary oxidation products of cod oil at simulated
digestion conditions after addition of fungal lipase (although
now considered inadequate to represent human lipolysis23).
They also noticed higher amounts of secondary oxidation
products after the addition of bile salts, pointing to the
importance of micelles for the oxidation pathway.42 In
addition, Tullberg et al. connected the marked increase in 4-
hydroxy-nonenal, originating from n-6 PUFAs, in cod liver oil
digestates to the presence of rabbit gastric lipase. On the other
hand, the presence of gastric lipase had no effect on the marker
of n-3 PUFAs degradation, 4-hydroxy-hexenal, during the
gastric phase, while the marker increased during the intestinal
phase.28 This was in agreement with the positional distribution
of fatty acids in cod liver oil, which had 18:2n-6 mainly in sn-
1,3 positions and DHA solely in sn-2 position.49 Compared to
the previous studies, the present work demonstrated that
primary oxidation products arose during digestion, and the
extent depended on the lipid structure. They degraded to
hydroxides or epoxides in the simulated gastrointestinal tract
medium. Our work excluded the necessity of epithelial cells for
propagation, pointing to the importance of micelles. The
decrease in oxidized molecules during simulated digestion
conditions can be ascribed to their chemical instability, as
mentioned previously. This degradation would continue
oxidation cascade in the gastrointestinal lumen or in micelles,
while epoxides and ketones can in addition covalently bind
proteins. Liberation of oxidized fatty acids from position sn-3
would likely facilitate this process. At the same time, hydrolysis
of position sn-3 decreases the protection of PUFA in position
sn-2. While epoxides are reactive and expected to degrade prior
to absorption, hydroxides, on the other hand, are proven to be
absorbed during digestion,50 hence potentially degrading at
other sites. Therefore, hydroperoxidation can exert damage
locally or after absorption. Further research on the improve-
ment of simulated digestion conditions (amount of required
pancreatin or presence of epitelial cells) to more accurately
represent the small intestine is therefore required to better
understand the dynamics of hydroperoxide degradation.
In conclusion, the present study investigated the simulated
digestion of DHA-EE, DHA-TAG, and AAB- and ABA-type
enantio- and regiopure TAGs containing DHA and EPA. As
previously observed in auto-oxidation trial, DHA-EE showed
higher oxidative stability than DHA-TAG in simulated
digestion conditions. While DHA-TAG showed an increase
in oxidation species during simulated digestion, it generated
MAG 22:6, which would grant absorption of PUFA, whereas
DHA-EE was only partially hydrolyzed at our digestion
conditions. All AAB- and ABA-type TAGs generated MAGs
and DAGs, the latter also present as oxidized species.
Simulated digestion determined a decrease in oxidized
TAGs. In general, EPA was more resistant to oxidation and
less resistant to digestion compared to DHA. While the higher
stability of PUFAs in sn-2 has been reported previously, at
simulated digestion conditions, only EPAsn-2 was clearly the
most stable. In addition, our results hint at a higher
degradation rate of EPAsn-1 hydroperoxides compared to
EPAsn-3. Finally, only DHAsn-2 and EPAsn-2 generated MAG
22:6 and MAG 20:5, which are more efficiently absorbed than
the corresponding free fatty acids, giving relevance to our
finding regarding bioavailability.
■ ASSOCIATED CONTENT
*sı Supporting Information
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acs.jafc.3c02207.
Identification of undigested compounds and of digestion
and oxidation products; fatty acid composition of DHA-
containing structured lipids and EPA-containing ABA
and AAB structures, DHA-TAG, and DHA-EE; HPLC-
qTOF ion currents of DHAsn-2 before and after
digestion; ratio between oxygenated DAGs and TAGs
in digestates of ABA- and AAB-type TAGs and DHA-
TAG; 1H NMR spectra of ABA and AAB structures,
DHA-EE, and DHA-TAG prior to and after digestion;
average score values on the second principal component
of the PCA model computed with HPLC-qTOF data of
structural lipids after removal of MAG PUFA; and
selective gradient excitation 1H NMR spectra of ABA-
and AAB-structures, DHA-EE, and DHA-TAG prior to
and after digestion (PDF)
■ AUTHOR INFORMATION
Corresponding Author
Kaisa M. Linderborg − Food Sciences, Department of Life
Technologies, University of Turku, FI-20014 Turku,
Finland; orcid.org/0000-0003-1977-7322; Phone: +
358 50 439 5535; Email: kaisa.linderborg@utu.fi
Authors
Gabriele Beltrame − Food Sciences, Department of Life
Technologies, University of Turku, FI-20014 Turku, Finland
Eija Ahonen − Food Sciences, Department of Life
Technologies, University of Turku, FI-20014 Turku, Finland
Journal of Agricultural and Food Chemistry pubs.acs.org/JAFC Article
https://doi.org/10.1021/acs.jafc.3c02207
J. Agric. Food Chem. 2023, 71, 10087−10096
10094
Annelie Damerau − Food Sciences, Department of Life
Technologies, University of Turku, FI-20014 Turku,
Finland; orcid.org/0000-0002-3495-2419
Haraldur G. Gudmundsson − Faculty of Pharmaceutical
Sciences, University of Iceland, IS-107 Reykjavík, Iceland
Gudmundur G. Haraldsson − Faculty of Physical Sciences,
University of Iceland, IS-107 Reykjavík, Iceland
Complete contact information is available at:
https://pubs.acs.org/10.1021/acs.jafc.3c02207
Author Contributions
G.B.: investigation, formal analysis, visualization, writing�
original draft, and funding acquisition; E.A.: methodology,
investigation, and writing�review and editing; A.D.: con-
ceptualization, methodology, investigation, validation, and
writing�review and editing; H.G.G.: investigation and
writing�review and editing; G.G.H.: methodology and
writing�review and editing; K.L.: conceptualization, writ-
ing�review and editing, supervision, project administration,
and funding acquisition.
Funding
A personal financial grant to Gabriele Beltrame from the Niemi
Foundation is acknowledged. Personal financial grants to
Annelie Damerau from the Finnish Cultural Foundation and to
Eija Ahonen from the Niemi Foundation, the Finnish Cultural
Foundation, and the Finnish Food Research Foundation are
acknowledged. This work was carried out as part of the project
“Omics of oxidation�Solutions for better quality of
docosahexaenoic and eicosa-pentaenoic acids” funded by the
Academy of Finland (grant number 315274, PI Kaisa
Linderborg).
Notes
The authors declare no competing financial interest.
■ ACKNOWLEDGMENTS
The authors would like to thank Erika Tatiana Cortes Macias
for her help in the GC-FID analysis and Hafdís Haraldsdóttir
for her help in the interpretation of the NMR spectra.
■ ABBREVIATIONS
ANOVA, analysis of variance; BHT, dibutylhydroxytoluene;
CDCl3, deuterated chloroform; DAG, diacylglycerol; DHA,
docosahexaenoic acid; DMAP, 4-dimethylaminopyridine;
DMSO-d6, deuterated dimethylsulfoxide; DCl, 1-ethyl-3-(3-
dimethylaminopropyl)carbodiimide; EE, ethyl ester; EPA,
eicosapentaenoic acid; GC-FID, gas chromatography coupled
with flame ionization detection; HPLC-qTOF, high-perform-
ance liquid chromatography with triple quadrupole-time-of-
flight detection; MAG, monoacylglycerol; NMR, nuclear
magnetic resonance; PCA, principal component analysis;
PC1(2), principal component 1(2); PUFA, polyunsaturated
fatty acid; TAG, triacylglycerol
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