# Incomplete lipid extraction as a possible cause for underestimation of lipid oxidation in emulsions

Katharina Münch, Sten ten Klooster, Isabelle van Kouwen, Claire

Berton-Carabin, Karin Schroën

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## To cite this version:

Katharina Münch, Sten ten Klooster, Isabelle van Kouwen, Claire Berton-Carabin, Karin Schroën. Incomplete lipid extraction as a possible cause for underestimation of lipid oxidation in emulsions. European Journal of Lipid Science and Technology, 2024, 126 (6), 10.1002/ejlt.202300229. hal-04698288

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RESEARCH ARTICLE

# Incomplete lipid extraction as a possible cause for underestimation of lipid oxidation in emulsions

Katharina Münch<fnref n="1" /> | Sten ten Klooster<fnref n="1" /> | Isabelle van Kouwen<fnref n="1" /> | Claire Berton-Carabin<fnref n="1" />,<fnref n="2" /> | Karin Schroën<fnref n="1" />

Correspondence

Katharina Münch and Karin Schroën,
Laboratory of Food Process Engineering,
Bornse Weilanden 9, 6708 WG Wageningen,
The Netherlands.

Email: katharina.muench@wur.nl and
karin.schroen@wur.nl

Funding information

Dutch Research Council, Grant/Award
Number: 731.017.301; Nederlandse
Organisatie voor Wetenschappelijk
Onderzoek

Abstract

Lipid oxidation deteriorates the sensory and nutritional quality of food emulsions containing polyunsaturated fatty acids. Classically, different extraction solvents are used as a first step to measure lipid oxidation in emulsions. However, it is unclear how the applied extraction method influences the measured lipid oxidation values. In this work, we systematically examined the performance of common solvent mixtures such as chloroform, methanol, and hexane (or isooctane)-isopropanol on lipid extraction from emulsions stabilized with different emulsifiers (Tween 20 (T20), whey proteins, and pea proteins) and oxidation levels, and how this, in turn, affected the measured hydroperoxide concentrations. Chloroform-methanol was the most effective solvent (lipid yield >93 wt.%). When using hexane-isopropanol, extraction yields were consistently high for T20- and pea protein-based emulsions (>60 wt.%), but in whey protein-based emulsions, values as low as 26 wt.% were measured. In case of incomplete extraction, hydroperoxide concentrations measured by colorimetric methods need to be corrected for this effect. When using ¹H NMR to assess lipid oxidation, the actual amount of extracted lipids is intrinsically taken into account. This highlights not only the importance of the extraction method in determining lipid oxidation in emulsions but also that of the actual analysis method.

**Practical application:** This study highlights that the lipid extraction yield can vary depending not only on the emulsion composition (e.g., type of emulsifier) but also on the oxidative state of the emulsion and the extraction solvent used. If this is overlooked, errors can be made in the hydroperoxide determination. Although these effects can be corrected for, this is not standard procedure, which implies that awareness on this matter should be increased. It is also important to point out that depending on the solvent used, the different lipid classes (including various lipid oxidation products) may be extracted at different levels. Chloroform-methanol should be preferred for extraction of all lipid and lipid oxidation-derived molecules, including aldehydes.

<footnote n="1">¹Wageningen University & Research, Laboratory of Food Process Engineering, Wageningen, The Netherlands</footnote>

<footnote n="2">²INRAE, UR BIA, Nantes, France</footnote>
KEYWORDS

¹H NMR, hydroperoxide assay, lipid extraction, lipid oxidation, methodology

1 | INTRODUCTION

Food emulsions and other natural or manufactured products contain-
ing polyunsaturated fatty acids are prone to lipid oxidation, which
deteriorates the quality of the food and thus is a concern in food
production.[1] Lipids in emulsions are especially prone to oxidation due
to the high interfacial area in emulsions where lipid oxidation is initi-
ated, and a lot of work in the field has been dedicated to preventing
this by various strategies.[2-8]

To measure lipid oxidation in emulsions, lipid extraction via organic solvents is a classical preliminary step for many methods used in literature.[9] One of the most frequently used methods to measure the hydroperoxide (primary oxidation products) concentration is the colorimetric method described by Shantha and Decker.[10] The original method used hexane-isopropanol (HIP) as an extraction solvent (ratio 3:2 v/v) and chloroform-methanol (7:3 v/v) for further measurements. Over time, the method was extended to the use of other solvent mixtures, for example, isooctane-isopropanol (IOIP), chloroform-methanol in the ratio of 2:1 v/v (Folch), or a ratio of 3:1 instead of 3:2.[11-13] As most of these methods assume a total extraction of the lipids (any lipid molecules present in the oil, including lipid oxidation products), the results may be skewed for incomplete extraction since they are expressed per kg of oil.[9,14] Besides, extraction solvents differ in polarity, and depending on the solvent used, classes of lipids that vary in polarity may be extracted more preferentially, and thus be under or overrepresented in the extract.

In addition, emulsifiers may influence the extraction of lipids by
interacting with the extraction solvents, for example, by forming a net-
work of denatured proteins thereby limiting the extraction of lipids.[15]
From this, we suspect that both the choice of the extraction method as
well as the emulsifier may have pronounced effects on the amount of
extracted oil (and composition), and thus on the determined concen-
tration of oxidation products that depend on the extracted amount of
oil. Surprisingly, to date, this has not been investigated systematically,
and that is therefore the goal of the present work.

To shed light on this matter, we measured the lipid extraction yield for different commonly used solvents, namely, HIP, IOIP, and Folch, for emulsions stabilized by Tween 20 (T20), whey protein isolate (WPI), or pea protein isolate (PPI), freshly prepared and under oxidizing incubation conditions. The extracts were analyzed for oxidation products using the colorimetric method described by Shantha and Decker and the ¹H NMR method described by Merkx et al.[10,16] from which concentrations of oxidation products can be determined independently of the extraction yield. Results were compared and put into perspective for possible implications of the extraction method.[3,8] On the basis of our outcomes, we advise on methods to obtain accurate lipid oxidation

results for different extraction solvents considering also the emulsifier
used.

2 | MATERIALS AND METHODS

2.1 | Materials

Rapeseed oil (kindly supplied by Unilever) was stripped from impurities and tocopherols by using alumina powder (MP EcoChrome ALUMINA N, Activity: Super I, Biomedicals). This procedure has proved useful to accelerate lipid oxidation in model systems and holds the advantage of standardizing the properties of the oil.[17,18] PPI, protein content 78% (N x 5.6; NUTRALYS S85F, Roquette), WPI, protein content 97.0%-98.4% (BiPro, Davisco), and Tween 20 (T20, Sigma Aldrich) were selected as emulsifiers. Methanol, 2-propanol, and *n*-hexane were supplied by Actu-All Chemicals. 2,2,4-trimethylpentane (isooctane) was obtained from Alfa Aesar. Deuterated chloroform (CDCL₃) with 0.03% tetramethylsilane, deuterated dimethylsulfoxide, and deuterated 4 Å mol sieves were purchased from Euriso-top. Hydrochloric acid (2 N) was obtained from Merck. Butanol, ethyl-acetate, chloroform, sodium phosphate dibasic (Na₂HPO₄), sodium phosphate monobasic (NaH₂PO₄), sodium chloride, sodium azide, sodium dodecyl sulfate, 2,4-dinitrophenylhydrazine (DNPH), thiocyanate (NH₄SCN), guanidine hydrochloride, trichloroacetic acid (TCA) 40%, 0.144 M iron(II)sulfate heptahydrate (FeSO₄·7H₂O), 0.132 barium chloride (BaCl₂), ascorbic acid, and a bicinchoninic acid (BCA) kit (BCA1-1 KT) were purchased from Sigma-Aldrich. Ultrapure water obtained from a Milli-Q system (Millipore Corporation) was used throughout all experiments.

## 2.2 | Preparation of continuous phase solutions

The pea protein solution was prepared by dispersing 10 wt.% PPI in 10 mM phosphate buffer (pH = 7.0) and hydrated for at least 48 h at 4°C. The soluble protein fraction was obtained by centrifuging (16 000 × g, 30 min) the protein suspension and collecting the supernatant, which was then centrifuged again under the same conditions. The protein concentration of the supernatant was determined with the Dumas method.[19] The supernatant, further referred to as “soluble pea protein” (sPPI), was then diluted to a concentration of 1 wt.% protein in the final emulsion. For WPI and T20, a solution was directly prepared in 10 mM phosphate buffer (pH = 7.0) in order to reach 1 wt.% in the final emulsion and was stirred for 2 h before being stored for 48 h at 4°C prior to usage.
<table><caption>**TABLE 1** Overview of the solvents used for the different emulsions and time points and the measurements performed at those time points.</caption><thead><tr><th>Emulsifier</th><th>Time points (days)</th><th>Used extraction solvent</th><th>NaCl</th><th>Measurements</th></tr></thead><tbody><tr><td>T20</td><td>All</td><td>HIP 3:1</td><td>-</td><td>Lipid extraction yield; NMR</td></tr><tr><td rowspan="2">WPI</td><td>All</td><td>HIP 3:1</td><td>-</td><td>Lipid extraction yield; NMR</td></tr><tr><td>0; 9/10</td><td>HIP/IOIP 3:2; Folch</td><td>+</td><td>Lipid extraction yield; NMR; colorimetric</td></tr><tr><td rowspan="2">sPPI</td><td>All</td><td>HIP 3:1</td><td>-</td><td>Lipid extraction yield; NMR</td></tr><tr><td>0; 9/10</td><td>HIP/IOIP 3:2; Folch</td><td>-</td><td>Lipid extraction yield; NMR; colorimetric</td></tr></tbody></table>

Note: Ratios v/v: 3:1/3:2.[3,8]

Abbreviations: Folch, chloroform-methanol; HIP, hexane-isopropanol; IOIP, isooctane-isopropanol; sPPI, "soluble" pea protein isolate fraction; T20, Tween20; WPI, whey protein isolate.

## 2.3 | Preparation of the emulsion

A coarse emulsion was prepared by mixing 10 wt.% stripped rapeseed oil with the aqueous phase (sPPI/WPI/T20, final emulsifier concentration in the emulsion 1 wt.%) using a high speed blender (S18N-19G, Ultra-turrax R, IKA-Werke GmbH & Co.) at 11 000 rpm, for 1 min. The coarse emulsion was then passed through a high pressure homogenizer (M-110Y Microfluidizer equipped with a F12Y interaction chamber, Microfluidics) at 400 bar, which was repeated to a total of five times to obtain the final emulsion. The coil of the system was cooled throughout homogenization by ice water to prevent heating up of the emulsion during preparation. The emulsions were stored in 20-mL vials (6 mL/vial) with 0.02 wt.% sodium azide against microbial growth and horizontally rotated in an oven at 40°C in the dark. Samples were taken at days 0, 1, 3, 6/7, and 9/10 and stored at -80°C till further analysis; at least two emulsions were prepared independently.

## 2.4 | Measurements

Table 1 summarizes all the measurements and solvents used for the different emulsions (T20, WPI, or sPPI) and time points considered.

### 2.4.1 | Lipid extraction yield

After thawing emulsion aliquots (0.3 mL) at room temperature (RT), lipids were extracted by adding 1.5 mL of a mixture of HIP (in a ratio of 3:1 v/v), solvent to emulsion ratio 5:1 v/v, as described by Shantha and Decker[10]. The emulsion and solvent were added to Eppendorf vials (2 mL), and the weight of both emulsion (before freezing) and solvent was noted. The mixtures were vortexed (Digital Vortex Mixer, Fisher Scientific) three times for 20 s and centrifuged (5424R, Eppendorf) at 4700 × g for 8 min. After centrifugation, the lipid-containing phase (hexane phase) was collected, added to another Eppendorf vial, and weighed. The Eppendorf vials were then placed in an extraction evaporator (Reacti-Therm, Thermo Scientific), flushing the tubes with nitrogen gas (25°C) for 1.5 h. The residual lipid mass was recorded and the lipid extraction yield was reported as the percentage of lipids present in the hexane phase compared to the total amount of oil present in the emulsion aliquot.

In addition, HIP was used in a ratio of 3:2 v/v, isopropanol-isooctane in a ratio of 3:2 v/v, and chloroform-methanol in a ratio of 2:1 v/v as extractions solvents. Furthermore, NaCl was added to the WPI-based emulsion before extraction with the different solvents to facilitate the extraction of lipids.[20] For all solvents and ratios, the procedure remained the same as described for the HIP 3:1 (Table 1), and samples were taken at representative points in time.

For each emulsion, all measurements were performed in triplicate for each solvent and time point.

### 2.4.2 | Lipid oxidation measurements

After extracting the lipids as described in Section 2.2, lipid oxidation was measured using the colorimetric method of Shantha and Decker and the ¹H NMR method of Merkx et al.[10,16]

**Colorimetric assay**

Briefly, 10 µL of the hexane or isooctane phase were mixed with 1.4 mL methanol-butanol (2:1 v/v) and 15 µL of assay reagent (Fe(SCN)₂). After 20 min incubation at RT, the absorbance was measured (510 nm) and the hydroperoxide concentration was calculated using a cumene peroxide calibration curve (0–160 µM). The measurements were performed in triplicate for each emulsion. The chloroform extracts were not analyzed by this method as chloroform gives a high absorbance at 510 nm (Figure S1) due to the enhanced formation of Fe³⁺ via trichloromethyl radicals.[21]

#### ¹H NMR

The dried lipid extract (see Section 2.2) was re-solubilized in a NMR-compatible solvent (deuterated chloroform-deuterated dimethylsulfoxide; 5:1 v/v) in a ratio of 3:1 v/v. The measurements were performed in duplicate for all samples (HIP 3:1 solvent).

For both lipid oxidation methods, the hydroperoxide concentration was expressed as mmol per kg of oil. In the case of the colorimetric method, it is common practice to assume that 100% of the lipids present in the sample are extracted to the nonpolar phase. In the present work, we checked whether this assumption is sound by comparing with the mass of extracted lipids. For the NMR method, the total lipid content can be determined by integration of the triglyceride signal (4.46–3.93 ppm) and using it as reference.
### 2.4.3 | Protein oxidation measurements

Protein oxidation in the sPPI- and WPI-based emulsions was determined by measuring the carbonyl content according to the DNPH method described by Levine et al. and Sante-Lhoutellier et al.[22,23] Proteins were first precipitated by mixing an emulsion aliquot with isopropanol (1:10) followed by a centrifugation step at 15 000 × g (5 min, RT) to obtain the protein pellet, which was next dispersed in either 500 µL of 10 mM DNPH in 2 N HCL or only in 2 N HCL (blank). After incubation for 60 min in the dark, proteins were precipitated again with 500 mL of 40 wt.% TCA solution (10 min on ice). The dispersion was centrifuged at 15 000 × g for 5 min at RT, and the pellet was washed with 1 mL of ethanol/ethyl acetate 1/1 v/v (2×), then once with 1 mL of 2-propanol, and finally dissolved in 1 mL of 6 M guanidine hydrochloride at 37°C. Centrifugation under the same conditions removed the insoluble matter, if any, and the absorbance of the supernatant was measured at 370 nm. A molar absorption coefficient of 22 000 M⁻¹ cm⁻¹ was used to calculate the protein-bound carbonyl content. The soluble protein content in the supernatant was determined by the BCA-assay using sPPI or bovine albumin standard (for WPI-based samples) as calibration curve.[24] The results were expressed in mmol carbonyl per kg soluble protein for two independent measurement replicates.

### 2.4.4 | Statistics

Standard deviations were calculated from triplicate measurements of the same emulsion (lipid extraction, lipid oxidation via calorimetric method) or from duplicate measurements (relative middle phase height, protein oxidation, and lipid oxidation via NMR method). Significant differences between samples were determined using IBM SPSS statistics software with one-way ANOVA and post hoc with the Tukey HSD method to compare means. Significance was established with *p* < 0.05.

## 3 | RESULTS

In the following sections, the results are described for the different solvents and emulsifiers. The average droplet sizes ($d_{3,2}$) varied between 0.07 and 0.30 µm and did not change over time, with WPI-emulsions showing the smallest ($d_{3,2}$: 0.07–0.08 µm) and PPI-emulsions the largest values ($d_{3,2}$: 0.20–0.30 µm) (see Table S1).

### 3.1 | Extraction system—opaque layer

For T20-stabilized emulsions, a clear phase separation was obtained upon extraction, whereas for protein-stabilized emulsions, an opaque layer (referred to hereafter as the middle layer) was observed between the upper (hexane/isooctane or methanol) and lower phases (isopropanol or chloroform). The thickness of the middle layer was similar

for all extraction methods and time points for the sPPI-emulsion (data not shown), whereas it varied in size per time point, extraction method, and even between samples of one-time point for the WPI-emulsion (Figure 1). Overall, the thickness of the middle layer was larger for the WPI-emulsions than for the sPPI-emulsions. Figure 1(ii) shows that the addition of NaCl decreased the thickness of the middle layer for all solvents used and the layer seemed more compact.

### 3.2 | Lipid extraction yield for different emulsifiers

Figure 2 shows the obtained yields for HIP (3:1 v/v) extraction from emulsions stabilized by T20 (a), WPI (b), or sPPI (c) over time. The extracted lipids represented >75 and >60 wt.% of the oil present in the emulsions, for the T20- and sPPI-based emulsions, respectively, with no differences between time points. In contrast, the lipid extraction yields for the WPI-stabilized emulsion were high in the fresh emulsion (around 70 wt.%) but then decreased to values as low as 26 wt.% on days 1 and 3 (Figure 2). At prolonged incubation, large variations were found (Figure 2, ii) indicating difficult lipid extraction from such aged systems.

### 3.3 | Effect of extraction method on lipid extraction yield

Figure 3 summarizes the lipid extraction yields obtained for emulsions stabilized by WPI or sPPI at day 0 or at the end of the incubation period (10 days) with different extraction methods, including the addition of a saturated NaCl solution (0.05 vol.% of total extraction system) (Figure 3, striped blue bars). For all solvents and emulsions, the lipid extraction yields were between 75 and 100 wt.%. The Folch 2:1 method led to significantly higher lipid extraction yields in all cases (75–86 vs. 95–100 wt.%). Similar lipid extraction yields were obtained with HIP 3:1, HIP 3:2, and IOIP 3:2 for both emulsions (WPI(+) and sPPI) and time points (0 and 10 days). The sPPI-emulsion generally showed a higher lipid extraction yield compared to the WPI-emulsion. In most cases, the addition of salt had no significant effect; only for the HIP solvent (3:2 v/v) the lipid extraction yield was significantly lowered, albeit the difference remained limited.

### 3.4 | Influence of the lipid extraction yields on the measured lipid oxidation levels

Figure 4 shows the measured hydroperoxide concentration in the protein-based emulsions for the different extraction methods and hydroperoxide assays. For all emulsifiers and solvents, the hydroperoxide concentration was the highest when measured with the ¹H NMR method, whereas it was substantially lower when measured with the colorimetric method. When these values were corrected for the actual lipid extraction yields (i.e., using the actual mass of extracted lipid instead of assuming complete extraction; see Section 2.4.2) the values
<figure><img src="image_4.png" /><figcaption>**FIGURE 1** Relative middle layer height of the whey protein isolate (WPI)-emulsion extracted (i) with hexane-isopropanol (HIP) solvent at different time points and (ii) at day 10 with the different solvents (HIP/isooctane-isopropanol (IOIP)/Folch), ratios (3:1/3:2), and with or without the addition of a saturated NaCl-solution (0.05 vol.% of total extraction system). The error bars show the standard deviations of two analyzed tubes of the same emulsion (except day 0 where only one tube was analyzed; see Figure S2).</figcaption></figure>

<figure><img src="image_5.png" /><figcaption>**FIGURE 2** Lipid extraction yields for emulsions stabilized by (i) Tween 20 (T20), (ii) whey protein isolate (WPI), or (iii) soluble pea protein (sPPI), as function of incubation time. In each panel, the different symbols correspond to independently prepared emulsions. The error bars denote the standard deviations of three independent measurements of a given emulsion.</figcaption></figure>
<figure><img src="image_6.png" /></figure>

**FIGURE 3** Lipid extraction yields for emulsions stabilized by whey protein isolate (WPI) (blue) and soluble pea protein (sPPI) (green) extracted with the different indicated solvents. For WPI-based systems, the extraction was conducted as such (plain blue bars) or with added NaCl (striped blue bars). Data for freshly prepared emulsion (panel (i)) or after 10 days of incubation (panel (ii)) are reported. The error bars represent the standard deviations (n ≥ 2) and significant differences are indicated with letters (p < 0.05; small letter: significance between WPI (with and without the addition of NaCl) and sPPI; capital letter: significance of solvent; apostrophe: significance between measured time points).

obtained with NMR for the sPPI-stabilized emulsions remained higher, whereas rather similar values to those determined by NMR method were obtained for the WPI-stabilized emulsions (see Section 4).

In addition, the aldehyde concentrations in the extracted lipids were measured with the NMR method after extraction with different solvents (Figure 5). The largest dependency on the used extraction solvent was found for the long *n*-alkanals (*n* > 5), of which the concentration varied significantly between 0.21–0.64 mmol/kg oil and 0.31–0.96 mmol/kg oil (IOIP < HIP 3:2 < HIP 3:1 < Folch) for the sPPI- and WPI-based emulsions, respectively. Folch extraction led to significantly higher aldehyde concentrations compared to the IOIP extraction

for both protein-stabilized emulsions and compared to HIP (3:1 or 3:2 ratio) for the sPPI-emulsion.

## 3.5 | Effect of lipid and protein oxidation on the lipid extraction yield

To investigate if lipid and protein oxidation are connected to the lipid extraction yield, we plotted both in Figure 6. For sPPI-based emulsions (Figure 6i), the lipid extraction yields were relatively high for all oxidation levels, whereas the lipid extraction yields for the WPI-based

<figure><img src="image_7.png" /></figure>

**FIGURE 4** Hydroperoxide concentration after 10 days of incubation of the soluble pea protein (sPPI) and whey protein isolate (WPI)-emulsions measured with the NMR (purple) or colorimetric (red) methods, with (hashed) or without (filled) correction for the lipid extraction yield for different solvents (hexane–isopropanol [HIP] 3:1 or 3:2, isooctane–isopropanol [IOIP] 3:2). The error bars represent the standard deviation from two (NMR) or three (colorimetric method) independent replicates. Capital letters indicate significant (p < 0.05) differences between the data obtained by the individual measurement methods after extraction with different solvents; small letters indicate per solvent and emulsifier significant (p < 0.05) differences between the NMR and colorimetric method (with and without correction).
<figure><img src="image_8.png" /><figcaption>**FIGURE 5** Concentrations of aldehyde classes (1: small *n*-alkanals (*n* < 3); 2: long *n*-alkanals (*n* > 3); 3: undefined (9.7–9.6 ppm); 4: hydro(pero)xy-alkenals; 5: dienals; 6: alkenals; 7: undefined (9.4–9.3 ppm); primary *y*-axis) and total aldehyde concentration (secondary *y*-axis) for the soluble pea protein (sPPI)-emulsion (i) and the whey protein isolate (WPI)-emulsion (ii) at day 10 extracted using Folch 2:1 (brown), hexane–isopropanol (HIP) 3:1 (yellow filled), HIP 3:2 (yellow striped), or isooctane–isopropanol (IOIP) 3:2 (orange).[16] The error bars represent the standard deviation of two NMR measurements. Letters indicate significant (*p* < 0.05) differences between the NMR measurements with different solvents.</figcaption></figure>

emulsions (Figure 6ii) were much more scattered. They were found to be high (60–80 wt.%) at low and, sporadically, very high protein oxidation levels, whereas they were noticeably low (24–55 wt.%) at moderate (protein) to high (lipid) oxidation levels.

## 4 | DISCUSSION

It is first clear that both the extraction solvent used and the analysis method lead to differences in the amount of oxidation products

measured (both hydroperoxides and aldehydes), which was herein exemplified for emulsions stabilized with a surfactant (T20) or protein ingredients (sPPI and WPI). Although the lipid extraction yield in T20-stabilized emulsions was not affected by the extraction solvent used, the situation was very different for protein-stabilized emulsions. There, the formation of a three-layered medium was observed upon extraction with a middle layer consisting of an organogel-like phase. This is most likely formed by a denatured protein network, which can possibly hamper the full extraction of lipids into the nonpolar phase of the system (in this study, chloroform, *n*-hexane, or isooctane).[14,25]

<figure><img src="image_9.png" /><figcaption>**FIGURE 6** Influence of protein oxidation expressed as carbonyls (blue filled symbols) and of lipid oxidation expressed as hydroperoxides (measured with ¹H NMR; green striped symbols) on the lipid extraction yield for three independently prepared soluble pea protein (sPPI)-(i) and WPI-(ii) emulsions. Errors denote standard deviations of two (carbonyls and hydroperoxides) or three (lipid yield) independent measurements of the same emulsion.</figcaption></figure>
reactions in heterogeneous food systems. *Journal of Food Science*, 65(8), 1270-1282. https://doi.org/10.1111/j.1365-2621.2000.tb10596.x

5. Laguerre, M., Bayrasy, C., Panya, A., Weiss, J., McClements, D. J., Lecomte, J., Decker, E. A., & Villeneuve, P. (2015). What makes good antioxidants in lipid-based systems? The next theories beyond the polar paradox. *Critical Reviews in Food Science and Nutrition*, 55(2), 183-201. https://doi.org/10.1080/10408398.2011.650335

6. Ghelichi, S., Hajfathalian, M., Yesiltas, B., Sørensen, A. D. M., García-Moreno, P. J., & Jacobsen, C. (2023). Oxidation and oxidative stability in emulsions. *Comprehensive Reviews in Food Science and Food Safety*, 22, 1864-1901. https://doi.org/10.1111/1541-4337.13134

7. Waraho, T., McClements, D. J., & Decker, E. A. (2011). Mechanisms of lipid oxidation in food dispersions. *Trends in Food Science & Technology*, 22, 3-13. https://doi.org/10.1016/j.tifs.2010.11.003

8. Hennebelle, M., Villeneuve, P., Durand, E., Lecomte, J., van Duynhoven, J., Meynier, A., Yesiltas, B., Jacobsen, C., & Berton-Carabin, C. (2024). Lipid oxidation in emulsions: New insights from the past two decades. *Progress in Lipid Research*, 94, 101275. https://doi.org/10.1016/j.plipres.2024.101275

9. Christie, W. W., & Han, X. (2010). Lipid extraction, storage and sample handling. In *Lipid analysis: Isolation, separation, identification and lipidomic analysis* (pp. 55–66). Woodhead Publishing.

10. Shantha, N. C., & Decker, E. A. (1994). Rapid, sensitive, iron-based spectrophotometric methods for determination of peroxide values of food lipids. *Journal of AOAC International*, 77(2), 421-424. https://doi.org/10.1093/jaoac/77.2.421

11. Donnelly, J. L., Decker, E. A., & McClements, D. J. (1998). Iron-catalyzed oxidation of menhaden oil as affected by emulsifiers. *Journal of Food Science*, 63(6), 997-1000. https://doi.org/10.1111/j.1365-2621.1998.tb15841.x

12. Mancuso, J. R., McClements, D. J., & Decker, E. A. (2000). Iron-accelerated cumene hydroperoxide decomposition in hexadecane and trilaurin emulsions. *Journal of Agricultural and Food Chemistry*, 48, 213-219. https://doi.org/10.1021/jf990757d

13. Tong, L. M., Sasaki, S., Julian, D., & Decker, E. A. (2000). Mechanisms of the antioxidant activity of a high molecular weight fraction of whey. *Journal of Agricultural and Food Chemistry*, 48, 1473-1478. https://doi.org/10.1021/jf991342v

14. ten Klooster, S., Boerkamp, V., Lazaridi, E., Yang, S., Takeuchi, M., Berton-Carabin, C., Schroën, K., Janssen, H.-G., Friedrich, H., Hohlbein, J., Duynhoven, J., & Hennebelle, M. (2022). Lipid oxidation in food emulsions: Analytical challenges and recent developments. In *Lipid oxidation in food and biological systems* (pp. 3-29). Springer.

15. Radin, N. S. (1988). Lipid extraction. In Bouton, A.a., Baker, G.B., Horrocks, L.A. (Eds.), *Lipids and Related Compounds. Neuromethods*, vol 7. Humana Press.

16. Merkx, D. W. H., Hong, G. T. S., Ermacora, A., & van Duynhoven, J. P. M. (2018). Rapid quantitative profiling of lipid oxidation products in a food emulsion by 1H NMR. *Analytical Chemistry*, 90(7), 4863-4870. https://doi.org/10.1021/acs.analchem.8b00380

17. Berton, C., Genot, C., & Ropers, M. H. (2011). Quantification of unadsorbed protein and surfactant emulsifiers in oil-in-water emulsions. *Journal of Colloid & Interface Science*, 354(2), 739-748. https://doi.org/10.1016/j.jcis.2010.11.055

18. Hoppenreijs, L. J. G., Berton-Carabin, C. C., Dubbelboer, A., & Hennebelle, M. (2021). Evaluation of oxygen partial pressure, temperature and stripping of antioxidants for accelerated shelf-life testing

of oil blends using 1H NMR. *Food Research International*, 147, 110555. https://doi.org/10.1016/j.foodres.2021.110555

19. Shea, F., & Watts, C. E. (1939). Dumas method for organic nitrogen. *Industrial and Engineering Chemistry—Analytical Edition*, 11(6), 333-334. https://doi.org/10.1021/ac50134a013

20. Folch, J., Lees, M., & Sloane Stanley, G. H. (1957). A simple method for the isolation and purification of total lipides from animal tissues. *Journal of Biological Chemistry*, 226(1), 497-509. https://doi.org/10.1016/S0021-9258(18)64849-5

21. Mihaljević, B., Katušin-Ražem, B., & Ražem, D. (1996). The reevaluation of the ferric thiocyanate assay for lipid hydroperoxides with special considerations of the mechanistic aspects of the response. *Free Radical Biology and Medicine*, 21(1), 53-63. https://doi.org/10.1016/0891-5849(95)02224-4

22. Levine, R. L., Garland, D., Oliver, C. N., Amici, A., Climent, I., Lenz, A. G., Ahn, B. W., Shaltiel, S., & Stadtman, E. R. (1990). Determination of carbonyl content in oxidatively modified proteins. *Methods in Enzymology*, 186(C), 464-478. https://doi.org/10.1016/0076-6879(90)86141-H

23. Sante-Lhoutellier, V., Aubry, L., & Gatellier, P. (2007). Effect of oxidation on in vitro digestibility of skeletal muscle myofibrillar proteins. *Journal of Agricultural and Food Chemistry*, 55(13), 5343-5348. https://doi.org/10.1021/jf070252k

24. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Frovenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., & Klenk, D. C. (1985). Measurement of protein using bicinchoninic acid. *Analytical Biochemistry*, 150, 76-85. https://doi.org/10.1016/0003-2697(85)90442-7

25. Radin, N. S. (1981). Extraction of tissue lipids with a solvent of low toxicity. *Methods in Enzymology*, 72(C), 5-7. https://doi.org/10.1016/s0076-6879(81)72003-2

26. Hara, A., & Radin, N. S. (1978). Lipid extraction of tissues with a low-toxicity solvent. *Analytical Biochemistry*, 90(1), 420-426. https://doi.org/10.1016/0003-2697(78)90046-5

27. Joshi, D. R., & Adhikari, N. (2019). An overview on common organic solvents and their toxicity. *Journal of Pharmaceutical Research International*, 28(3), 1-18. https://doi.org/10.9734/jpri/2019/v28i330203

28. Münch, K., Schroën, K., & Berton-Carabin, C. (2024). Relevance of various components present in plant protein ingredients for lipid oxidation in emulsions. *Journal of the American Oil Chemists' Society*, https://doi.org/10.1002/aocs.12790

## SUPPORTING INFORMATION

Additional supporting information can be found online in the Supporting Information section at the end of this article.

**How to cite this article:** Münch, K., ten Klooster, S., van Kouwen, I., Berton-Carabin, C., & Schroën, K. (2024). Incomplete lipid extraction as a possible cause for underestimation of lipid oxidation in emulsions. *European Journal of Lipid Science & Technology*, 126, e2300229. https://doi.org/10.1002/ejlt.202300229