# Faba bean and pea protein ingredients: Endogenous lipid accumulation and lipid oxidative state

Jolijn S.A. Koomen, Lucie Ribourg-Birault, Alice Kermarrec, Bérengère Marais, Sophie Le Gall, Adeline Boire, Anne Meynier, Claire Berton-Carabin

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## ► To cite this version:

Jolijn S.A. Koomen, Lucie Ribourg-Birault, Alice Kermarrec, Bérengère Marais, Sophie Le Gall, et al.. Faba bean and pea protein ingredients: Endogenous lipid accumulation and lipid oxidative state. Food Research International, 2025, 217, pp.116884. 10.1016/j.foodres.2025.116884. hal-05358075

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Submitted on 13 Nov 2025

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# Faba bean and pea protein ingredients: Endogenous lipid accumulation and lipid oxidative state

Jolijn S.A. Koomen<sup>a</sup>, Lucie Ribourg-Birault<sup>a</sup>, Alice Kermarrec<sup>a</sup>, Bérengère Marais<sup>a,b</sup>, Sophie Le Gall<sup>a,b</sup>, Adeline Boire<sup>a</sup>, Anne Meynier<sup>a</sup>, Claire Berton-Carabin<sup>a,c,*</sup>

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## ARTICLE INFO

**Keywords:**

Plant proteins

Pea

Faba bean

Endogenous lipids

Lipid oxidation

Hydroperoxides

## ABSTRACT

Pulses such as faba bean and pea are a promising source of proteins in the current protein transition. For their use in food products, they are processed into protein ingredients through dry or wet fractionation routes. The use of these ingredients is currently hampered by their inherent off-flavours, partly arising from lipid oxidation. In this study, both commercial and pilot-scale-produced faba bean and pea protein ingredients were used to determine their endogenous lipid profile, as well as the lipids' oxidative status. All protein ingredients contained considerable amounts of endogenous lipids compared to the raw material (4.4–6.6 g/100 g dry matter for concentrates and 7.4–7.5 g/100 g dry matter for isolates), and more than half these lipids were phospholipids. In addition, minor lipophilic compounds (tocopherols, carotenoids) were present in substantial amounts (240–2700 µg/g lipids) that varied both with the seed cultivar or growing conditions and with the seed-to-ingredient processing method. The endogenous lipids had undergone oxidation, as high and highly variable levels of hydroperoxides were found (2.7–57 µmol/g lipids). Yet, levels of secondary lipid oxidation markers were fairly low, and no strong occurrence of off-flavour associated volatiles, e.g., hexanal, was detected. Nonetheless, considering the high initial hydroperoxide concentration, it is expected that these off-flavours can quickly be formed upon hydroperoxide decomposition if these ingredients are applied in typical food matrices, such as plant-based dairy alternatives. These results substantiate the origin of lipid-derived off-flavours in pulse protein applications.

## 1. Introduction

Pulses are a promising source of proteins that can help partly shift human consumption from animal- to plant-derived food products. This shift is often referred to as the protein transition and has gained interest in recent years due to its proposed benefits to planetary, human, and animal health (Aiking & de Boer, 2020; Duluins & Baret, 2024). Pulses are legume seeds and therefore have an additional benefit compared to other alternative proteins, as legumes can fixate atmospheric nitrogen. Their cultivation can therefore aid in repairing the current disordered nitrogen cycle, as well as decrease the need for fertilisers (Herridge et al., 2008; Köpke & Nemecek, 2010).

Faba bean (*Vicia faba* L.) and yellow pea (*Pisum sativum* L.) seeds are starch-rich pulses that together represent about one-third of the European legume seed production (van Loon et al., 2023). For human consumption, faba bean and pea can be harvested as fresh or dry seeds. The

dry seeds, so-called pulses, contain approximately 20–30 wt% protein, 60–65 wt% carbohydrates, and 1–3 wt% lipids (Hall et al., 2017; Singh, 2017). In processed food products such as plant-based beverages and dairy and meat analogues, ingredients derived from dried seeds are used (Goldstein & Reifen, 2022). These ingredients are produced using different fractionation techniques, which aim to increase the protein content by removing non-protein components such as starch (Boye et al., 2010; Eze et al., 2022). First, the seeds are dehulled and ground into a flour. Then, one of two main fractionation routes, called wet and dry fractionation, is typically applied. Different wet fractionation methods exist, but most often proteins are extracted in alkaline conditions and recovered through an isoelectric precipitation step (Boukid et al., 2021; Lam et al., 2018). The recovered protein suspension then needs to be (spray-)dried to obtain a protein isolate that generally contains more than 70 wt% proteins (Boye et al., 2010). The pH adjustment, heating, and drying steps can induce protein denaturation and aggregation,

<footnote><sup>a</sup> INRAE, UR BIA, F-44300, Nantes, France</footnote>

<footnote><sup>b</sup> INRAE, PROBE/CALIS Research Infrastructures, BIBS Facility, F-44300, Nantes, France</footnote>

<footnote><sup>c</sup> Wageningen University & Research, Laboratory of Food Process Engineering, 6700, AA, Wageningen, the Netherlands</footnote>

<footnote>* Corresponding author at: INRAE, UR BIA, F-44300, Nantes, France.</footnote>

<footnote>E-mail address: claire.berton-carabin@inrae.fr (C. Berton-Carabin).</footnote>
leading to a loss of protein functionality in the protein isolate (Geerts et al., 2017; Möller et al., 2022; Yang et al., 2024). In addition, the use of water and energy-consuming drying steps is currently challenged due to environmental concerns. Therefore, dry fractionation has been gaining in interest as a more sustainable process compared to wet fractionation, although it produces an ingredient with lower protein purity (Geerts et al., 2018; Schutyser et al., 2015). Dry fractionation generally relies on a difference in size and density between the protein-rich particles and starch granules (Assatory et al., 2019; Pelgrom et al., 2013; Schutyser & van der Goot, 2011). Accordingly, so-called air classification can be used to separate a fine fraction, enriched in protein, from a coarse fraction, enriched in starch (Jones et al., 1959). The fine fraction is a protein concentrate and contains 45–70 wt% proteins (Arntfield & Maskus, 2011).

Previous research has shown that pulse protein concentrates and isolates can have promising properties to stabilise food matrices, such as emulsions (Felix et al., 2018; Karaca et al., 2011; Krause et al., 2023; Stone et al., 2015; Vogelsang-O'Dwyer et al., 2021). However, their current use in commercial food products is still limited. This is mainly due to their inherent off-flavours (Roland et al., 2017; Sharan et al., 2021). It has been suggested that the involved volatile off-flavour molecules would, for a large part, originate from lipid oxidation (Jiang et al., 2016; Karolkowski et al., 2023; Murat et al., 2013; Rajhi et al., 2022; Sessa & Rackis, 1977). Besides lipid oxidation, amino acid degradation, Maillard reactions, and polyphenols are also responsible for the formation of off-flavours in such ingredients (Cosson et al., 2021; Roland et al., 2017; Trikusuma et al., 2020). In pulses, lipid oxidation is suggested to occur mostly enzymatically, as lipoxygenases are present (Baysal & Demirdöven, 2007; Yang et al., 2017). In addition, autoxidation can occur through free radical chain reactions involving unsaturated fatty acids and oxygen (Frankel, 1980). Both pathways lead to the formation of hydroperoxides, which are primary lipid oxidation products. The decomposition of hydroperoxides, catalysed notably by pro-oxidant metals, leads to the formation of a diverse range of secondary lipid oxidation products (Schaich, 2005). Some of these secondary lipid oxidation products, such as hexanal and 2-nonenal, are volatile and cause the previously mentioned negative flavour attributes (Eriksson, 1975; Frankel, 1983; Jacobsen, 1999).

The lipid content in pulse flours is relatively low at 1–3 wt%, but previous authors have found it to be sufficient to produce noticeable levels of volatile lipid oxidation products (Jiang et al., 2016; Xu et al., 2019). Recent research has shown that endogenous pulse lipids accumulate during the protein fractionation process (Keuleyan et al., 2023). A commercial pea protein concentrate was found to contain 6 g lipids per 100 g of dry matter (DM), and an even higher content of almost 12 g lipids per 100 g DM was found in a pea protein isolate. The volatile flavour profile of pulse protein ingredients has been investigated before, however, the lipid profile at the origin of this flavour development was not characterised (Liu et al., 2023; Sharan et al., 2022). A characterisation of both the composition, as well as the oxidative state of these endogenous lipids, is key to better understand the origin of off-flavour molecules.

The aim of this work was therefore to characterise in depth the endogenous lipids, as well as their oxidative status, in faba bean and pea protein ingredients obtained from different cultivars and processes. The characterisation of lipids included both major (lipid classes and fatty acid composition) and minor components (tocopherols and lipophilic pigments), as well as several lipid oxidation products, completed by the analysis of headspace volatile compounds. Both commercially available and pilot-scale-produced protein ingredients were used in this study. The commercial ingredients were chosen as they are most relevant for food applications, whereas the pilot-scale-produced ingredients allowed to look into the effect of seed cultivar on the endogenous lipid profile and oxidative state.

# 2. Materials and methods

## 2.1. Chemicals

Chloroform, ethanol, n-hexane, isopropanol, methanol, and methyl-tert-butyl ether were purchased from Biosolve (Dieuze, France). Ferrous ammonium sulfate, heptadecanoic acid (C17:0), and hydranal-composite 5 were obtained from Fluka Analytical (Honeywell, Saint-Germain-en-Laye, France). Acetonitrile, ammonia solution (30 %), anhydrous sodium sulfate, cyclohexane, diethyl ether, and toluene were acquired from Carlo Erba Reagents (Val de Reuil, France). 2-Mercaptoethanol, boron trifluoride (BF₃) in methanol (14 w/v%), butylated hydroxytoluene (BHT), ethyl acetate, ethylenediaminetetraacetic acid (EDTA), hydrogen peroxide (H₂O₂), neutral lipid standards (FA 16:0; MAG 16:0; 1,2-DAG 16:0; 1,3-DAG 16:0; and TAG 16:0), phosphoric acid (85 wt%), tetraethoxypropane (TEP), thiobarbituric acid (TBA), trichloroacetic acid (TCA), and xylenol orange were purchased from Sigma Aldrich (Merck KGaD, Darmstadt, Germany). Bolt™ MES SDS running buffer, Bolt™ Bis-Tris plus gels (4–12 % gradient), and Novex Sharp pre-stained protein standard were obtained from Invitrogen (Thermo Fisher Scientific, Waltham, USA). Sodium dodecyl sulfate (SDS) solution (20 %) and tris(hydroxymethyl)aminoethane were bought from Euromedex (Souffelweyersheim, France). Ammonium sulfate was from VWR (Radnor, USA), glycerol was from Acros Organics (Thermo Fisher Scientific), sodium chloride was from Fisher Scientific (Thermo Fisher Scientific), and sulfuric acid (96 %) was from PanReac (ITW Reagents, Monza, Italy). As standards, a fatty acid methyl ester (FAME) mix C4-C24 was obtained from Supelco (Merck KGaD), phospholipid standards (phosphatidic acid, phosphatidylcholine, phosphatidylethanolamine, and phosphatidylinositol) from Avanti (London, UK), precision plus protein unstained standard from Bio-rad (Marnes-la-Coquette, France), and α/β/γ/δ tocopherol set from Calbiochem (Merck KGaD). Lastly, bromophenol blue sodium salt and Coomassie brilliant blue G250 were purchased from Serva (Serva Electrophoresis GmbH, Heidelberg, Germany). All the chemicals and reagents used were of analytical grade. Ultrapure water obtained with a MilliQ device (Millipore, Merck KGaD) was used for all the experiments.

## 2.2. Samples

Commercial samples of faba bean and pea protein ingredients were kindly provided by ingredient suppliers and are indicated with 'C'

<caption>**Table 1**</caption>

Protein ingredient samples and their abbreviation. After sample reception, the protein ingredients were stored in the dark at room temperature and a<sub>w</sub> 0.438. NA = information not available.

<table><thead><tr><th>Seed type</th><th>Ingredient type</th><th>Abbreviation</th><th>Seed variety</th><th>Production</th></tr></thead><tbody><tr><td rowspan="10">Faba bean</td><td rowspan="4">Commercial protein concentrate</td><td>FPC_C1</td><td>NA</td><td>Nov 2023</td></tr><tr><td>FPC_C2</td><td>NA</td><td>Aug 2022</td></tr><tr><td>FPC_C3</td><td>NA</td><td>Sep 2023</td></tr><tr><td>FPC_P1</td><td>Nairobi</td><td>Nov 2023</td></tr><tr><td rowspan="4">Pilot-scale protein concentrate</td><td>FPC_P2</td><td>Diva</td><td>Nov 2023</td></tr><tr><td>FPC_P3</td><td>Navara</td><td>Nov 2023</td></tr><tr><td>FPC_P4</td><td>Navi</td><td>Nov 2023</td></tr><tr><td>FPC_P5</td><td>Victus</td><td>Nov 2023</td></tr><tr><td rowspan="2">Commercial protein isolate</td><td>FPI_C1</td><td>Trumpet</td><td>May 2023</td></tr><tr><td>Pilot-scale protein isolate</td><td>FPI_P1</td><td>Nairobi</td><td>Nov 2023</td></tr><tr><td rowspan="4">Pea</td><td rowspan="2">Commercial protein concentrate</td><td>PPC_C1</td><td>NA</td><td>Nov 2023</td></tr><tr><td>PPC_C2</td><td>NA</td><td>Oct 2022</td></tr><tr><td rowspan="2">Pilot-scale protein concentrate</td><td>PPC_C3</td><td>NA</td><td>Oct 2023</td></tr><tr><td>PPC_P1</td><td>Furious</td><td>Nov 2023</td></tr></tbody></table>
(Table 1). All commercial protein concentrates came from the same supplier, but differed in production year or production process. C1 and C3 ingredients were produced in 2023, and C2 in 2022. An extra heat treatment in dry state was applied to the C3 protein concentrates, after which the ingredient was ground. In addition to the commercial ingredients, pilot-scale fractionation was applied by IMPROVE (Dury, France) on faba bean and pea seeds of different varieties. This yielded protein concentrates, FPC and PPC, and one faba bean protein isolate, FPI_P1, indicated with the letter 'P' (Table 1). The faba bean and pea protein concentrates were prepared from different varieties of seeds obtained from Agri Obtentions (Guyancourt, France) (FPC_P1 to FPC_P4 and PPC_P1) and RAGT (Rodez, France) (FPC_P5).

## 2.3. Compositional analyses

### 2.3.1. Moisture content

Equilibration at $a_W$ 0.438 was done at room temperature in absence of direct light in a desiccator over a saturated $K_2CO_3$ solution for at least 2 weeks, and equilibrated ingredients were stored (less than 1 year) at this $a_W$ in the dark at room temperature. This storage period was found to have no significant influence on the oxidative state of the ingredients (data not shown). The moisture content of the equilibrated powders was measured in independent triplicates using a Karl Fisher apparatus (Volumetric KF Titrator V30S, Mettler-Toledo, Viroflay, France) equipped with a sampler tray and oven at 105 °C (Stromboli, Mettler-Toledo). The equilibration step led to a relatively stable moisture content for all samples of 8–10 g per 100 g (Supplementary information 2, Table S2.1 and S2.2), in contrast to larger differences seen before equilibration (4–10 g per 100 g). In addition, the equilibration ensured a stable dry matter content during all subsequent analyses.

### 2.3.2. Protein content and composition

The protein content of equilibrated powders was calculated from their nitrogen content analysed by solid Dumas analysis in a CNS elemental analyser (Vario Micro Cube, Elementar, Langenselbold, Germany). The PPC protein content was calculated using a nitrogen-to-protein conversion factor of 5.6 (Keuleyan et al., 2023) The nitrogen-to-protein conversion factor for FPC and FPI samples were derived from their amino acid composition (analysed by an external lab by UHPLC-MS after acid and oxidative hydrolysis, Supplementary information 1 Table S1.1) and were equal to 5.4 and 5.5, respectively (Supplementary information 1 Table S1.2). All compositional analyses were done in independent triplicates and expressed in g per 100 g dry matter (DM) as the average value ± standard deviation.

In addition to the protein content, the protein composition was determined through SDS-PAGE (Laemmli, 1970), which may notably point to the presence of enzymes associated with lipid oxidation. Samples (2 mg protein) were added to 1 mL of Laemmli buffer $1 \times (0.0625 \text{ M Tris, } 2 \% \text{ SDS, } 10 \% \text{ glycerol, } 0.005 \% \text{ bromophenol blue})$ and head-over-tail rotated overnight. The following day, 100 µL of the hydrated ingredient were mixed with 100 µL of reducing Laemmli buffer $2 \times (0.125 \text{ M Tris, } 4 \% \text{ SDS, } 20 \% \text{ glycerol, } 0.01 \% \text{ bromophenol blue, } 16 \% 2-mercaptoethanol)$ and boiled in water for 5 min. Samples (8 µL) were loaded in the wells of 4–12 % Bis-tris Plus gels (Bolt™, Invitrogen). Precision plus protein unstained standard (Bio-rad) and Novex Sharp pre-stained protein standard (Invitrogen) were applied to each gel in a separate well (5 µL) as molecular weight marker. The gels were mounted in a Life Technologies Mini Gel Tank (Invitrogen) and submerged in MES SDS running buffer (Bolt™, Invitrogen). The sample migration was done at 200 V according to the manufacturer guidelines. Afterwards, the gels were removed and washed twice for about 1 h in a 50 % ethanol and 2 % phosphoric acid solution. The third washing step was done in 2 % phosphoric acid for 1 h. Then, 50 mL of sensibilisation solution (17 % ethanol, 15 % ammonium sulfate, 2 % phosphoric acid) were added to each gel. After 20 min, 100 mg of Coomassie brilliant blue (G250, Serva) were added and the gels were left to colour overnight. The next day, the

gels were washed with water until the background staining was removed. The gels were scanned (ImageScanner III, GE Healthcare, Freiburg, Germany) and the obtained image was processed with Image Lab (V6.1, Bio-rad) to determine the molecular weight of each band.

### 2.3.3. Analyses of lipids and lipophilic compounds

#### 2.3.3.1. Lipid content.
The lipid content of equilibrated powders was determined gravimetrically after lipid extraction. The Folch lipid extraction method (Folch et al., 1957) was chosen as it was found to have the highest pulse lipid recovery (Caprioli et al., 2016). The Folch protocol used a mixture of chloroform-methanol and was adapted as previously described (Keuleyan et al., 2023). In addition to their adaptations, the initial sampling size was increased to 2 g powder, and the volume of extracting solvent was adapted to maintain a 1:20 (w/v) sample-to-solvent ratio. The round flask containing the lipid extract was covered with aluminium foil during drying under nitrogen to prevent photo-oxidation. Lipid extracts were stored in the dark at -80 °C until further analysis of lipid extracts.

#### 2.3.3.2. Lipid classes.
The different types of neutral lipids (triacylglycerides (TAG), diacylglycerides (DAG), monoacylglycerides (MAG), and free fatty acids (FFA)) and phospholipids (PL) present in the lipid extracts were analysed using normal-phase high-performance liquid chromatography (HPLC) coupled to an evaporative light scattering detector (ELSD). Two independent injections were executed on each lipid extract to quantify neutral lipids, analysed at 0.5 mg L$^{-1}$ chloroform, and PL, analysed at 1 mg mL$^{-1}$ chloroform, separately. For both series, 30 µL of the prepared lipid extract were injected into an UltiMate® 3000 RS-HPLC System (Thermo Scientific Dionex, France) equipped with an Uptisphere® CS Evolution (2.6 µm, 150 mm × 4.6 mm, Interchim, Montluçon, France). Eluent A was $CHCl_3$ and eluent B a mixture of $CH_3OH/CHCl_3/NH_4OH$ (30 %) (92/1/7, v/v). The elution profile used was a multistep binary linear gradient: 0 min 0 % B; 3 min 20 % B; 6 min 50 % B; 6.1 min 100 % B; and isocratic on 100 % B for 1 min for neutral lipids or 0 min 0 % B; 4 min 25 % B; 6 min 30 % B; 14 min 100 % B; and isocratic on 100 % B for 2 min for PL. The flow rate was 1.5 mL min$^{-1}$. The column was equilibrated with 100 % A for 3 min in between injections. Detection of lipid classes was carried out by an ELSD (SEDEX LT-ELSD 85LT, Sedere, Alfortville, France). Air was used as nebulizer gas (pressure 3.5 bars, drift tube temperature 40 °C) and the photomultiplier sensitivity was set to a gain value of 6. The Chromeleon Chromatography Management software (7.2.10 ES, Thermo Scientific) was used to control the system and process the data. Lipid class quantification was performed using calibration curves made with pure standards (quadratic fit).

#### 2.3.3.3. Fatty acid composition.
The fatty acid (FA) composition of lipid extracts was investigated after acid-catalysed methylation to fatty acid methyl esters (FAME) as described by Morrison and Smith (1964) and adapted by Viau and Meynier (2024b). The formed FAMEs were separated with gas chromatography (GC Clarus 690, Perkin Elmer) and detected by a flame ionisation detector (FID) (Viau & Meynier, 2024a). The GC capillary column (DB225 J&W Scientific, 0.25 µm, 30 m × 0.32 mm, Avantor, VWR) had a stationary phase of 50 % cyanopropylphenyl and 50 % methylpolysiloxane. The Chromeleon Chromatography Management software (7.2.10 ES, Thermo Scientific) was used to control the system and process the data. Identification was done by comparing the analyte retention time with that of a standard mixture (FAME mix C4-C24 Supelco 18,919-1AMP). Quantification was performed using the internal standard (C17:0) and applying correction factors depending on the FAMEs response (C18:0 taken as reference). Each lipid extract was analysed in duplicate (total $n = 6$) and the results were expressed as FA proportion ± standard deviation. In addition to the results on the total lipid fraction, FA composition was similarly assessed on two
fractionated lipid extracts (Supplementary information 4) prepared as previously described (Viau et al., 2024).

### 2.3.3.4. Tocopherol content.
Normal phase HPLC separation combined with a fluorescence detector was used to determine the tocopherol type and content in lipid extracts solubilised in hexane (Ribourg-Birault & Meynier, 2024b). An UltiMate® 3000 RS HPLC System (Thermo Scientific Dionex) with a normal phase column (Polar Advantage II, 3 µm, 25 cm × 3 mm, Thermo Scientific) was used for the analysis. The Chromleon Chromatography Management software (7.2.10 ES, Thermo Scientific) was used to control the system and process the data. Tocopherol quantification was performed using calibration curves made with pure standards (linear fit). Each lipid extract was analysed once (total n = 3). The results were expressed as average concentration in µg tocopherol per g lipid ± standard deviation.

### 2.3.3.5. Lipophilic pigments.
The UV-visible absorbance of lipid extracts adjusted to 5 mg mL⁻¹ diethyl ether between 200 and 700 nm (1 nm interval) were recorded with a UV-visible spectrophotometer (UV-1800, Shimadzu). Diethyl ether was used as a blank. UVProbe (version 2.33, Shimadzu) was used to control the spectrophotometer and process the data. The obtained spectra were corrected for the turbidity contribution (Winder & Gent, 1971). Each lipid extract was analysed once (total n = 3), and one representative absorbance spectrum is shown in Fig. 6 or Supplementary information 5 (Figs. S5.1, S5.2, and S5.3).

### 2.3.4. Other components

Additional non-lipid compounds were characterised in equilibrated powders to obtain a full characterisation of the protein ingredients. The ash content was determined gravimetrically after oven carbonisation at 550 °C. Cell wall polysaccharides, starch, saccharose, and raffinose family oligosaccharides were quantified using methods previously described by Lahaye et al. (2020), Le Gall et al. (2016), and Gangola et al. (2014). The sum of these components was taken as total carbohydrates for protein concentrates, for protein isolates, only cell wall polysaccharides were quantified. In addition, the iron and copper content were quantified by an external lab through plasma atomic emission spectroscopy and expressed as the measured value ± standard error of the measurement.

## 2.4. Lipid oxidation measurements

### 2.4.1. Hydroperoxides

Hydroperoxides (HPX) were quantified by the ferrous ion xylenol orange (FOX2) assay (Ribourg-Birault & Genot, 2024). The absorbance spectrum of the reaction medium was recorded between 450 and 650 nm (1 nm interval) (UV-1800, Shimadzu). UVProbe (V2.33, Shimadzu) was used to control the spectrophotometer and process the data. Each lipid extract was analysed once (total n = 3). The results were expressed as average H₂O₂ equivalent concentration in µmol per g lipids and per g of DM powder ingredient ± standard deviation. As an additional marker of primary lipid oxidation products, conjugated dienes were quantified (Supplementary information 6) as previously detailed (Ribourg-Birault et al., 2024).

### 2.4.2. Thiobarbituric acid reactive substances

Secondary lipid oxidation products were analysed as reported by Ribourg-Birault and Meynier (2024a). On the one hand, all thio-barbituric acid reactive substances (TBARS) were quantified spectro-photometrically through a colorimetric analysis. On the other hand, malondialdehyde (MDA) – TBA complexes specifically were separated on an inverse phase C18 HPLC column (Synchronis C18, 5 µm, 150 × 4.6 mm, Thermo Scientific) and detected with a diode array detector at 532 nm (UltiMate® 3000 RS HPLC system, Thermo Scientific Dionex). Each lipid extract was analysed once (total n = 3). The results were

expressed as average MDA equivalent concentration in nmol per g lipids ± standard deviation.

### 2.4.3. Volatile compounds

Volatile secondary lipid oxidation products were analysed with solid-phase microextraction (SPME) coupled to GC with mass spectrometry (MS) and FID as reviewed by Jacobsen (2024). The samples (1 g) were placed in a hermetically sealed vial of 22.4 mL. Two different incubation protocols were used. The vials were incubated at 35 °C for 5 min or 50 °C for 15 min, after which the volatile compounds present in the headspace were adsorbed onto a SPME fibre (TriPlus RSH system, Thermo Fisher Scientific). The SPME fibre (carbon WR/PDMS, Thermo Fisher Scientific) was exposed for 15 min at 35 °C or for 30 min at 50 °C (penetration depth 4 cm) and desorbed for 5 min in the GC (Trace 1310, Thermo Fisher Scientific) injector (splitless, 250 °C). The following temperature gradient was used: 0–5 min at 35 °C; raised at 10 °C min⁻¹ up to 210 °C; raised at 40 °C min⁻¹ up to 240 °C and held at 240 °C for 4 min. The GC was coupled to an FID detector (225 °C) fed by air and hydrogen flows of 350 and 35 mL min⁻¹, respectively. In addition, an MS detector was coupled for identification of volatile compounds (ISQ 7000 Single Quadrupole, Thermo Fisher Scientific). Mass spectra were obtained with 70 eV electron impact ionisation (ion source 250 °C) for m/z 25–250. The obtained spectra were compared with the NIST library to identify the volatile compounds. The Chromleon Chromatography Management software (7.2.10 ES, Thermo Scientific) was used to control the system and process the data. Each sample was analysed in independent duplicates and one representative chromatogram is shown in Fig. 9.

## 2.5. Statistical analysis

Statistical analysis of the data was performed in RStudio (V4.3.2, Boston, USA) using a multifactorial analysis of variance (ANOVA) with a Tukey post-hoc test. Significant differences (*P* < 0.05) were indicated using different letters. In addition, a Spearman correlation test was used to analyse the correlation between different variables.

# 3. Results and discussion

## 3.1. Composition of the samples

### 3.1.1. Lipid accumulation

Previous work on pulse protein ingredients has shown that endogenous lipids accumulate during protein fractionation, even for pulses that are relatively poor in lipids (Gueguen, 1983; Keuleyan et al., 2023). The Folch extraction method was chosen to quantify these lipids, since this method allowed for the fullest lipid recovery in legume seeds (Caprioli et al., 2016). The concentration of endogenous lipids in pulse protein ingredients was confirmed by the high lipid contents: FPCs contained 4.4–5.6, FPIs 7.4–7.5, and PPCs 5.4–6.6 g per 100 g DM (Fig. 1). Both the seed species and the applied fractionation process were found to significantly influence the lipid content. The FPC and PPC lipid contents were similar to those reported by Felix et al. (2018) (FPC at 4.8 g per 100 g), Karolkowski et al. (2023) (FPC at 4.6–4.9 g per 100 g), and Shand et al. (2007) (PPC at 5.8 g per 100 g). However, most studies still report a much lower lipid content of similar protein ingredients (Karaca et al., 2011; Vioque et al., 2012). This is most likely due to the lipid extraction method employed to assess the lipid content. The Soxhlet method is predominantly used in literature, which does not allow for a full recovery of especially polar lipids (Xiao et al., 2012). Even though the total lipid content of pulse protein ingredients is not comparable due to the difference in extraction methods, previous research confirmed the accumulation of endogenous lipids in dry-fractionated pulse protein concentrates (Bhatty & Christison, 1984; Z. Li et al., 2024; Rempel et al., 2019; Skylas et al., 2023; Sosulski & McCurdy, 1987). Funke et al. (2022) found a correlation between protein and lipid content during dry fractionation of lentils, which was ascribed to the association of protein
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Proximate composition of faba bean and pea protein ingredients, expressed relative to dry matter (DM). The protein content was calculated with a nitrogen-to-protein conversion factor of 5.4 for faba bean concentrates (FPC), 5.5 for faba bean isolates (FPI), and 5.6 for pea protein concentrates (PPC). Carbohydrates includes cell wall polysaccharides, starch, saccharose, and raffinose family oligosaccharides for protein concentrates and cell wall polysaccharides for protein isolates. The values are displayed as the average of three independent replicates ± standard deviation. Data used to create the figure are given in Supplementary information 2. FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>

bodies and small oil bodies. Nonetheless, other studies did not report a further lipid accumulation upon wet fractionation (Z. Li et al., 2024; Sosulski & McCurdy, 1987). This could be explained by the fact that lipids can be trapped inside more complex structures in pulse protein isolates (Keuleyan et al., 2023), and therefore their extractability with the Soxhlet method is decreased compared to pulse flours and protein concentrates.

### 3.1.2. Non-lipid components

In addition to the lipid content, the complete proximate composition of the protein ingredients was determined (Supplementary information 2). Several non-lipid components can affect the lipid oxidative state, such as lipid-modifying enzymes (as detailed later) in the protein fraction and pro-oxidant metal ions in the ash fraction. Iron and copper were quantified in all ingredients and were present in noticeable amounts in faba bean concentrates (51–72 mg iron and 19–33 mg copper per kg DM), isolates (136–142 mg iron and 24–38 mg copper per kg DM), and pea protein concentrates (63–91 mg iron and 15–19 mg copper per kg DM).

Protein was the main constituent in all analysed ingredients (Fig. 1). Using nitrogen-to-protein conversion factors derived from the ingredients' amino acid composition, the protein content of FPCs and PPCs was 55–61 and 48–59 g per 100 g DM, respectively (Supplementary information 1). The commercial FPCs had a higher protein content compared to commercial PPCs. This could be explained by the higher protein content in faba bean seeds and the larger faba bean starch granules, facilitating dry fractionation through air classification (Gueguen & Cerletti, 1994; Hall et al., 2017; Schutyser et al., 2015). All pilot-scale-produced protein concentrates (faba bean FPC P1 to FPC P5, as well as pea PPC P1) had a similar protein content around 60 g per 100 g DM. This is explained by the fact that the applied air classification parameters were optimised for each seed cultivar to reach this target value. The FPIs had a higher protein content at 78–81 g per 100 g DM, as expected for a wet fractionated ingredient (Z. Li et al., 2024). The protein composition was analysed through SDS-PAGE analysis (Fig. 2) to identify lipid-modifying enzymes, as faba bean or pea lipase, phospholipase, lipoxygenase (LOX), and hydroperoxide lyase (HPL) activities have been previously reported (Atwal et al., 1979; Hornostaj & Robinson, 2000; Lampi et al., 2020; Yang et al., 2017). Lipase is capable of cleaving ester bonds and releasing free fatty acids (FFA) from glycerol-bound neutral lipid species, and has been described to have a molecular weight of around 210 kDa in faba bean (Dundas et al., 1978). Weak

<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Protein profile of faba bean (A) and pea (B) protein ingredients on SDS-PAGE gels (4–12 % gradient) under reducing conditions. Marker = Novex Sharp prestained protein standard (Invitrogen, A) and precision plus protein unstained protein standard (Bio-rad, B). LOX = lipoxygenase, HPL = hydroperoxide lyase, FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>
bands were seen in this region on the electrophoresis gels, but their intensity was too low to estimate an exact molar mass. A faint band was observed slightly above 100 kDa for faba bean samples, which could correspond to a LOX of 106 kDa, previously reported for faba bean (Warsame et al., 2020). LOXs are enzymes responsible for the formation of hydroperoxides (HPX) from unsaturated fatty acids during enzymatic oxidation. In addition, a clear band around 95 kDa was seen in all faba bean protein ingredients, attributed to a second LOX enzyme, found to have a molecular mass of 96–97 kDa (Clemente et al., 2000; Warsame et al., 2020). In pea protein ingredients, a corresponding band at 94 kDa was seen. It has previously been reported that pea LOX has a slightly lower molecular weight corresponding to faba bean LOX (Szymanowska et al., 2009). A band around 70 kDa was observed in all samples, which could correspond to a heat shock protein as found by Warsame et al. (2020) but could also be a phospholipase previously reported in faba bean leaves by Kyong Kim et al. (1994). Similar to lipase, phospholipase is able to cleave the ester bond present in phospholipids to release FFA. The last enzyme, hydroperoxide lyase, has been purified from pea seeds in previous research and catalysed the decomposition of HPX into secondary lipid oxidation products (Hornostaj & Robinson, 2000). A band possibly corresponding to this enzyme was seen around 55 kDa. In addition to these specific bands, bands of storage proteins were annotated as described in previous research (Horstmann & Müntz, 1986; Keivananahr et al., 2021; Nikolić et al., 2012; Warsame et al., 2018, 2020). These bands were very similar among samples of same seed species, showing the conservation of major proteins between cultivars (Singhal et al., 2016).

### 3.1.3. Lipid composition

The low recovery of the Soxhlet method for polar lipids is especially problematic in pulse samples, as phospholipids have been found to make up an important portion of the total lipids present in legume seeds and their derived protein ingredients (Coxon & Wright, 1985; Gueguen, 1983; Haydar & Hadziyev, 1973; Keuleyan et al., 2023; Miyazawa et al., 1975; Yoshida et al., 2008). HPLC-ELSD analysis was employed to quantify the lipid classes present in the lipid extracts. The majority of lipids were found to be phospholipids, representing 55–77 % of the identified lipid classes (Fig. 3). The highest phospholipid content of 77 % was found in the pilot-scale-produced pea protein concentrate, PPC_P1. The main phospholipids were phosphatidylcholine (PC) and

phosphatidylinositol (PI), and in some samples phosphatidic acid (PA). Yoshida et al. (2008) confirmed that PC is the main phospholipid in faba bean seeds, however, they found a more important contribution of phosphatidylethanolamine (PE) than PI. Conversely, in the present work, PE only accounted for 6–18 % of all phospholipids. PA represented 7–19 % of total lipid classes in protein concentrates, but was remarkably decreased to 2–3 % in the lipid extracts of both faba bean protein isolates (Fig. 3). FPI_P1 was made from the same seed cultivar and batch as FPC_P1 – both starting from the same faba bean flour – from which could be concluded that this decrease was caused by the applied fractionation process. PA had the lowest retention factor (R<sub>F</sub> value) in the complementary high-performance thin layer chromatography (HPTLC) analysis (Supplementary information 3 Fig. S3.2), showing its relatively higher polarity compared to other phospholipids. This higher polarity is expected to have led to a partial solubilisation of PA during wet fractionation, leading to its lower recovery in the protein isolates. Similarly, Keuleyan et al. (2023) found a lower PA content in a commercial pea protein isolate at 0.2 g per 100 g DM, compared to a commercial pea protein concentrate at 0.4 g per 100 g DM.

Besides phospholipids, neutral lipids were present in all lipid extracts and represented 23–45 % of identified lipid classes (Fig. 3). The ratio of neutral lipids to phospholipids was similar to previous studies on pea and faba bean seeds, although this ratio has been found to vary considerably depending on seed cultivar (Coxon & Wright, 1985; Yoshida et al., 2007; Yoshida et al., 2009). Triacylglycerols (TAG) were the main neutral lipids present in the extracts, in addition to a small amount of free fatty acids (FFA). The FFA content was the highest in FPC_C2 and FPC_C3 at 6–8 % of identified lipid classes. These commercial concentrates were either stored for an extended period of time, FPC_C2, or heat-treated, FPC_C3. The sample storage of FPC_C3 could have induced FFA formation due to lipase activity previously described in faba bean (Yang et al., 2017). In addition, the heat treatment step of FPC_C2 could have caused lipid degradation if high enough temperatures were used (> 200 °C), also leading to FFA formation (Nawar, 1969). Diacylglycerols were found in trace amounts, which could be seen from complementary HPTLC analysis (Supplementary information 3 Fig. S3.1).

For all samples, part of the gravimetrically determined lipid content could not be found with HPLC analysis. This fraction generally represented around 15–25 % of the lipid extract, although it could reach up to 40 % in FPC_C2. The HPTLC analysis confirmed the presence of unidentified lipid compounds as well (Supplementary information 3). A part of these unknown lipids could correspond to hydrocarbons (Yoshida et al., 2008) and other unsaponifiable compounds, such as phytosterols and squalene (Miyazawa et al., 1974; Ryan et al., 2007; Singh et al., 2017). In addition, a small amount of glycolipids could be present, yet below the limit of detection of the HPLC-ELSD method (Haydar & Hadziyev, 1973; Keuleyan et al., 2023). In addition to lipid substances, a small amount of non-lipid components, for example oleosins, might have contaminated the lipid extract as well, leading to an overestimation of the lipid content (Li et al., 2025).

In addition to the lipid classes, the fatty acid (FA) composition was analysed in detail since the degree of FA unsaturation largely influences the lipids' sensitivity to oxidation (Gardner, 1989; Holman & Elmer, 1947). The main FAs were linoleic acid (C18:2 n-6), oleic acid (C18:1 n-9), palmitic acid (C16:0), stearic acid (C18:0), and α-linolenic acid (C18:3 n-3) (Fig. 4). These fatty acids are given in decreasing order, except that pea samples contained more C18:3 n-3 compared to C18:0 (Grela & Günter, 1995). These FA compositions are consistent with previous literature for faba bean (Yoshida et al., 2008) and pea seeds (Coxon & Wright, 1985; Miyazawa et al., 1975). Small differences in FA composition were found according to the seed cultivar, as previously reported by Yoshida et al. (2007). It is important to note that comparisons of FA composition results should only be made between studies using the same lipid extraction protocol, as the Soxhlet extraction has been found to yield an unrepresentable FA composition when applied to

<figure><img src="image_7.png" /><figcaption>**Fig. 3.** Lipid content and distribution in different lipid classes expressed as g per 100 g of dry material (DM). TAG = triacylglycerols; FFA = free fatty acids; PI = phosphatidylinositol; PG = phosphatidylglycerol; PC = phosphatidylcholine; PA = phosphatidic acid; PE = phosphatidylethanolamine. Values are displayed as the average of three independent lipid extractions ± standard deviation. FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Fatty acid composition expressed as percentage of total fatty acids. Values are displayed as the average of three independent lipid extractions. The coefficient of variation of each major class was maximum 3 %. Other includes C14:0, C15:0, C20:0, and C20:1 (n-9), each representing less than 2 % of the total fatty acids. FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>

legume seeds (Caprioli et al., 2016). In the present work, a lipid fractionation test of the commercial faba bean heat-treated concentrate, FPC_C3, and pilot-scale-produced isolate, FPI_P1, showed that neutral lipids contained a considerable fraction of C18:3 n-3 and C18:2 n-6 at 3–4 and 44–50 %, respectively. The phospholipid fraction contained only 0.9–1.1 % of C18:3 n-3 and 44–46 % of C18:2 n-6 (Supplementary information 4 Fig. S4.1). Such a difference in FA composition between lipid classes was also reported in pea seeds by Haydar and Hadziyev (1973).

Besides the major lipid compounds, minor lipophilic components can influence lipid oxidative stability as well (Shahidi & Zhong, 2010). Tocopherols are notably important in that respect, since these lipophilic antioxidants can protect lipids against oxidation by breaking radical chain reactions, although a prooxidant activity has also been reported in food systems depending on the tocopherol concentration (Barouh et al., 2022). The lipid extracts contained variable tocopherol contents, ranging from 800 to 2700 µg per g lipids (Fig. 5). These contents were remarkably high when compared to typical values in vegetable oils, found to range between 200 and 600 µg per g lipids (Elisia et al., 2013). The tocopherol content was the lowest for the commercial faba bean protein isolate, FPI_C1 at 238 ± 5 µg per g lipids, and the commercial pea protein concentrate from 2022, PPC_C2 at 411 ± 7 µg per g lipids. The major tocopherol isomer was γ-tocopherol, representing more than 85 % of total tocopherols, and α- and δ-tocopherol were found in much lower amounts. Tocopherols in similar samples have, to the best of our knowledge, only been analysed in Keuleyan et al. (2023). They also found high values for a commercial pea protein concentrate (502 µg per g lipids) and even higher for a commercial pea protein isolate (808 µg per g lipids). In the present study, FPC_P1 and FPI_P1 were prepared from the same seed cultivar and batch. From these samples, it can thus be concluded that both total pulse lipids and the associated tocopherols were more strongly accumulated throughout wet fractionation compared to dry fractionation. The commercial faba bean protein isolate, FPI_C1, had the lowest tocopherol content among all tested samples. This may be due to a low tocopherol content of the raw material, and/or to the applied wet fractionation process, although no information on this matter is available owing to the fact that this was a commercial sample.

To better compare these values to data available in literature, the tocopherol contents were converted into mg per kg of ingredient, which gave a range of 32–132 mg per kg for all ingredients except the commercial faba bean protein isolate and the commercial pea protein

<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Tocopherol content and distribution in different isoforms expressed in µg per g extracted lipids. Alpha-tocopherol (unless shown with the corresponding bars) and gamma-tocotrienol were present in trace amounts. Values are displayed as the average of three independent lipid extractions ± standard deviation. Columns indicated with a different letter have a significantly different total tocopherol content (P < 0.05). FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>

concentrate that was stored for an extended period of time. These two samples had lower contents of 16 and 22 mg per kg, respectively. These values could then be compared to those typically found in faba beans and pea seeds: 66–74 and 90–97 mg per kg seed, respectively (Grela & Günter, 1995; Yoshida et al., 2007; Yoshida et al., 2010). Some protein ingredients had a remarkably lower tocopherol content compared to those values found in their respective seeds. This might be due to tocopherol degradation during storage, as commercial ingredients produced in 2022, FPC_C2 and PPC_C2, were found to contain less tocopherols compared to 2023-produced ingredients from the same supplier, FPC_C1 and PPC_C1. However, the seed cultivar(s) used to obtain these commercial ingredients could have been different as well. The seed cultivar influenced the tocopherol content, as seen from the varying tocopherol content of pilot-scale-produced ingredients produced from 5 distinct faba bean cultivars. In addition to seed cultivar, seed growing conditions could be of importance as well, and were not necessarily aligned for the different seeds used in the present project, although all seeds used for pilot-scale-produced ingredients were grown in the same year. Pilot-scale-produced protein ingredients were produced from winter (FPC_P1, FPC_P2, and PPC_P1) or spring (FPC_P3, FPC_P4, and FPC_P5) varieties, a difference that could not correspondingly explain contrasting tocopherol contents. The growing location of the raw material also varied; FPC_P5 seeds were grown in Canada, whereas the other pilot-scale-produced ingredients were made from seeds grown in France. Previous research showed that tocopherol contents in plants are dependent on growing conditions, as the tocopherol content increases upon stressful conditions such as high light, water deficit, cold temperatures (Hasanuzzaman et al., 2014; Munné-Bosch & Alegre, 2002). However, contrasting results may be found in the literature: Yoshida et al. (2007) found no significant difference in tocopherol content across pea cultivars harvested from different locations, while Padhi et al. (2017) reported minor variation of 48–57 mg per kg seed among yellow pea seed varieties grown in Canada.

Similarly to tocopherols, carotenoids can act as primary antioxidants when present in lipid extracts, although high concentrations may lead to a prooxidant effect (Burton & Ingold, 1984; Elisia et al., 2013). One of the main carotenoids, β-carotene, has been found to synergistically increase the tocopherol antioxidant capacity (Palozza & Krinsky, 1992).
Carotenoids are lipophilic pigments and can therefore be analysed by measuring the absorbance of the lipid extract between 400 and 500 nm (Lichtenthaler & Buschmann, 2001). Their absorbance profile is characterised by three peaks (Fig. 6 and Supplementary information 5). The height of the absorbance peaks was correlated with the tocopherol content (R = 0.68, P < 0.001, Supplementary information 5 Fig. S5.4), i. e., samples that contained a high tocopherol content also contained more carotenoids. As the tocopherol content, the carotenoid content has previously been found to be influenced both by seed cultivar and growing conditions (Edelenbos et al., 2001).

## 3.2. Oxidative state of endogenous lipids

The oxidative state of endogenous lipids in pulse protein ingredients has, to the best of our knowledge, not been reported in literature. By deploying advanced microscopy methods, Brüls-Gill et al. (2024) observed oxidised lipids present within protein-lipid aggregates from several pulse protein isolates in the continuous phase of oil-in-water emulsions. These oxidised lipids were visualised through coupling with a fluorescent probe, but not quantified. They were present at the start of sample incubation, suggesting that the endogenous lipids in the protein ingredients had an initial degree of lipid oxidation. Consequently, the present work aimed to quantify both primary and secondary lipid oxidation markers in the obtained lipid extracts.

### 3.2.1. Primary lipid oxidation products

All lipid extracts contained elevated hydroperoxide (HPX) levels (Fig. 7). More specifically, all samples but the pilot-scale-produced pea protein concentrate, PPC_P1, had an HPX content that greatly exceeded the maximum peroxide value allowed for refined vegetable oils set at 5 µmol HPX per g lipids in the Codex Alimentarius. These results are therefore consistent with the observations of Brüls-Gill et al. (2024). It should, however, be considered that compared to vegetable oils that

contain mainly triacylglycerols, the protein ingredients had a lipid content of 4.4–7.5 g lipids per 100 g DM. Therefore, the HPX content was also expressed per g DM of the total ingredient. All samples then had an HPX content below 5 µmol HPX per g DM of powder (Fig. 7). In addition to the HPX content, primary lipid oxidation was also assessed by analysing conjugated dienes (Supplementary information 6 Fig. S6.1).

The samples had a highly variable HPX content, ranging from 2.7 to 57 µmol HPX per g lipids. The HPX content was negatively correlated with the tocopherol content (R = -0.78, P < 0.001, Supplementary information 6 Fig. S6.2), which could be indicative of the fact that tocopherols effectively displayed an antioxidant effect in these samples. As the tocopherol content may depend on the seed growing conditions, the HPX content differences could have similarly been affected. In addition, variation in enzymatic activity between samples could also be at play. Lipase activity could have caused the formation of FFA, known substrates for LOX. However, no correlation was found between the FFA and HPX contents. Ingredients from different seed cultivars were found to have different LOX band intensity in the SDS-PAGE analysis (Warsame et al., 2020). Nonetheless, the band intensity is correlated with LOX content, not directly with its activity. In addition, LOX activity would be important in case HPX were formed through enzymatic oxidation. The quantification of HPX content does not allow to determine whether HPX formation was primarily initiated through autooxidation or enzymatic oxidation.

Lipid fractionation allowed to look further into the distribution of HPX among the different lipid classes (Supplementary information 4 Fig. S4.2). For instance, the neutral lipids of the commercial faba bean heat-treated concentrate (FPC_C3) and pilot-scale-produced isolate (FPI_P1) were found to contain a 4- to 7-fold higher HPX content compared to their phospholipid fractions. The higher oxidative state of neutral lipids could be an indication that lipid oxidation occurred mostly enzymatically, as Haydar et al. (1975) found that neutral pea lipids were more susceptible to LOX oxidation than polar lipids, even in absence of lipase activity. In their study, LOX was not able to oxidise the pea phospholipids, except upon addition of phospholipase. Phospholipase was not unambiguously identified on the SDS-PAGE profiles, and further research is needed regarding its characterisation in pulses (Fig. 2). The higher HPX content of neutral lipids could also be explained by their higher degree of unsaturation (Haydar & Hadziyev, 1973), making them more susceptible to autooxidation as well.

<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Hydroperoxide content in µmol H₂O₂ equivalent per g extracted lipids (left axis) or per g of dry matter (DM) of powder ingredient (right axis). Values are displayed as the average of three independent lipid extractions ± standard deviation. Columns indicated with a different letter have a significantly different hydroperoxide content per g of extracted lipids (P < 0.05). FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>

<figure><img src="image_11.png" /><figcaption>**Fig. 6.** Absorbance spectra of lipid extracts from various protein ingredients at 5 mg lipids per mL in diethyl ether. One representative replicate per sample type is shown. The absorbance spectra of all of the other samples can be found in Supplementary information 5. FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>
### 3.2.2. Secondary lipid oxidation products

The HPX content of samples showed that lipid oxidation had occurred to a substantial extent in the protein ingredients. However, the content of primary lipid oxidation products is not directly linked to typical off-flavours associated with lipid oxidation. For off-flavours to occur, HPX need to decompose into secondary lipid oxidation products. The formation of the latter can be assessed using the reaction of carbonyl compounds with thiobarbituric acid (TBARS), which notably measures the formation of malondialdehyde (MDA). MDA is the main carbonyl compound formed through lipid oxidation, originating mostly from α-linolenic acid (C18:3 n-3) degradation (Schaich, 2005). TBARS ranged from 10 to 91 nmol per g lipids, with the highest amounts found in the commercial pea protein concentrates (Fig. 8). Complementary HPLC-DAD analysis quantified specifically MDA-TBA complexes, with values of 12–64 nmol per g lipids, which was slightly lower than the total TBARS concentrations for some samples (Fig. 8). From the lower MDA-TBA results, the TBARS content in the pea-based samples may have been overestimated due to species-specific interferents of the colorimetric assay. For the faba bean-derived samples, the TBARS and MDA-TBA quantification corresponded very well, confirming the reliability of the TBARS method to quantify secondary lipid oxidation products (Viau et al., 2016).

All samples had a relatively low MDA content around 12–64 nmol per g lipids when compared to the found HPX concentrations (Supplementary information 6 Fig. S6.3). This was surprising, as HPX decomposition could have taken place in the ingredient owing to the presence of pro-oxidant metals (section 3.1.2). Nevertheless, the TBARS/MDA quantification showed that HPX degradation was minimal. A possible explanation is that in the dry state of the powder ingredients, HPX do not come into contact with the metal ions (Karel, 1980). This would explain their relatively high stability. A potential consequence of these outcomes is the fact that, once the protein ingredient will be applied in a high-α<sub>W</sub> media, e.g., an aqueous suspension, the high initial HPX concentration may quickly lead to secondary oxidation product formation.

Previous research focussing on lipid oxidation in similar ingredients merely measured the headspace volatile profile (Liu et al., 2023; Murat et al., 2013; Sharan et al., 2022; Sumner et al., 1981). The volatile profile in the headspace of our ingredients was analysed, as a complementary analysis of secondary lipid oxidation products (Jacobsen, 1999; Nielsen et al., 2004; Villiere et al., 2007). In particular, the peak corresponding

to hexanal was scrutinized, as it is a typical marker of linoleic acid (C18:2 n-6) oxidation (Frankel, 2012; Genot et al., 2003; Schaich, 2005). When the adsorption of the volatile compounds on the SPME fibre was conducted at 35 °C, the volatilome did not show a typical lipid oxidation profile, with the hexanal peaks being only minor (Fig. 9, Supplementary information 7). It should be noted that, in this type of analysis, peak height is not directly correlated with the concentration of a compound in the sample, as it depends on the partitioning of the compound in question between sample-headspace-fibre, and on the possible competition for adsorption on the SPME fibre between multiple volatiles. Nonetheless, the absence of a typical lipid oxidation profile in the SPME analysis was in line with the low TBARS results and led to the conclusion that pulse protein powder ingredients do not show elevated levels of secondary lipid oxidation products.

To promote the partitioning of the volatile compounds potentially present in the powders towards the headspace, and thus the SPME fibre, the analysis was repeated with an exposure of the SPME fibre to the headspace of the sample at 50 °C, as this temperature was typically used in literature on similar ingredients (Akkad et al., 2019; Azarnia et al., 2011; Oomah et al., 2014; Sharan et al., 2021). For most samples, the SPME fibre exposure temperature did not change the allure of the chromatogram. Yet, the commercial faba bean protein isolate FPI_C1 sample did show a larger hexanal peak after SPME fibre exposure at 50 °C, as well as the appearance of several other lipid oxidation markers: 2-heptanone, 2,4-nonenal, 2-nonenal, and octadienone (Fig. 9). These differences seen at higher temperature could have two explanations. On the one hand, it is possible that the higher temperature allowed for a favoured partitioning of the volatile compounds towards the headspace (Drabińska & Jeleń, 2022). On the other hand, the FPI_C1 sample had the highest HPX content of all samples at 57 µmol per g lipids, and the exposure to this temperature could have led to a decomposition of HPX during the SPME fibre exposure time (Schaich, 2020). The TBARS content in this sample was relatively low, suggesting that the volatile compounds were preferentially formed from HPX decomposition during the SPME fibre exposure, rather than initially present in the sample. It is therefore recommended to carry out SPME analysis at moderate temperature, to avoid modifying the oxidative state of the sample.

## 4. Conclusion

A major outcome of this study was that faba bean and pea protein ingredients contained considerable amounts of endogenous lipids compared to the seeds they originate from. More than half of these lipids were phospholipids, which may largely influence the functionality of the ingredients, notably in stabilising interface-dominated systems such as foams and emulsions. The phospholipid profile was altered with wet fractionation, as phosphatidic acid was absent in protein isolates. In addition, the lipid fraction contained considerable amounts of tocopherols, which, as prominent antioxidants in the plant realm, could help stabilising these ingredients and food matrices prepared thereof against lipid oxidation. The ingredients' tocopherol content greatly depended both on the seed cultivar or growing conditions and the seed-to-ingredient processing method. Despite the generally high tocopherol content, the endogenous lipids present in the ingredient were readily prone to oxidation. Lipid oxidation led to a high and highly variable hydroperoxide (HPX) content in all samples, although the level of secondary lipid oxidation markers was fairly low. Consequently, off-flavour associated volatiles were not yet strongly present in the powdered protein ingredients. Nonetheless, considering the high initial HPX concentration of the endogenous lipids, it is expected that these off-flavours can quickly be formed upon HPX degradation in higher-α<sub>W</sub> media typically encountered in food matrices, such as plant-based dairy alternatives. In such conditions, HPX decomposition could be enhanced by the ingredients' content in pro-oxidant metals. These results thus shed light on the origin of lipid-derived off-flavours in pulse protein applications. To

<figure><img src="image_12.png" /><figcaption>**Fig. 8.** Malondialdehyde (MDA) equivalent concentrations as determined with a colorimetric analysis (thiobarbituric acid reactive substances, TBARS) and through HPLC separation and diode array detection (MDA). Values are displayed as the average of three independent lipid extractions ± standard deviation. Bars indicated with a different letter have a significantly different malondialdehyde equivalent concentration per g of extracted lipids (*P* < 0.05). FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>
<figure><img src="image_13.png" /><figcaption>**Fig. 9.** Flame ionisation detector (FID) response after solid-phase microextraction (SPME) (SPME fibre exposure conducted at 35 or 50 °C) and GC separation of volatile compounds present in the headspace of the protein ingredients FPI C1 (A) and PPC P1 (B). Peaks were annotated based on the mass spectrometry response by comparing fragmentation profiles with the NIST library. The chromatogram of remaining samples can be found in Supplementary information 7. FPC = faba bean protein concentrate; FPI = faba bean protein isolate; PPC = pea protein concentrate. The final letter in the sample code stands for the origin of the sample (C = commercial; P = produced at pilot-scale). For a full description of the different samples, see section 2.2 and Table 1.</figcaption></figure>

improve the flavour of pulse protein applications, it is key to better understand and hereafter control lipid oxidation reactions. In addition, lipid accumulation during protein fractionation is an integral part of pulse protein ingredient off-flavour, as the endogenous lipid content in protein ingredients is non-negligible. Analysing the composition of these endogenous lipids in detail should be an essential part of research assessing off-flavour formation through lipid oxidation.

## CRediT authorship contribution statement

**Jolijn S.A. Koomen:** Writing – review & editing, Writing – original draft, Visualization, Validation, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Lucie Ribourg-Birault:** Writing – review & editing, Validation, Methodology, Investigation, Data curation. **Alice Kermarrec:** Writing – review & editing, Validation, Methodology, Investigation, Data curation. **Bérengère Marais:** Validation, Investigation, Data curation. **Sophie Le Gall:** Writing – review & editing, Validation, Methodology, Investigation. **Adeline Boire:** Writing – review & editing, Visualization, Supervision, Project administration, Methodology, Funding acquisition, Conceptualization. **Anne Meynier:** Writing – review & editing, Visualization, Supervision, Project administration, Methodology, Funding acquisition, Conceptualization. **Claire Berton-Carabin:** Writing – review & editing, Visualization, Supervision, Project administration, Methodology, Funding acquisition, Conceptualization.

## Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

This work benefitted of the financial support of the French government through the National Research Agency (ANR) as part of France 2030 in the framework of LETSPROSEED ANR-22-PELG-002, and the financial support of INRAE for the Ph.D. thesis of JK. The authors would like to thank Joëlle Davy and Sophie Laurent for their technical and analytical support. Carbohydrate analyses were performed on the BIBS instrumental platform (UR1268 BIA, IBiSA, Phenome-Emphasis-FR ANR-11-INBS-0012, PROBE, CALIS infrastructures, Biogenouest, DOI:

10.15454/1.5572358121569739E12; http://www.bibs.inrae.fr/bibs_eng/).

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.foodres.2025.116884.

## Data availability

Data will be made available on request.

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