Natural Oleosomes from Nuts and Seeds: Structural Function and Potential for Pharmaceutical Applications
1Faculty of Pharmacy and Pharmaceutical Sciences, University of Alberta, Edmonton, AB T6G 2E1, Canada; mallilli@ualberta.ca (M.C.M.III); msalami2@ualberta.ca (M.S.); shengna2@ualberta.ca (S.Z.); elmahrou@ualberta.ca (S.R.E.-M.); kirtypal@ualberta.ca (K.S.); siraki@ualberta.ca (A.G.S.); aelkadi@ualberta.ca (A.O.S.E.-K.)
2Department of Pharmacy, Faculty of Pharmacy, University of Santo Tomas, Manila 1015, Philippines; oavillalobos@ust.edu.ph
3Department of Food Science, Engineering and Technology, College of Agriculture & Natural Resources, Karaj Campus, University of Tehran, Karaj 31587-77871, Iran
4China Z. Pharmaceutical Productivity Centre, Beijing 101111, China
5Faculty of Science, Department of Chemistry, University of Alberta, Edmonton, AB T6G 2G2, Canada; serpe@ualberta.ca
6Faculdade de Ciências Farmacêuticas, Universidade de São Paulo, São Paulo 05508-000, Brazil; chacra@usp.br
Abstract
Background/Objectives: Oleosomes, plant-derived lipid nanostructures comprising a triacylglycerol core surrounded by a phospholipid monolayer and interfacial proteins, provide sustainable alternatives to synthetic lipid vesicles. This study compares solvent-free aqueous extractions of oleosomes from five nuts (almond, macadamia, walnut, hazelnut, pine) and five seeds (flaxseed, sunflower, hemp, sesame, canola/rapeseed) to understand how botanical origin influences composition and physicochemical behavior. Methods: Oleosomes were isolated using solvent-free aqueous extraction. Extraction yield, lipid content, protein content, particle size, polydispersity, and zeta potential were determined using standard analytical assays and dynamic light scattering techniques. SDS–PAGE was performed to evaluate interfacial protein profiles and oleosin abundance. Results: Extraction yields ranged from 8.4% (flaxseed) to 59.5% (walnut). Oleosome diameters spanned 424 nm to 3.9 µm, and all oleosome dispersions exhibited negative zeta potentials (–26 to –57 mV). SDS–PAGE revealed abundant 15–25 kDa oleosins in seed oleosomes but relatively sparse proteins in nut oleosomes. Seed oleosomes were smaller and exhibited stronger electrostatic stabilization, while nut oleosomes formed larger droplets stabilized primarily through steric interactions due to lower oleosin content. Conclusions: Variation in oleosin abundance and interfacial composition leads to distinct stabilization mechanisms in nut and seed oleosomes. These findings establish a predictive basis for tailoring oleosome size, stability, and functionality, and highlight their potential as natural nanocarriers for food, cosmetic, and pharmaceutical formulations.
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Keywords: oleosomes, sustainable nanocarriers, food technology, cosmetics, green nanotechnology, natural materials
Article notes
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Received 2025 Nov 25; Revised 2026 Jan 13; Accepted 2026 Jan 20; Collection date 2026 Feb.
1. Introduction
Oleosomes, also called plant oil bodies, are intracellular lipid storage organelles composed of a hydrophobic triacylglycerol (TAG) core surrounded by a phospholipid monolayer and structural proteins such as oleosin, caleosin, and steroleosin [1]. These proteins impart remarkable stability through steric and electrostatic mechanisms that prevent coalescence and oxidation.
Due to their amphiphilic nature, oleosomes self-stabilize without the need for synthetic surfactants, aligning with sustainable “green” formulation strategies. Compared to conventional lipid nanoparticles such as solid lipid nanoparticles (SLNs), nanostructured lipid carriers (NLCs), or emulsions that rely on chemical surfactants, oleosomes offer inherent biocompatibility and solvent-free processing, which is attractive for food, cosmetic, and pharmaceutical applications [1,2,3].
Vegetable oils are long recognized as safe excipients. Sesame, castor, peanut, sunflower, and olive oils are routinely used as vehicles for parenteral, topical, and otic dosage forms [3,4]. Oleosomes encapsulate these same triglycerides within nanostructured droplets stabilized by natural proteins, integrating the safety of edible oils with the structural sophistication of colloidal nanocarriers.
Although oleosomes have been examined primarily in individual seeds, such as rapeseed or soybean [5,6], a systematic comparison of nut- and seed-derived oleosomes under identical aqueous extraction conditions remains limited, particularly regarding their potential use in food emulsions and nutraceutical delivery systems.
Recent advances have expanded oleosome research beyond food science into diverse industrial sectors, including paints, coatings, lubricants, films, gels, inks, waxes, and even road construction materials, where their natural interfacial films support the “green” replacement of synthetic surfactants [2,7].
Multiple studies have demonstrated that oleosomes can encapsulate and stabilize a range of lipophilic nutraceuticals and pharmaceutical bioactives, including cannabidiol, curcumin, and β-carotene, resulting in improved protection, colloidal stability, and delivery compared to bulk oils or simple emulsions [8,9]. In pharmaceutical contexts, oleosomes serve as biocompatible, excipient- and surfactant-free lipid carriers, offering a natural alternative to synthetic lipid nanoparticle systems, such as solid lipid nanoparticles and nanostructured lipid carriers [2,7]. Significantly, interfacial characteristics that govern pharmaceutical performance, including droplet size, surface charge, and interfacial protein composition, vary substantially across botanical sources, particularly between nuts and seeds [10]. Despite this recognized variability, systematic cross-source comparisons of oleosomes extracted under identical, solvent-free conditions remain limited.
This study tests the hypothesis that nut-derived oleosomes, which are richer in lipid and lower in protein, yield higher extraction efficiency and predominantly steric stabilization. In contrast, seed-derived oleosomes, which contain higher protein levels, produce smaller, electrostatically stabilized droplets [1,5,10,11,12]. By analyzing the physicochemical characteristics and protein banding profiles, this work provides an integrated framework for understanding how the botanical origin influences the structure, stability, and functional performance of oleosomes.
2. Materials and Methods
2.1. Materials
2.1.1. Seeds and Nuts
Five nuts (almond, macadamia, walnut, hazelnut, and pine) and five seeds (flaxseed, sunflower, hemp, sesame, and canola/rapeseed) were sourced from certified commercial suppliers that provide food-grade, lot-traceable products. Unless otherwise specified, all materials were stored in resealable zipper bags at 25 °C until use.
Sunflower seeds (Elan, Tootsi Impex Inc., Lot 25 216) were purchased from a supplier located in Montreal, QC, Canada (H4S 1P4). Hemp seeds (Hemp Hearts, Fresh Hemp Foods Ltd., Lot 250206) were obtained from Winnipeg, MB, Canada (R2R 1V4). Flaxseed (Bob’s Red Mill, Bob’s Red Mill Natural Foods Inc., Lot 24-120-WHQ-0-16) was procured from Milwaukee, OR, USA (97222). Sesame seeds (Kevala, Kevala International LLC, Lot KEX-01205) were sourced from Dallas, TX, USA (75367). Canola/rapeseed (Apache Seeds, Canterra Seeds, Lot CS3100tf) was obtained through UFA, Lethbridge, AB, Canada (T1H 6M1).
The nut samples included macadamia nuts (Going Nuts, Lot L2523/25), pine nuts (Going Nuts, Lot L2523/27), and hazelnuts (Going Nuts, Lot L2223/86), all sourced from Going Nuts, Calgary, AB, Canada (T2G 1Y6). Almonds (Kirkland, Costco Wholesale Canada Ltd., Lot 284601) were purchased from Ottawa, ON, Canada (K2E 1C5). Walnuts (Co-op Gold, Federated Co-operatives Limited, Lot 135844) were obtained from Saskatoon, SK, Canada (S7K 3M9).
2.1.2. Reagents and Consumables
Sodium bicarbonate (NaHCO3, ≥99%, Caledon Laboratories Ltd., Georgetown, ON, Canada), sodium hydroxide (NaOH, ACS grade, Caledon Laboratories Ltd., Georgetown, ON, Canada), Tris–HCl buffer (Bio-Rad Laboratories, Hercules, CA, USA), hexane (≥95%, Sigma-Aldrich, St. Louis, MO, USA), Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA), Coomassie Brilliant Blue R-250 (Bio-Rad Laboratories, Hercules, CA, USA), methanol (≥99.8%, Fisher Scientific, Fair Lawn, NJ, USA), and glacial acetic acid (≥99.7%, Fisher Scientific, Fair Lawn, NJ, USA) were used as received. Ultrapure water (18.2 MΩ·cm) was obtained from a Milli-Q purification system (MilliporeSigma, Burlington, MA, USA). Two-ply cotton cheesecloth (nominal pore size ≈ 150 µm) was used for coarse filtration.
2.1.3. Instruments
Homogenization was performed using a Thermomix blender (Vorwerk, Germany). Centrifugation was performed using a Beckman Coulter Avanti J-20 XP with a JLA-16.250 rotor and a Thermo Scientific Sorvall ST Plus centrifuge with an F15-6×100y rotor and 50 mL conical tube adapters. pH was monitored using a Thermo Scientific Accumet AB250 pH/conductivity meter, which was calibrated daily with pH 7.00 and 10.00 buffers. Particle size, polydispersity index, and zeta potential were measured using a Malvern Zetasizer Ultra Red (Malvern Panalytical, Worcestershire, UK) operating under dynamic light scattering (DLS) and electrophoretic light scattering (ELS) modes. Microscopy was conducted using an ECHO Revolve revolve2-k2-1601 microscope (Discover Echo, San Diego, CA, USA). Lyophilization was carried out using a Labconco FreeZone freeze dryer.
2.2. Aqueous Extraction of Oleosomes
2.2.1. Seed Soaking
Batches of dry seeds were soaked for 24 h at 4 °C at a seed-to-liquid ratio of 1:7 w/w using one of the following media: (i) 0.1 mol/L NaHCO3 adjusted to pH 9.5 with 1.0 mol/L NaOH, or (ii) Milli-Q water adjusted to pH 7.0 or 8.0 with 0.1 mol/L NaOH. All soaking was performed in sealed containers at 4 °C, and pH was verified at both the start and end of the soaking process, consistent with previous reports that mildly alkaline conditions promote oleosome release while preserving interfacial integrity [1,13,14].
The soaking conditions (24 h at 4 °C, 1:7 w/w solid-to-liquid ratio) were selected based on commonly used oleosome extraction protocols, which report that extended cold hydration maximizes oleosome release while preserving interfacial protein integrity and minimizing enzymatic degradation [1,13,14].
Sodium bicarbonate buffer (pH 9.5) was selected as the standard extraction medium, as mildly alkaline conditions (pH 8–9.5) have been shown to promote efficient oleosome release while preserving interfacial protein integrity across a wide range of oilseeds [1,13,14]. Flaxseed was treated as a specific exception because its high mucilage content resulted in reduced dispersion quality under pH 9.5 conditions. For this source, pH 8 water produced more stable and reproducible oleosome preparations, consistent with prior reports on flaxseed oleosome extraction [6,15].
2.2.2. Homogenization and Coarse Filtration
Pre-soaked seeds and fresh extraction medium were combined to maintain a 1:7 w/w ratio at 4 °C. The slurries were then blended at approximately 7500 rpm for 60 s. Homogenates were filtered through two layers of cheesecloth to remove insoluble solids. Filtrates were kept at 4 °C and processed on the same day.
2.3. Isolation and Washing of Oleosomes
Crude filtrates were centrifuged at 10,000× g for 30 min at 4 °C in polypropylene copolymer (PPCO) bottles, which were filled to approximately 80% of the nominal volume and balanced within ±0.1 g. The upper cream layer was collected. The cream was washed by dispersing it in fresh medium matching the soak chemistry, followed by centrifugation at 8000× g for 30 min at 4 °C in 50 mL conical polypropylene tubes, ensuring a balance of 0.1 ± 0.05 g. Brake and acceleration were set to medium. The final washed cream was either stored at 4 °C for short-term analysis or frozen for lyophilization.
2.4. Lyophilization
Cream aliquots were frozen at −80 °C and then lyophilized at approximately one mbar until a constant mass was achieved, typically over 24 to 36 h. Constant mass was defined as less than 1 mg change over a 2 h interval at the end of the run. Dry powders were stored sealed at −20 °C with desiccant.
Lyophilization was performed solely as an analytical preparation step to enable accurate normalization of lipid and protein content on a dry-mass basis and to ensure reproducible compositional comparisons across botanical sources.
2.5. Lipid Content Determination
Approximately 100–200 mg of lyophilized oleosome cream was extracted three times with hexane (20 mL solvent per g of dry sample, i.e., 2.0–4.0 mL solvent for 100–200 mg) under ambient temperature. For each extraction cycle, the sample was vortexed for 1 min to resuspend the matrix, followed by centrifugation at a minimum of 3000× g for 10 min at ~25 °C to separate the solvent phase from solids. The supernatants from all three cycles were pooled, and the solvent was evaporated under a gentle nitrogen stream at a temperature of ≤40 °C until dryness. The residue was further dried in a desiccator to constant weight and then weighed.
2.6. Protein Quantification by Bicinchoninic Acid (BCA)
Protein concentrations were measured using the Pierce BCA Protein Assay Microplate Protocol (Thermo Fisher Scientific, Product #23252). BSA standards ranging from 25 to 1000 µg/mL were prepared in the same matrix as the samples. Samples were lightly clarified by brief centrifugation to minimize lipid carryover. Absorbance at 562 nm was measured after incubation according to the specified times outlined in the kit. Concentrations were calculated using four-parameter logistic fits with a target R2 ≥ 0.99 and corrected for sample dilution. Spike recovery acceptance criteria were set at 80–120%.
Protein concentration was reported as µg/mL of oleosome dispersion to provide a comparative measure of surface-associated protein abundance under identical dilution conditions used for particle size and zeta potential measurements. This approach enables direct correlation between protein content and colloidal behavior, which is central to evaluating stabilization mechanisms in oleosome systems [1,2].
2.7. SDS–PAGE Protein Profiling
SDS–PAGE (sodium dodecyl sulfate polyacrylamide gel electrophoresis) was used to profile oleosome-associated proteins. Samples were mixed 1:1 with Laemmli sample buffer, and 5 to 20 µg of protein was loaded per lane on 15% gels. Gels were stained with Coomassie Brilliant Blue R-250 for 30 to 60 h and then destained in a methanol–acetic acid–water solution until the background cleared. The ladder spanned 10 to 250 kDa.
2.8. Particle Size and Polydispersity Index (PDI) by DLS
Oleosome dispersions were prepared at 1:10 w/v in Milli-Q water immediately before analysis to provide a standardized, low-ionic-strength medium. This approach minimizes ionic screening effects, enabling the direct comparability of scattering behavior across botanical sources, regardless of the extraction buffer composition. DLS was performed at 25 °C using a Zetasizer Ultra in backscatter mode at 173° with auto-attenuation. Dispersant parameters were RI 1.333 and viscosity 0.8872 mPa·s. Each replicate was measured in technical triplicate with 10 to 100 sub-runs per measurement as selected by the instrument.
2.9. Zeta Potential by Electrophoretic Light Scattering
Electrophoretic mobility was measured at 25 °C using the Zetasizer Ultra (Malvern Panalytical, Worcestershire, UK) with DTS1070 folded capillary cells. Dispersions were prepared at a 1:10 w/v ratio in Milli-Q water to ensure a common, low-ionic-strength medium that minimizes ionic screening and allows direct comparison of electrophoretic mobility across oleosome sources. Samples were equilibrated for 2–3 min before measurement. The Smoluchowski model was applied. Each extraction was measured in technical triplicate.
2.10. Qualitative Loading of Model Compounds into Oleosomes
Oleosomes were qualitatively evaluated for their ability to associate with model compounds of varying polarity using visual loading experiments. Selected compounds were added to freshly prepared oleosome dispersions at a 1:10 (w/w) drug-to-oleosome ratio. The mixtures were incubated for 3 h at room temperature under constant stirring to allow interaction between the compounds and the oleosome phase.
Following incubation, samples were centrifuged at 8000× g for 30 min at 4 °C to concentrate the oleosome-rich cream layer. The recovered cream was then resuspended in Milli-Q water at a 1:1 (w/w) ratio and gently mixed to remove loosely associated compounds. The dispersion was centrifuged again under the same conditions to re-collect the oleosome fraction. Only a single washing step was performed.
2.11. Microscopy
Dispersions at 1:10 w/v were imaged in brightfield using the ECHO Revolve with a 0.30 NA objective. A 10 µL drop was placed on a glass slide, covered with a No. 1.5 coverslip, and allowed to settle for approximately one minute. Then, it was imaged using a fixed camera exposure and gain. Only linear global brightness and contrast adjustments were applied.
2.12. Statistical Analysis
Quantitative data are presented as mean ± standard deviation (SD), with the number of replicates indicated in each figure legend. Extraction yield, lipid content, and protein content were measured in triplicate (n = 3), whereas particle size, polydispersity index, and zeta potential were determined from five independent extractions (n = 5). Differences among oleosomes from different botanical sources were evaluated using the Kruskal–Wallis nonparametric test, with a significance level of p < 0.05 (α = 0.05). Graphs were generated using BioRender Graph, and schematic illustrations were created with BioRender (https://www.biorender.com, accessed on 3 November 2025).
3. Results
3.1. Extraction Yield and Lipid Composition
Extraction efficiency differed significantly among botanical sources (Kruskal–Wallis H = 18.857, p = 0.026), ranging from 8.4% to 59.5% (Figure 1). The highest cream yields were obtained from walnut (59.5%), macadamia (58.8%), and pine (36.4%), consistent with their lipid-dense, readily disrupted tissues. The superior performance of nut-derived materials likely reflects a looser parenchymal architecture and softer lipid bodies, which facilitate oleosome release during homogenization, in agreement with structural observations in almonds and hazelnuts [10,12]. Similar trends have been observed in tree nut oleosomes, which often exhibit larger oil bodies and more fragile cell matrices that are more conducive to release [2,11]. In contrast, flaxseed (8.4%) and hemp (25.6%) exhibited lower extraction efficiencies, which can be attributed to the presence of surface mucilage and more pronounced lignocellulosic barriers that impede oleosome release [6,15].
The oil content differed significantly among sources (Kruskal–Wallis H = 28.925, p < 0.001), ranging from 21% to 85% (Figure 2). Sesame, hazelnut, and almond oleosomes exhibited more than 80% lipid, whereas flaxseed and hemp were more protein-rich (21–58%). High lipid-to-protein ratios (L/P > 40) characterize sterically stabilized nut oleosomes, while lower ratios (L/P approximately 15–25) indicate electrostatically stabilized seed oleosomes, consistent with earlier reports on plant oil bodies [1,2,7].
To further rationalize these trends, the influence of the hydration medium was examined by comparing extractions performed in ultrapure water (pH 7–8) and sodium bicarbonate buffer (pH 9.5). Extractions conducted in ultrapure water generally yielded lower cream yields and visibly fewer stable dispersions, with partial oil leakage from the oleosomes, indicating disruption of the native interfacial structure. In contrast, extractions using the sodium bicarbonate buffer produced higher and more reproducible yields with intact, well-dispersed oleosome creams for all nuts and seeds. This conclusion was supported by multiple lines of evidence, including a thicker cream layer and a reduced free-oil phase, stable dispersions over 24–48 h at room temperature, uniform droplet morphology observed via light microscopy, and narrower particle-size distributions with more negative zeta potential values, consistent with enhanced colloidal stability. Therefore, all subsequent experiments and discussions are based on buffer-extracted samples, except for flaxseed, where pH 8 water extraction produced the most stable dispersion and was retained as the optimized condition. The choice of a mildly alkaline sodium bicarbonate buffer (pH 9.5) is consistent with previous reports, which show that pH 8–9.5 conditions promote the solubilization of interfacial and storage proteins, facilitate oleosome release, and maintain membrane integrity without excessive denaturation [1,13,14].
The high cream yields observed for walnut (59.5%) and macadamia (58.8%) are consistent with their lipid-dense tissues and favorable cellular architecture, which facilitate oleosome release and cream phase separation during aqueous homogenization and centrifugation. Tree nut oleosomes are known to exhibit larger droplet sizes and lower interfacial protein density which promotes efficient creaming behavior [1,10,12].
Differences in lipid and protein composition among oleosome extracts reflect botanical variations in lipid-to-protein ratio and co-extracted matrix components. Oleosomes from sesame, hazelnut, canola, and almond exhibited lipid contents exceeding 80%, consistent with lipid-rich oleosomes containing relatively lower proportions of surface-associated proteins [2]. In contrast, flaxseed and hemp oleosomes were comparatively protein-rich, which has been attributed to their mucilage- and protein-rich seed matrices, which promote increased protein association and non-lipid carryover during aqueous extraction [6,13,16].
3.2. Protein Composition and SDS–PAGE Profiles
3.2.1. Band Identification
SDS–PAGE profiles displayed characteristic protein bands across all samples. Prominent bands between 15–25 kDa correspond to oleosins, major structural proteins that stabilize the TAG-phospholipid interface. Intermediate bands at 30–35 kDa represent caleosins, which contribute to oxidative and mechanical resilience. Fainter high-mass bands (40–60 kDa) correspond to steroleosins or associated enzymes (Figure 3). Protein bands were assigned to oleosins, caleosins, and steroleosins based on their well-established molecular weight ranges reported for plant oleosomes.
In addition to the characteristic oleosin bands (15–25 kDa), higher-molecular-weight bands in the ~70–250 kDa range were observed. These bands are attributed to co-extracted storage proteins, oleosome-associated enzymes, and protein complexes or aggregates that remain associated with the oleosome interface following aqueous extraction. Previous studies have shown that such high-molecular-weight bands can arise from oligomeric or disulfide-linked protein assemblies, as well as partial association of non-oleosin proteins with the phospholipid monolayer of plant oil bodies [1,11,13].
While BCA quantifies total protein content and particle size and zeta potential describe bulk colloidal behavior, SDS–PAGE provides molecular-level resolution of oleosome-associated proteins, particularly the relative abundance of low-molecular-weight oleosins. Oleosomes enriched in oleosins are expected to exhibit smaller droplet sizes and stronger electrostatic stabilization, whereas oleosome systems with reduced oleosin abundance are expected to form larger, sterically stabilized droplets. This interpretation would be challenged if oleosin-rich samples exhibited large droplet sizes and weak surface charge, or if oleosin-poor samples formed small, highly charged droplets.
3.2.2. Comparative Patterns
Seed oleosomes (sunflower, sesame, canola/rapeseed, hemp, flaxseed) showed strong, well-defined 15–25 kDa bands, confirming high oleosin abundance. In contrast, nut oleosomes (almond, hazelnut, walnut, pine, macadamia) exhibited weaker low-mass bands and more prominent medium-to-high-mass bands, indicating reduced oleosin density and thicker lipid layers.
These profiles correlate with physicochemical measurements: seed oleosomes, rich in oleosin, produced smaller, more stable droplets; nut oleosomes, lower in oleosin, exhibited larger droplets stabilized sterically. Such patterns mirror earlier findings in sunflower and rapeseed oleosomes [5,13].
Comparable oleosin distributions (15–25 kDa) have also been described in soy, walnut, and jujube oleosomes, where the protein-to-lipid ratio directly influenced droplet stability [10,17]. The strong preservation of these structural proteins supports the universal interfacial role of oleosins, as reported by Nikiforidis and [1] and Fu et al. [6].
3.2.3. SDS–PAGE as Corroborative Structural Evidence
Protein content differed significantly among oleosome sources (Kruskal–Wallis H = 28.277, p < 0.001) (Figure 4). SDS–PAGE profiles aligned closely with quantitative BCA and size measurements. Samples with high protein content (for example, sunflower, 1910 µg/mL, and canola or rapeseed, 1715 µg/mL) exhibited prominent low-molecular-mass oleosin bands, which correlated with smaller droplet sizes and higher zeta potentials. In contrast, hazelnut (468 µg/mL) and walnut (440 µg/mL), which contained less interfacial protein, formed larger, more weakly charged droplets. These trends align with previous work on tree nuts, hemp, and jujube oleosomes, where the protein-to-lipid ratio has been shown to influence interfacial coverage and colloidal stability directly [10,11,16,17]. The co-presence of oleosin, caleosin, and steroleosin bands confirms preservation of the native interfacial protein ensemble. Collectively, these data indicate that the aqueous extraction process maintains protein integrity, reinforcing its solvent-free advantage for various industrial applications.
3.3. Particle Size and PDI
Hydrodynamic diameters differed significantly among oleosome sources (Kruskal–Wallis H = 47.562, p < 0.001; n = 5) (Figure 5), ranging from a submicron mean diameter of ~424 nm in sunflower to ~3.9 µm in hazelnut (Figure 6a,b). Polydispersity indices spanned 0.16–0.54, with relatively narrow size distributions for walnut, pine and macadamia (PDI ≈ 0.16–0.18) despite their larger droplet sizes (1.9–2.5 µm), and broader distributions for sunflower and flaxseed (PDI ≈ 0.40–0.54). Sunflower was the only system that formed predominantly submicron droplets (<500 nm). In contrast, all other seed and nut dispersions exhibited volume-weighted mean diameters in the low micrometer range (~1.1–3.0 µm). All extracts carried moderately high negative zeta potentials (about −26 to −57 mV), consistent with electrostatically stabilized oil-in-water dispersions. Systems combining a smaller droplet size with higher interfacial protein content, such as sunflower and canola, also showed higher BCA-derived protein levels (approximately 1.7–1.9 mg/mL). In contrast, larger droplets in hazelnut, walnut and hemp coincided with lower or intermediate protein contents. Functionally, the sub-500 nm sunflower oleosomes are well suited for stable nanoemulsions and uniform dispersions used in cosmetic, pharmaceutical coating, or food formulations, where smaller droplets and narrow size distributions support improved colloidal stability and more homogeneous delivery of bioactives [2,5,8,9]. In contrast, the larger droplets typical of nut-derived systems, such as hazelnut and walnut, may be advantageous for topical or depot applications, where slower diffusion and more occlusive behavior support sustained release [10,11,18].
3.4. Zeta Potential and Colloidal Stability
All samples showed negative zeta values (−26.5 to −57.4 mV) (Figure 7). This observation aligns with previous reports showing that oleosomes’ negatively charged phospholipid interfaces contribute to long-term dispersion stability [11,16]. Similar electrostatic stabilization has been exploited in functional emulsion systems and cosmetic formulations [2]. Hemp (−57 mV) and sunflower (−52 mV) exhibited the highest magnitudes due to surface phosphate and carboxylate groups [16]. Nut oleosomes, with lower zeta magnitudes, were stabilized by steric hindrance.
Compared with synthetic SLNs stabilized by non-ionic surfactants (−25 to −45 mV), oleosomes achieve similar electrostatic stability without the use of surfactants [3].
Combining zeta and size data highlights a clear inverse correlation: higher magnitude negative zeta values correspond to smaller, more stable droplets, in line with earlier reports on hemp and nut oil bodies [10,11,16].
3.5. Qualitative Visual Association of Model Compounds with Oleosomes
Oleosomes exposed to model compounds spanning a wide polarity range exhibited visible coloration following incubation and a single aqueous washing step. Hydrophobic compounds produced intense coloration of the oleosome-rich cream layer, whereas hydrophilic compounds resulted in more uniform coloration throughout the dispersion. (Table 1) Amphiphilic compounds showed intermediate behavior, consistent with partial partitioning between lipid and aqueous domains. These observations provide qualitative evidence that natural oleosomes can visually associate with compounds across a broad range of polarities. (Figure 8) Because the assessment was based solely on visual inspection, the results are interpreted as a qualitative visual association rather than a quantitative measure of drug loading.
| Model Compound | Polarity Classification | Visual Property |
|---|---|---|
| Mitoxantrone | Hydrophilic | Blue |
| Mitomycin C | Hydrophilic | Purple |
| Vitamin B12 | Hydrophilic | Red |
| Betanin | Hydrophilic | Red-violet |
| Rifampicin | Amphiphilic | Orange |
| Doxorubicin | Amphiphilic | Red |
| Curcumin | Hydrophobic | Yellow |
4. Discussion
This study presents a systematic, solvent-free comparison of oleosomes extracted from nuts and seeds under standardized aqueous conditions, establishing clear structure–function relationships that link botanical origin to extraction yield, lipid and protein composition, particle size, surface charge, and interfacial protein profiles. Two dominant stabilization regimes were identified: protein-rich, electrostatically stabilized seed oleosomes and lipid-rich, sterically stabilized nut oleosomes.
Seed-derived oleosomes (flaxseed, sunflower, hemp, sesame, and canola/rapeseed) consistently formed smaller, highly charged, protein-dense droplets governed primarily by electrostatic stabilization. In contrast, nut-derived oleosomes (from almond, macadamia, walnut, hazelnut, and pine) yielded larger, lipid-rich droplets that are stabilized predominantly through steric mechanisms. (Table 2) These distinctions reflect inherent botanical differences in cellular architecture and the composition of storage tissues. Lipid-rich nut matrices favor the formation of larger oleosomes that readily separate into the cream phase, resulting in higher apparent yields and lipid contents. In contrast, flaxseed and hemp contain mucilage- and protein-rich matrices that promote increased interfacial protein association and electrostatic stabilization [1,13,16].
| Parameter | Seeds | Nuts |
|---|---|---|
| Yield (%) | 8–54 | 30–60 |
| Lipid content (%) | 21–85 | 63–82 |
| Protein (µg/mL) | 325–1910 | 440–1500 |
| Size (nm) | 424–3036 | 1265–3907 |
| PDI | 0.28–0.54 | 0.16–0.36 |
| Zeta Potential (mV) | −33 to −57 | −29 to −47 |
Protein concentration determined by BCA analysis served as a mechanistic indicator of oleosome interfacial composition rather than an absolute compositional metric. Higher protein concentrations reflected an increase in surface-associated proteins that contribute charged functional groups, thereby enhancing electrostatic stabilization. In contrast, lower protein concentrations were consistent with lipid-rich oleosomes stabilized predominantly by steric effects. Reporting protein content per dispersion volume enabled direct comparison with particle size and zeta potential data obtained under identical preparation conditions [1,13,16].
The concordance between SDS–PAGE profiles, BCA-derived protein concentrations, and particle size and zeta potential measurements further supports the proposed stabilization mechanisms. SDS–PAGE provided molecular-level resolution of interfacial protein composition, particularly in terms of oleosin abundance, which could not be inferred from bulk protein quantification or colloidal measurements alone. Together, these complementary analyses confirm that stabilization behavior arises from genuine interfacial compositional differences rather than analytical artifacts.
Variations in soaking time or solid-to-liquid ratio have been reported to primarily influence absolute extraction yield rather than the relative physicochemical characteristics of oleosomes. Accordingly, while changes in hydration conditions may affect recovery efficiency, the comparative structure–function trends observed across botanical sources are expected to remain unchanged under the standardized extraction conditions employed in this study [2,5].
The physicochemical attributes quantified here are directly relevant to oleosome performance in pharmaceutical systems. Parameters such as droplet size, lipid-to-protein ratio, interfacial protein integrity, and colloidal stability play crucial roles in formulation behavior and bioactive performance [9]. The present findings, therefore, provide a pre-formulation framework for selecting oleosome sources with properties that align with pharmaceutical requirements.
From a formulation perspective, oleosome suitability is inherently application dependent. Sunflower oleosomes appear particularly well suited for dispersed pharmaceutical systems due to their smaller droplet size, relatively narrow size distribution, high interfacial protein content, and strong negative zeta potential. In contrast, lipid-rich nut oleosomes such as hazelnut, almond, and walnut may be better suited for applications requiring higher lipid loading or depot-type behavior [1,8,9,10,16].
The qualitative loading experiments further support the functional versatility of oleosomes as natural carrier systems. The observed visual association of both hydrophilic and hydrophobic model compounds highlights the multifunctional nature of the oleosome interface, in which the lipid core, phospholipid monolayer, and interfacial proteins collectively enable interactions across a broad range of polarities. Although loading was assessed qualitatively, these observations reinforce the adaptability of oleosomes as carrier platforms and motivate future quantitative studies addressing loading capacity, release kinetics, and bioavailability.
Overall, this work establishes a coherent framework linking botanical origin to oleosome structure, stabilization mechanism, and functional potential under solvent-free conditions. Oleosomes have also been reported as natural alternatives to synthetic surfactant-based lipid nanoparticles in topical, oral, and transdermal applications, and the Generally Recognized as Safe (GRAS) status of their constituent oils supports the development of tunable interfaces for sustainable “green nanocarrier” design [19,20]. By delineating electrostatically stabilized seed oleosomes and sterically stabilized nut oleosomes as distinct formulation classes, the study provides a rational basis for source selection. It supports the development of oleosomes as sustainable alternatives to synthetic lipid carriers across various applications, including pharmaceutical, nutraceutical, cosmetic, and food.
5. Conclusions
This study compared oleosomes from nuts and seeds extracted in aqueous media under standardized conditions, showing that organic-solvent-free processing preserves their native triacylglycerol–phospholipid–protein architecture. Quantitative sizing, PDI, zeta potential, and SDS–PAGE analysis support the existence of two stabilization regimes: seed-derived oleosomes are smaller, protein-rich, and primarily electrostatically stabilized, whereas nut-derived oleosomes are larger, lipid-dense, and stabilized mainly by steric effects. This structure–function map links botanical origin to droplet size, charge, and stability, and can guide source selection by application.
Industrial relevance is broad. In foods, sub-500 nm, highly charged seed oleosomes are suitable for clear beverages, dairy alternatives, and the delivery of antioxidants or vitamins, with improved colloidal stability. In cosmetics, larger nut oleosomes enable the formulation of occlusive creams, masks, and depot formats with slower diffusion and a pleasant sensory feel. In pharmaceuticals and nutraceuticals, both classes offer biocompatible carriers for lipophilic actives, with tunable release characteristics achieved through adjustments in droplet size, interfacial protein density, and pH or ionic strength, thereby enhancing delivery and increasing bioavailability.
Manufacturing can be scaled via continuous or batch blending at low temperatures (approximately 4 °C), disc-stack or tubular centrifugation, aqueous washing, and freeze- or spray-drying, with quality control based on particle size, PDI, and zeta potential. Overall, oleosomes are promising, organic-solvent-free, bio-based carriers that can replace or reduce the use of synthetic carriers in food, cosmetic, and pharmaceutical applications.
Future studies will focus on quantitative loading capacity, release kinetics, and in vivo performance to further validate oleosomes as pharmaceutical nanocarriers.
Acknowledgments
The author, Marlon C. Mallillin III, is supported by a scholarship from the Department of Science and Technology—Science Education Institute (DOST-SEI). The author Shengnan Zhao is supported by a scholarship from the China Scholarship Council (CSC). The authors also wish to thank Seohyeon Park for her technical assistance with aspects of the experimental work.
Abbreviations
The following abbreviations are used in this manuscript:
| BCA | Bicinchoninic acid |
| GRAS | Generally recognized as safe |
| kDa | Kilodalton |
| mV | Millivolt |
| NLC | Nanostructured lipid carriers |
| PDI | Polydispersity index |
| PPCO | Polypropylene copolymer |
| SDS–PAGE | Sodium dodecyl sulfate polyacrylamide gel electrophoresis |
| SLN | Solid lipid nanoparticles |
| TAG | Triacylglycerol |
Institutional Review Board Statement
Not applicable.
Informed Consent Statement
Not applicable.
Data Availability Statement
The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding authors.
Conflicts of Interest
The authors declare no conflicts of interest.
Funding Statement
This research received no external funding.
Footnotes
Footnote Group
References
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Associated Data
Data Availability Statement
The original contributions presented in this study are included in the article. Further inquiries can be directed to the corresponding authors.