https://orcid.org/0000-0002-9779-0357
Figures
Abstract
Globally, there is urgent search and research interest towards alternative sources of polyunsaturated omega-3 fatty acids (ω-3 FA) due to sustainability challenges faced by dependence on fish and plant-derived sources. Limited information exists on insects despite their promising potential. This work analyzed oils from seven insect species commonly consumed globally compared to commercial sesame oil using coupled gas chromatography-mass spectrometry (GC-MS) and liquid chromatography-mass spectrometry (LC-MS). Oil yields from insects ranged between 11.1 mg/g to 115.3 mg/g. Over 67 fatty acids, α-tocopherol [vitamin E] and eight flavonoids were identified. All insect oils contained ω-3 FA [i.e., α-linolenic acid (ALA), eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA)]. The fifth instar of desert locust [Schistocerca gregaria] had the highest levels of ALA (9.4 µg/mg), EPA (1.0 µg/mg), and DHA (3.1 µg/mg) compared to sesame oil with ALA (7.9 µg/mg) only. Polyunsaturated fatty acids were dominant in sesame oil. Over 50% of monounsaturated fatty acids were present in S. gregaria, Spodoptera frugiperda and Bactrocera dorsalis. Gryllus bimaculatus oil contained mostly saturated fatty acids (51.1%). The levels of α-Tocopherol [vitamin E] were significantly higher in all the insect oils except S. frugiperda. Flavonoids such as quercetin and luteolin were abundantly present in S. gregaria and G. bimaculatus oils. These insects are widely consumed across the globe and can be farmed using fewer resources. Thus, widening the research scope would offer new opportunities to advance use of edible insects to address interconnected nutritional and health challenges.
Citation: Cheseto X, Tanga CM (2026) Omega-3 fatty acids and bioactives in aqueous-extracted oils from seven edible insects. PLoS One 21(8): e0356807. https://doi.org/10.1371/journal.pone.0356807
Editor: Amel Mohamed El Asely, Benha University, EGYPT
Received: January 13, 2026; Accepted: August 7, 2026; Published: August 26, 2026
Copyright: © 2026 Cheseto, Tanga. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The minimal data set is available at Figshare via https://doi.org/10.6084/m9.figshare.32583804.
Funding: The authors gratefully acknowledge the financial support from Bill & Melinda Gates Foundation (INV-032416); European Commission (NESTLER project: 101060762 and INNOECOFOOD project: 101136739); the Curt Bergfors Foundation Food Planet Prize Award; the Swedish International Development Cooperation Agency (Sida); the Swiss Agency for Development and Cooperation (SDC); the Australian Centre for International Agricultural Research (ACIAR); the Government of Norway; the German Federal Ministry for Economic Cooperation and Development (BMZ); and the Government of the Republic of Kenya. The views expressed herein do not necessarily reflect the official opinion of the donors. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Omega-3 fatty acids (ω-3 FA), such as α-linolenic acid (ALA), eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), are essential dietary lipids known to support heart health, brain function, mood balance, and may offer anti-diabetic and anti-cancer benefits [1–3]. Their primary sources have traditionally included fatty fish, flaxseeds, chia seeds, and walnuts [4–6]. However, concerns about overfishing, environmental degradation, and resource-intensive crop production are raising concerns about the long-term sustainability of these foods. This has intensified the search for alternative, reliable sources of lipids and ω-3 FA [7,8].
Beyond ω-3 FAs, dietary lipids can serve as carriers of important bioactive compounds, including fat-soluble vitamins and phytochemicals, which contribute to their overall nutritional and functional value [9,10]. Vitamin E (tocopherols), for instance, acts as a lipid-soluble antioxidant that protects cellular components from oxidative damage and supports immune function [11]. In edible insect oils, α-tocopherol concentrations have been reported to range from approximately 2–11 µg/mg of oil, with species such as Schistocerca gregaria, Ruspolia differens, and Gryllus bimaculatus showing particularly high levels [12–15]. Similarly, flavonoids, including quercetin, luteolin, and kaempferol, exhibit antioxidant, anti-inflammatory, and cardioprotective effects, with reported levels in insect oils ranging from 0.2 to 1.3 µg/mg of oil depending on the species [14,15]. Despite their recognized importance, these bioactive constituents remain underexplored in many alternative lipid sources.
In this context, edible insects have emerged as promising and sustainable alternatives. They are characterized by low environmental footprint, efficient feed conversion, and ability to thrive on waste streams [16–18]. Beyond their well-known protein content, many insects’ species contain oils naturally enriched with ω-3 FA and other bioactive compounds [8,19], positioning them as an underexplored yet potentially valuable contributor to future dietary lipid sources
While insect protein has been extensively studied and recognized for its high nutritional value, insect oils, the second most abundant nutrient in many species, have largely been overlooked [19,20]. These oils represent valuable byproducts of protein processing and offer considerable potential for application in food, feed, biofuel production, pharmaceuticals, and cosmetics [14,21,22]. Despite this promise, the nutraceutical properties of insect oils, particularly their content of ω-3 FA content, vitamin E and flavonoids remain poorly characterized.
In this study, we assessed the fatty acid and bioactive compound profiles of oils extracted from seven edible insect species representing three orders (Orthoptera, Diptera, and Lepidoptera) commonly consumed across Africa. Oils were obtained using an aqueous extraction method selected for its affordability, environmental benefits, and safety. We quantified and compared oil yields and analyzed (i) fatty acid compositions including essential ω-3 FA, (ii) vitamin E content, and (iii) flavonoid profiles. For B. dorsalis adult, B. dorsalis L3 and C. capitata adult, vitamin E and flavonoid analyses were not performed due to limited sample availability. All results were compared with commercial sesame oil (Sesamum indicum), a locally available in Kenya, where its widely cultivated in (western, coastal, semi-arid regions),and primarily used as seed oil valued for its nutritional, economic importance and with a well-characterized profile of fatty acids, vitamin E, and bioactive compounds, making it a suitable reference for evaluating the functional potential of edible insect-derived oils [23,24]. It is also more affordable than most imported oils like sunflower, corn, and palm oils, which are not only costly but also frequently subject to adulteration, much like high-end extra virgin olive oil [25].
Materials and methods
Insects
In this study, insects representing three orders (Orthoptera, Diptera, and Lepidoptera), were selected to expand our existing database of characterized oils. Previous analyses from our group include oils from the adult desert locust (Schistocerca gregaria), the adult African bush-cricket (Ruspolia differens) and termite alates (Macrotermes spp) [14,15]. The newly selected species were reared on standardized diets using established protocols and production systems. Insects were harvested at defined developmental stages, as outlined in Table 1. All rearing and sample preparation were conducted at the International Centre of Insect Physiology and Ecology (icipe) in Nairobi, Kenya.
Plant oil
A 500 mL bottle of cold-pressed, unrefined sesame oil (Chepper industries, Nakuru, Kenya) was purchased from a local supermarket in Nairobi, Kenya.
2.3. Extraction of insect oils
For each insect species and the corresponding developmental stage targeted (Table 1), oil was extracted 0.5 kg of wet insect biomass using an aqueous extraction method adapted from previously described methods [14,26]. The insects were killed by freezing at −80 °C for 3 h followed by homogenization in 1000 mL of distilled deionized water for 5 min. The slurry was transferred to a 1.5 l conical flask and heated at 80 °C for 3 h with gentle magnetic stirring, followed by filtration through a double-layered gauze cloth.
The filtrate was transferred into a separating funnel and left undisturbed overnight to allow phase separation. The upper layer was collected, mixed with 0.9% NaCl solution (20 mL) (Sigma-Aldrich, St. Louis, MO, USA) and centrifuged at 14000 rpm for 10 min. The resulting supernatant (insect oil), recovered, weighed and stored at −20 °C until analysis. Three independent extractions were performed for each species using separate insect batches. Oil yield was calculated using the following equation:
GC-MS analysis of fatty acids
Fatty acid profiles of the insect oils and sesame oil were determined after derivatization to fatty acid methyl esters (FAMEs) using a sodium methoxide (15 mg/mL in dry methanol; Sigma-Aldrich, St. Louis, MO, USA). Briefly, 100 mg of each sample was mixed with 500 µL of the sodium methoxide solution, vortexed for 10 s, sonicated for 10 min, and then incubated at 60 °C for 1 h. The reaction was terminated by adding 100 μL of deionized water, followed by 10 s vortexing. FAMEs were extracted with 1 mL of GC-grade hexane, centrifuged at 14000 rpm for 5 min, and the organic layer was dried over anhydrous Na2SO4(s) (Sigma-Aldrich, St. Louis, MO, USA). An aliquot (1 µL) of each extract was analyzed on a GC-MS system (7890A GC coupled to a 5975 C mass selective detector, Agilent Technologies, Inc., Santa Clara, CA, USA) fitted with a HP5 MS low bleed capillary column (30 m × 0.25 mm i.d., 0.25 µm film thickness; J&W, Folsom, CA, USA. Helium was used as the carrier gas at a flow rate of 1.25 mL/min. The oven temperature was ramped from 35 °C to 285 °C, while the MS ion source and quadrupole were maintained at 230 °C and 180 °C, respectively. A filament delay of 3.3 min was applied. For external quantification, a calibration curve was generated using an authentic methyl octadecenoate standard (≥ 99%, Sigma-Aldrich, St. Louis, MO, USA) across a concentration range of 0.2–125 ng/μL under identical conditions. Compound identities were confirmed based on retention times, authentic standards (where available), and comparison of spectral data with reference spectra from Adams, Chemecol, NIST 05, 08, and 11 libraries. All insect oil samples were analyzed in triplicate, with each replicate derived from an independent extraction batch. FAME concentrations were expressed as µg/mg of oil.
Extraction and GC-MS analysis of Vitamin E (α-tocopherol)
Analysis of samples for vitamin E (α-tocopherol), was performed following a previously established protocol [14] with minor modifications. Briefly, 300 mg of insect or plant oil was placed into a 10 mL glass vial, after which 5 mL of a solvent mixture (hexane: methanol: deionized water; 2:1:2, v/v) was added. The mixture was vortexed for 10 s, sonicated for 30 min, and centrifuged at 14,000 rpm for 5 min. The resulting supernatant was dried over anhydrous Na2SO4(s), evaporated to dryness under gentle stream of N2(g) and any residual fatty acids were derivatized using above mentioned procedure (GC-MS analysis of fatty acids). All analyses were performed in triplicate, with each replicate prepared from an independent insect oil batch.
Extraction and LC-MS analysis of Flavonoids
Flavonoids were extracted following a previously established protocol [14], with minor modifications. Briefly, 300 mg of each insect or plant oil sample was transferred into a 2 mL Eppendorf tube containing 1,000 µL of LC-MS-grade acetonitrile (Sigma-Aldrich, St. Louis, MO, USA). The mixture was vortexed for 10s, sonicated for 30 min, and centrifuged at 14,000 rpm for 15 min. The resulting acetonitrile-soluble supernatant was filtered through Whatman No. 42 filter paper and collected into a 2 mL autosampler vial for LC-MS analysis.
Chromatographic separation was performed on an Agilent 1260 Infinity HPLC system (Agilent Technologies, Palo Alto, CA, USA) coupled to an Agilent 6120 Single Quadrupole MS detector. Separation done on a ZORBAX SB-C18 column (4.6 × 250 mm, 3.5 µm) maintained at 40 °C. The mobile phases consisted of water (A) and acetonitrile (B), each containing 0.01% formic acid. The gradient program was as follows: 0.01 min, 5% B; 0.01–5 min, 5% B; 5–10 min, 5–20% B; 10–15 min, 20% B; 15–20 min, 20–80% B; 20–25 min, 80% B; 25–30 min, 80–100% B; 30–35 min, 100–37 min, 100–5% B; and 37–42 min, 5% B. The flow rate was 0.5 mL/min, and the injection volume was 3 µL.
The single-quadrupole MS was operated in two separate acquisition methods, one in ESI positive mode and the other in ESI negative mode, each scanning across an m/z range of 100–2,000. The instrument was set with the following parameters: capillary voltage, 3.0 kV; cone voltage, 30 V; extractor voltage, 5 V; RF voltage, 0.5 V; source temperature, 110 °C; desolvation temperature, 380 °C; and desolvation nitrogen flow, 400 L/h. Mass spectra were recorded for each chromatographic peak, and compounds were tentatively identified by comparing the spectra with reference databases, including METLIN, ChemSpider, ChemCalc, and the NIST MS/MS libraries.
Quercetin, luteolin, kaempferol, rutin, and apigenin were confirmed through co-injection with high-purity authentic standards (≥ 90–98% purity; Sigma-Aldrich, St. Louis, MO, USA). For quantification, a linear calibration curve (peak area vs. concentration, 1–100 ng/µL) was established for each standard. All measurements were performed in triplicate, with each replicate prepared from a separate batch of extracted oil.
Data analysis
Data was first entered into Microsoft Excel, and all statistical analyses were performed using R software version 3.5.0 [27]. To illustrate the relative distribution of fatty acids across the insect and plant oils, 100% stacked bar charts were created in Excel. Differences in the proportions of saturated, monounsaturated, and polyunsaturated fatty acids were assessed using Chi-square tests. Concentrations of individual fatty acids, flavonoids, oil yields, and vitamin E were compared using one-way ANOVA followed by the Student–Newman–Keuls (SNK) post hoc test. When the assumption of equal variance was violated, Welch’s ANOVA was applied. Normality and homogeneity of variance were checked using the Shapiro–Wilk test (P > 0.05) and Levene’s test (P > 0.05), respectively. Statistical significance was defined at P < 0.05. Flavonoid concentrations were reported in ng/g of oils as Mean ± standard error (SE) based on triplicate determinations. Data used to generate all the graphs are available from Figshare [28].
Results
Insect oil yields
Oil yields varied significantly across the seven insect species studied (Fig 1). Yields ranged from 11.1 ± 1.63 mg/g in C. capitata to 115.3 ± 1.19 mg/g in S. icipe. The two cricket species (S. icipe and G. bimaculatus) produced approximately twice the oil of S. gregaria L5 among the orthopterans. The fall armyworm (S. frugiperda) yielded 90.0 ± 4.71 mg/g, comparable to the crickets and 1.5 times higher than S. gregaria L5. Fruit flies (B. dorsalis and C. capitata) showed the lowest yields.
Error bars indicate the standard error. Bar graph with different letters is significantly different from each other (ANOVA followed by Student-Neuman-Keul’s (SNK’s) post-hoc test; p < 0.05, n = 3).
Fatty acid composition of insect and plant oils
Table 2 presents the identified FAMEs in insects and plant oils. A total of 66 FAMEs were identified across the insect and plant oils analyzed, with sesame oil containing 44 FAMEs. Among the insect oils, S. gregaria L5 and G. bimaculatus registered the highest total number of FAMEs (53 each), followed by S. icipe (51). In contrast, oils from B. dorsalis (adult, 41; L3, 40) and C. capitata (42) had FAME profiles comparable to sesame oil, while S. frugiperda contained 34 detectable FAMEs.
The detected fatty acids comprised 35 SFA, 16 MUFA, and 15 PUFA. SFA distribution was as follows: sesame (19), S. gregaria L5 (28), S. icipe (26), G. bimaculatus (28), S. frugiperda (17), B. dorsalis adult (20), L3 (20), and C. capitata (21). MUFA counts were sesame (14), S. gregaria L5 (12), S. icipe (12), G. bimaculatus (12), S. frugiperda (6), B. dorsalis adult (12), L3 (12), and C. capitata (12). PUFA counts were sesame (11), S. gregaria L5 (13), S. icipe (13), G. bimaculatus (13), S. frugiperda (11), B. dorsalis adult (9), L3 (8), and C. capitata (9).
Several fatty acids were detected exclusively in either insect or plant oils. Sixteen SFA were unique to insect oils, whereas one SFA (methyl 4,8-dimethylnonanoate) was exclusive to sesame oil. Two MUFA (methyl (10Z)-heptadecenoate and methyl (6Z)-octadecenoate) occurred only in insect oils, while two MUFA (methyl (5Z)-dodecenoate and methyl 9-heptadecenoate) were found solely in sesame oil. Additionally, four PUFA were detected exclusively in insect oils.
The relative proportions of SFA, MUFA, and PUFA across insect and plant oils are shown (Fig 2). Insect oils displayed generally similar percentages of these fatty acid classes, whereas sesame oil contained a higher proportion of PUFA. Among essential PUFAs, linoleic acid (LA; 18:2 n-6) ranged from 5.8 µg/mg in C. capitata adult to 49.8 µg/mg in S. icipe. S. gregaria L5 contained 40.5 µg/mg of LA, while all orthopteran insect oils (S. icipe, S. gregaria L5, and G. bimaculatus) exceeded 17.1 µg/mg.
Difference in the proportions of SFAs, MUFAs and PUFAs were assessed using the Chi-square test. Statistical significance is indicated as follows ** p < 0.01, *** p < 0.001.
ω-3 FA were detected at varying levels across the insect and plant oils (Table 2). ALA was most abundant in S. gregaria L5 (9.4 µg/mg), followed by sesame oil (7.9 µg/mg). EPA was highest in S. gregaria L5 (25.2 µg/mg), with lower concentrations in G. bimaculatus (1.4 µg/mg) and B. dorsalis L3 (1.3 µg/mg). (DHA) also peaked in S. gregaria L5 (3.2 µg/mg). The ω-6/ω-3 ratios in insect oils ranged from 1.2 to 3.9.
Vitamin E
Only α-tocopherol was detected across all oil samples, and its concentration varied among the different sources. S. gregaria L5 andS. icipe recorded higher α-tocopherol levels than sesame oil and S. frugiperda L5 (Fig 3). G. bimaculatus showed an intermediate α-tocopherol concentration (1.83 mg/ 100 g oil), which was numerically higher than that of sesame oil (1.00 mg/100 g oil) but did not differ significantly from either the sesame oil/ S. frugiperda L5 or the S. gregaria L5/S. icipe group.
On each bar graph, the minimum and maximum values of all the data are represented by the ends of the standard error whiskers. Bar graphs with different lower-case letters are significantly different from each other (ANOVA followed by Student-Neuman-Keul’s (SNK’s) post-hoc test; p < 0.05, n = 3).
Flavonoids
Eight flavonoids were identified with distinct qualitative distributions across the different oils (Table 3). Sesame oil contained three flavonoids; S. gregaria L5 (5); S. icipe (3); and G. bimaculatus (3). Sesame oil uniquely contained high levels of hesperidin and sesamoside, while S. gregaria L5 had apigenin and orientin in addition to the three flavonoids common to all insect oils.
Discussion
The extraction method strongly influenced overall oil yields and fatty acid profiles in this study. We employed aqueous extraction, intentionally selected for its safety, affordability, and ability to preserve thermolabile bioactive compounds such as flavonoids and vitamin E [14,26]. This method efficiently recovers polar and amphipathic lipids, including phospholipids, glycolipids, and lysophospholipids, but has limited efficiency for non-polar storage lipids such as triacylglycerols, which are the main carriers of SFAs [29,30]. Therefore, the SFAs reported here primarily reflect the polar lipid fraction rather than the total insect SFA content. Comparisons with previous studies using conventional extraction techniques (e.g., Soxhlet, Folch, or ultrasound-assisted extraction) or solvent-based methods should therefore be interpreted in this context [31]. Future studies incorporating multiple extraction methods would allow comprehensive characterization of the total lipid fraction in edible insects.
Oil yields differ widely among the seven insect species, reflecting variations in developmental stage, species-specific physiology, and extraction method. For example, the desert locust has previously been reported to yield 37 mg/g of oil at adult stage produced nearly twice this amount in the 5th instar using aqueous extraction [14]. Crickets (S. icipe and G. bimaculatus) exhibited the highest oil yields among Orthoptera, whereas the fruit fly species (B. dorsalis and C. capitata) yielded considerably less oil, consistent with their smaller body size and lower energetic requirements [32]. The fall armyworm (S. frugiperda) yielded moderate amounts of oil, highlighting its potential as an abundant and cost-effective biomass source [33,34].
Dietary composition played a critical role in shaping the fatty acid and bioactive compound profiles of insect oils. Desert locusts fed a mixed diet of wheat seedlings, maize leaves, and wheat bran, rich in essential fatty acid precursors and natural antioxidants, exhibited elevated levels of ω-3 FA (ALA, EPA, DHA), α-tocopherol, and flavonoids [35,36]. Crickets reared on complex high-energy diets including wheat bran, soybean, fish offal, pumpkin leaf, and carrot similarly demonstrated enhanced lipid accumulation and bioactive content [37]. In contrast, fall armyworm, fed exclusively on maize leaves, showed a more restricted fatty acid diversity and lower flavonoid content, illustrating the influence of dietary inputs on nutritional quality. Flavonoids such as quercetin, luteolin, kaempferol, and apigenin detected in insect oils likely reflect the presence of these compounds in plant-based diets and may be sequestered or metabolically transformed by insects’ ingestion [38,39]. These findings highlight the potential for targeted dietary manipulation during rearing to optimize the nutritional and functional properties of edible insect oils [40–42].
GC-MS analysis confirmed that edible insect oils contain a nutritionally and functionally significant diversity of SFA, MUFA, and PUFA. S. gregaria L5, G. bimaculatus, and S. icipe showed the highest number of FAMEs, consistent with earlier findings on the biochemical richness of orthopteran oils [14,43]. Conversely, B. dorsalis and C. capitata showed fewer FAMEs, like sesame oil. Interestingly, sesame oil displayed a greater diversity of FAMEs than previously documented, suggesting improved analytical sensitivity in this study [44].
Saturated fatty acids were the dominant class (~ 53%), consistent with their roles in insect physiology, including energy storage, membrane integrity, reproduction, and adaptation [45]. Palmitic acid (C16:0), a major SFAs, contributes to cuticle structure and protection [46]. Lauric acid (C12:0), detected in all species, offers antimicrobial protection and supports rapid energy mobilization. Targeted dietary manipulation and optimized rearing could increase the content of nutritionally or industrially valuable SFAs, such as lauric acid [47,48].
In humans, SFAs provide concentrated energy, support thermoregulation, and facilitate the absorption of fat-soluble vitamins, while also enhancing the texture and stability of formulated foods [19,49,50]. SFAs such as palmitic and stearic acid are widely used in cosmetics, soaps, emulsifiers, lubricants, polishes, and emerging biofuel technologies, emphasizing the commercial relevance of insect-derived lipids beyond food applications [51,52].
Unsaturated fatty acids, including MUFA and PUFA, are equally important in insects for maintaining membrane fluidity and for serving as precursors to pheromones and other signaling molecules [50]. In humans, they are known for their cardioprotective and metabolic benefits [1,53,54].
One of the most striking findings was the presence of all three major ω-3 FA, (ALA, EPA, and DHA) in insect oils, usually associated with marine sources (Table 4). S. gregaria L5 displayed particularly high EPA (2514 mg/100 g) and DHA (319 mg/100 g), levels comparable to or exceeding several commercial fish species (Table 4).
G. bimaculatus also contained noteworthy amounts. In contrast, plant oils, such as flaxseed, chia, and sesame, provide ALA but no DHA or EPA. Because human conversion of ALA to EPA and DHA is inefficient (<10%) [55], edible insects offer a more complete ω-3 FA profile, especially for regions with limited access to fish or marine oils. The elevated ω-3 FA levels in S. gregaria L5 may be influenced by ecological factors such as cannibalism, which warrants further study [56].
These findings position edible insects as credible, environmentally responsible alternatives to marine sources of ω-3 FA, particularly at a time when overfishing, habitat loss, and aquaculture resource demands are intensifying [63]. Their demonstrated health benefits, including cardioprotective, anti-inflammatory, neurocognitive, and dermatological effects [53,54], further support their integration into functional foods and nutraceutical formulations.
The ω-6/ω-3 ratios (1.2–3.9) recorded for insect oils fall within WHO recommendations (<5:1), suggesting favorable dietary potential [64,65]. Current intake recommendations call for 1.1–1.6 g/day ALA and 250–500 mg/day combined EPA + DHA for adults (higher for pregnancy/lactation) [66,67].
Beyond fatty acids, the analyzed insect oils were rich in micronutrients, notably α-tocopherol (vitamin E), which was abundant in several species. Vitamin E acts as a potent antioxidant, protecting cellular membranes from oxidative damage and supporting immune, neurological, and cardiovascular health [11,67]. The high values observed align with earlier reports in diverse insect taxa [13,68], indicating that edible insects can serve as a natural dietary source of vitamin E. The levels of vitamin E in insects are influenced by the rearing diet, with plant-derived substrates rich in tocopherols and other antioxidants contributing to higher concentrations [35–37] For example, S. gregaria L5, which fed on a combination of wheat seedlings, maize leaves, and wheat bran, exhibited higher vitamin E content compared to fall armyworm (S. frugiperda), reared solely on maize leaves, demonstrating the direct impact of dietary composition on micronutrient accumulation. This suggests that dietary manipulation during insect farming could optimize vitamin E content, enhancing the functional and nutraceutical quality of the oils.
Flavonoids including quercetin, luteolin, kaempferol, and apigenin were detected across species. In insects, flavonoids are typically sequestered from dietary plant sources, as insects generally cannot synthesize these compounds de novo [38,39]. The diversity and concentration of flavonoids observed in this study closely reflect the complexity and variety of substrates consumed, with species fed on mixed plant-based diets (e.g., G. bimaculatus and S. icipe) showing higher flavonoid diversity than species on more restricted diets. These polyphenols contribute antioxidant, anti-inflammatory, and metabolic regulatory benefits, potentially improving oxidative stability of the oils and providing additional health benefits upon consumption [69].
Flavonoids and vitamin E also protect insect tissues from oxidative stress, supporting survival, development, and reproductive fitness [29,70]. For humans, the combination of these bioactives with essential fatty acids enhances the nutritional and therapeutic potential of insect oils, positioning them as promising ingredients for functional foods, nutraceuticals, and cosmeceuticals.
To fully realize the commercial potential of edible insect oils, standardized analytical profiling of vitamin E, flavonoids, other phenolics, alkaloids, and measures of antioxidant capacity and quality indices is essential. Such profiling will enable regulatory compliance, quality assurance, and targeted dietary optimization, ensuring that insect oils meet both nutritional and industrial standards. Future studies should evaluate diet-composition relationships, bioavailability, and stability of these bioactives to strengthen their applicability in food, feed, and pharmaceutical industries.
Conclusions
Edible insects represent a nutrient-dense and environmentally sustainable source of high-value oils rich in essential ω-3 FA, vitamin E, and diverse flavonoids. The substantial variation in oil yields and fatty acid composition across species and developmental stages underscores opportunities to optimize rearing and extraction strategies. The 5th instar desert locust (S. gregaria) was particularly notable for providing appreciable levels of ALA, EPA, and DHA, offering a more complete ω-3 FA profile than common plant oils such as sesame. Other species also contributed valuable ω-3 FA, vitamin E, and antioxidant flavonoids, strengthening their potential as functional ingredients. These attributes position insect-derived oils as viable, eco-friendly complementary to conventional plant and marine fats, with broad applications in food, feed, and industrial sectors. Continued research comparing additional plant oils, optimizing production systems, and expanding bioactive profiling will be essential for fully realizing the potential of edible insects within sustainable nutrition and bioeconomy frameworks.
Acknowledgments
We thank Jactone Ooko, Erick Isaiah, Shem Ondiek and Mercy Kibii for raising and supplying the colonies of the various insects, Stella Wachira for oil extraction and Onesmus Wanyama for technical support. We also thank Dr. Isaac Osuga for critical review of the manuscript.
References
- 1. Holub BJ. Clinical nutrition: 4. Omega-3 fatty acids in cardiovascular care. CMAJ. 2002;166(5):608–15. pmid:11898942
- 2. Shahidi F, Ambigaipalan P. Omega-3 polyunsaturated fatty acids and their health benefits. Annu Rev Food Sci Technol. 2018;9:345–81. pmid:29350557
- 3. Mori TA. Marine OMEGA-3 fatty acids in the prevention of cardiovascular disease. Fitoterapia. 2017;123:51–8. pmid:28964873
- 4. Bhatt DL, Budoff MJ, Mason RP. A revolution in omega-3 fatty acid research. J Am Coll Cardiol. 2020;76(18):2098–101. pmid:33121717
- 5. Valerio-Coronel D, Sánchez-Tapia M, Torres-y Torres N. Beneficial effects of seeds and nuts. Rev REDNUTRICIÓN. 2021;12:788–92.
- 6. Simopoulos AP. Omega‐3 fatty acids in wild plants, nuts and seeds. Asia Pac J Clin Nutr. 2002;11(s6).
- 7. Li Y, Xiang N, Zhu Y, Yang M, Shi C, Tang Y, et al. Blue source-based food alternative proteins: exploring aquatic plant-based and cell-based sources for sustainable nutrition. Trends in Food Sci Tech. 2024;147:104439.
- 8. Lange KW, Nakamura Y. Edible insects as future food: chances and challenges. J Future Foods. 2021;1(1):38–46.
- 9. Patted PG, Masareddy RS, Patil AS, Kanabargi RR, Bhat CT. Omega-3 fatty acids: a comprehensive scientific review of their sources, functions and health benefits. Futur J Pharm Sci. 2024;10(1).
- 10. Xu R, Molenaar AJ, Chen Z, Yuan Y. Mode and mechanism of action of omega-3 and omega-6 unsaturated fatty acids in chronic diseases. Nutrients. 2025;17(9):1540. pmid:40362847
- 11. Aggarwal BB, Sundaram C, Prasad S, Kannappan R. Tocotrienols, the vitamin E of the 21st century: its potential against cancer and other chronic diseases. Biochem Pharmacol. 2010;80(11):1613–31. pmid:20696139
- 12. Adámková A, Mlček J, Kouřimská L, Borkovcová M, Bušina T, Adámek M, et al. Nutritional potential of selected insect species reared on the island of Sumatra. Int J Environ Res Public Health. 2017;14(5):521. pmid:28498340
- 13. Murugu DK, Onyango AN, Ndiritu AK, Osuga IM, Xavier C, Nakimbugwe D, et al. From farm to fork: crickets as alternative source of protein, minerals, and vitamins. Front Nutr. 2021;8:704002. pmid:34447775
- 14. Cheseto X, Baleba SBS, Tanga CM, Kelemu S, Torto B. Chemistry and sensory characterization of a bakery product prepared with oils from African edible insects. Foods. 2020;9(6):800. pmid:32570724
- 15. Cheseto X, Ochieng BO, Subramanian S, Tanga CM. Unravelling the nutritional and health benefits of marketable winged termites (Macrotermes spp.) as sustainable food sources in Africa. Sci Rep. 2024;14(1):9993. pmid:38693201
- 16. Ordoñez-Araque R, Egas-Montenegro E. Edible insects: a food alternative for the sustainable development of the planet. Int J Gastron Food Sci. 2021;23:100304.
- 17. Halloran A, Flore R, Vantomme P, Roos N. The role of edible insects in diets and nutrition in East Africa. In: Halloran A, Flore R, Vantomme P, Roos N, editors. Edible insects in sustainable food systems. Cham: Springer International Publishing; 2018. 1–479.
- 18. van Huis A. Edible insects are the future?. Proc Nutr Soc. 2016;75(3):294–305. pmid:26908196
- 19. van Huis A. Insects as food and feed, a new emerging agricultural sector: a review. JIFF. 2020;6(1):27–44.
- 20. Dreassi E, Cito A, Zanfini A, Materozzi L, Botta M, Francardi V. Dietary fatty acids influence the growth and fatty acid composition of the yellow mealworm Tenebrio molitor (Coleoptera: Tenebrionidae). Lipids. 2017;52(3):285–94. pmid:28083781
- 21. Brogan EN, Park Y-L, Shen C, Matak KE, Jaczynski J. Characterization of lipids in insect powders. LWT. 2023;184:115040.
- 22. Smetana S, Leonhardt L, Kauppi SM, Pajic A, Heinz V. Insect margarine: processing, sustainability and design. J Clean Prod. 2020;264:121670.
- 23. Pathak N, Rai AK, Kumari R, Thapa A, Bhat KV. Sesame crop: an underexploited oilseed holds tremendous potential for enhanced food value. Agric Sci. 2014;5:519–29.
- 24. Koitilio CE, Kinyua M, Kiplagat O, Kimno K. Sesame production characteristics and challenges in western Kenya. African J Edu Sci Tech. 2022;7.
- 25. Nanou E, Pliatsika N, Couris S. Rapid authentication and detection of olive oil adulteration using laser-induced breakdown spectroscopy. Molecules. 2023;28(24):7960. pmid:38138450
- 26. Tzompa-Sosa DA, Yi L, van Valenberg HJF, van Boekel MAJS, Lakemond CMM. Insect lipid profile: aqueous versus organic solvent-based extraction methods. Food Research Int. 2014;62:1087–94.
- 27.
R Core Team, R Development Core Team. A language and environment for statistical computing. Vienna, Austria: R Foundation for statistical computing; 2018.
- 28. Cheseto X, Tanga MC. Omega-3 fatty acids and bioactives in aqueous-extracted oils from seven edible insects. Figshare; 2026.
- 29. Saini RK, Prasad P, Shang X, Keum Y-S. Advances in lipid extraction methods-a review. Int J Mol Sci. 2021;22(24):13643. pmid:34948437
- 30. Genot C. Purification of lipids by cold-acetone precipitation. 2024. 31–6.
- 31. Pranil T, Moontree T, Moongngarm A, Loypimai P. Ultrasound-assisted extraction of edible cricket oils from Gryllus bimaculatus and Acheta domesticus. Food Measure. 2024;18(7):6145–55.
- 32. Chatterjee N, Perrimon N. What fuels the fly: energy metabolism in Drosophila and its application to the study of obesity and diabetes. Sci Adv. 2021;7(24):eabg4336. pmid:34108216
- 33. Sisay B, Sevgan S, Weldon CW, Krüger K, Torto B, Tamiru A. Responses of the fall armyworm (Spodoptera frugiperda) to different host plants: Implications for its management strategy. Pest Manag Sci. 2023;79(2):845–56. pmid:36301535
- 34. Emilly AO, Arnold W, Calleb OO, Ben M. A comparative evaluation of nutrient content of fall armyworm (Spodoptera frugiperda) larvae to other chicken feeds. Afr J Agric Res. 2022;18:27–34.
- 35. Millward DJ. The nutritional value of plant-based diets in relation to human amino acid and protein requirements. Proc Nutr Soc. 1999;58(2):249–60. pmid:10466163
- 36. das Graças Costa E, de Souza PM. Introduction to cereals. Cereal-based food products. Cham: Springer International Publishing; 2023. 1–24.
- 37. Straub P, Tanga CM, Osuga I, Windisch W, Subramanian S. Experimental feeding studies with crickets and locusts on the use of feed mixtures composed of storable feed materials commonly used in livestock production. Animal Feed Sci Tech. 2019;255:114215.
- 38. Shen N, Wang T, Gan Q, Liu S, Wang L, Jin B. Plant flavonoids: classification, distribution, biosynthesis, and antioxidant activity. Food Chem. 2022;383:132531. pmid:35413752
- 39. Dias MC, Pinto DCGA, Silva AMS. Plant flavonoids: chemical characteristics and biological activity. Molecules. 2021;26(17):5377. pmid:34500810
- 40. Li J, Deng J, Deng X, Liu L, Zha X. Metabonomic analysis of silkworm midgut reveals differences between the physiological effects of an artificial and mulberry leaf diet. Insects. 2023;14(4):347. pmid:37103160
- 41. Mahmoud MA, Abotaleb AO, Zinhoum RA. Evaluation of various diets for improved growth, reproductive and nutritional value of the yellow mealworm, Tenebrio molitor L. Sci Rep. 2025;15(1):15656. pmid:40325124
- 42. Muzzatti MJ, Harrison SJ, McColville ER, Brittain CT, Brzezinski H, Manivannan S, et al. Applying nutritional ecology to optimize diets of crickets raised for food and feed. R Soc Open Sci. 2024;11(12):241710. pmid:39635150
- 43. Murugu DK, Onyango AN, Ndiritu AK, Nyangena DN, Osuga IM, Cheseto X, et al. Physicochemical properties of edible cricket oils: implications for use in pharmaceutical and food industries. Future Foods. 2024;9:None. pmid:38932931
- 44. Hama JR. Comparison of fatty acid profile changes between unroasted and roasted brown sesame (Sesamum indicum L.) seeds oil. Int J Food Prop. 2016;20(5):957–67.
- 45. Kolobe SD, Manyelo TG, Malematja E, Sebola NA, Mabelebele M. Fats and major fatty acids present in edible insects utilised as food and livestock feed. Vet Anim Sci. 2023;22:100312. pmid:37736572
- 46. Kaczmarek A, Boguś M. The metabolism and role of free fatty acids in key physiological processes in insects of medical, veterinary and forensic importance. PeerJ. 2021;9:e12563. pmid:35036124
- 47. Ewald N, Vidakovic A, Langeland M, Kiessling A, Sampels S, Lalander C. Fatty acid composition of black soldier fly larvae (Hermetia illucens) - Possibilities and limitations for modification through diet. Waste Manag. 2020;102:40–7. pmid:31655329
- 48. Belghit I, Waagbø R, Lock EJ, Liland NS. Insect-based diets high in lauric acid reduce liver lipids in freshwater Atlantic salmon. Aquac Nutr. 2019;25:343–57.
- 49. West S, Monteyne AJ, van der Heijden I, Stephens FB, Wall BT. Nutritional considerations for the vegan athlete. Adv Nutr. 2023;14(4):774–95. pmid:37127187
- 50. Gede I, Aditama AS, Lorensia A, Virlando Suryadinata R, Raharjo DN. Association between body mass index and omega-3 fatty acid food intake. TMJ. 2022;45.
- 51. Adegbe AA, Larayetan RA, Omojuwa TJ. Proximate analysis, physicochemical properties and chemical constituents characterization of Moringa oleifera (Moringaceae) seed oil using GC-MS analysis. Am J Chem. 2016;6:23–8.
- 52. Franco A, Salvia R, Scieuzo C, Schmitt E, Russo A, Falabella P. Lipids from insects in cosmetics and for personal care products. Insects. 2021;13(1):41. pmid:35055884
- 53. Simopoulos AP. Omega-3 fatty acids in inflammation and autoimmune diseases. J Am Coll Nutr. 2002;21(6):495–505. pmid:12480795
- 54. Barry AR, Bishop KE, Pearson GJ, Koshman SL. Omega-3 fatty acids for cardiovascular disease prevention: a practice tool for pharmacists. Can Pharm J (Ott). 2022;155(3):169–74. pmid:35519086
- 55. Krzyzanowska J, Czubacka A, Oleszek W. Dietary phytochemicals and human health. In: Giardi MT, Rea G, Berra B, editors. Bio-farms for nutraceuticals: functional food and safety control by biosensors. Boston, MA: Springer US; 2010. 74–98.
- 56. Bazazi S, Buhl C, Hale JJ, Anstey ML, Sword GA, Simpson SJ, et al. Collective motion and cannibalism in locust migratory bands. Curr Biol. 2008;18(10):735–9. pmid:18472424
- 57. Rodriguez-Leyva D, Dupasquier CMC, McCullough R, Pierce GN. The cardiovascular effects of flaxseed and its omega-3 fatty acid, alpha-linolenic acid. Can J Cardiol. 2010;26(9):489–96. pmid:21076723
- 58. Ayerza R, Coates W. Chia: rediscovering a forgotten crop of the Aztecs. University of Arizona Press; 2022.
- 59. Callaway JC. Hempseed as a nutritional resource: an overview. Euphytica. 2004;140(1–2):65–72.
- 60. Crews C, Hough P, Godward J, Brereton P, Lees M, Guiet S, et al. Study of the main constituents of some authentic walnut oils. J Agric Food Chem. 2005;53(12):4853–60. pmid:15941326
- 61. Kris-Etherton PM, Harris WS, Appel LJ, American Heart Association. Nutrition Committee. Fish consumption, fish oil, omega-3 fatty acids, and cardiovascular disease. Circulation. 2002;106(21):2747–57. pmid:12438303
- 62. Gladyshev MI, Sushchik NN, Gubanenko GA, Makhutova ON, Kalachova GS, Rechkina EA, et al. Effect of the way of cooking on contents of essential polyunsaturated fatty acids in filets of zander. Czech J Food Sci. 2014;32(3):226–31.
- 63. Haque MM, Mahmud MN. Potential role of aquaculture in advancing sustainable development goals (SDGs) in Bangladesh. Aquac Res. 2025;2025:6035730.
- 64.
Fats and fatty acids in human nutrition. Report of an expert consultation. FAO Food Nutr Pap. 2010;91:1–166. pmid:21812367
- 65. Simopoulos AP. The omega-6/omega-3 fatty acid ratio: health implications. OCL. 2010;17(5):267–75.
- 66. Gebauer SK, Psota TL, Harris WS, Kris-Etherton PM. n-3 fatty acid dietary recommendations and food sources to achieve essentiality and cardiovascular benefits. Am J Clin Nutr. 2006;83(6 Suppl):1526S-1535S. pmid:16841863
- 67. Farhan Aslam M, Majeed S, Aslam S, Irfan JA. Vitamins: key role players in boosting up immune response-a mini review. Vitam Miner. 2017;06(01).
- 68. Finke MD. Complete nutrient content of four species of feeder insects. Zoo Biol. 2013;32(1):27–36. pmid:22689347
- 69. Mamta S, Saxena J, Nema R, Singh D, Gupta A. Phytochemistry of medicinal plants. J Pharmacogn Phytochem. 2013;1:168–82.
- 70. González-Tokman D, Villada-Bedoya S, Hernández A, Montoya B. Antioxidants, oxidative stress and reactive oxygen species in insects exposed to heat. Curr Res Insect Sci. 2025;7:100114. pmid:40519244