Animal studies were approved by the Norwegian Animal Research Authority (license number FOTS ID 30111). Male Wistar rats (Rattus Norvegicus), 5 weeks old, were purchased from Taconic (Denmark). Upon arrival, they were randomized and acclimatized for 1 week with unrestricted access to chow and water, under 12 h light/dark cycles at 22 ± 2 °C and 55 ± 5% humidity. At the start of the experiments, the rats were block-randomized to their respective interventions, with eight rats per group. The rats were fed low-fat diets providing 16% of total energy from fat (lard), 64% from carbohydrates, and 20% from protein. To these diets, 5% (w/w) of either CETO3^® (Grøntvedt Biotech AS, Norway) for the experimental group or soy oil for the control group was added, resulting in final diets with approximately 20% of energy from fat, 61% from carbohydrates, and 19% from protein. Feed was provided in fixed amounts, with leftovers weighed after 10 weeks, and weight gain was measured once weekly. A relatively high dietary inclusion level was chosen based on previous studies reporting inconsistent effects of monounsaturated fatty acids on blood lipid profiles. The oils were incorporated into the diets as a fixed proportion (5%) rather than as weight-adjusted doses (e.g., mg/kg body weight), following the feeding strategy established for this experiment. This approach may have resulted in some variations in individual intake due to differences in actual food consumption among animals. All animals survived the experiment. The rats were anesthetized with 2–5% isoflurane and decapitated, and blood was collected into EDTA tubes and centrifuged, with plasma stored at −80 °C. The organs were collected, weighed, snap-frozen, and stored at −80 °C.

Blood screening was performed at termination. Biochemical lipid analyses in rat fasting plasma were performed at Haukeland University Hospital using the Cobas 8,000 system (Roche Diagnostics), following standard laboratory protocols to measure triglycerides (TG), total cholesterol, LDL cholesterol, and high-density lipoprotein (HDL) cholesterol. Non-HDL cholesterol was calculated by subtracting HDL from total cholesterol. For hematology parameters, whole blood samples were analyzed at the Institute of Marine Research, Bergen, Norway, using a VetScan HM5 (Abaxis, Union City, CA, USA) to measure red blood cell count (RBC), hemoglobin (HGB), hematocrit (HCT), mean corpuscular volume (MCV), mean corpuscular hemoglobin (MCH), mean corpuscular hemoglobin concentration (MCHC), red cell distribution width (RDW), platelet count (PLT), mean platelet volume (MPV), platelet crit (PCT), platelet distribution width (PDW), white blood cell count (WBC), lymphocytes (LYM), monocytes (MON), and neutrophils (NEU).

The total FA composition in the liver and plasma was analyzed by Mitomega AS using ultrafast gas chromatography (UFGC) (Thermo Electron Corporation, Massachusetts, USA) (17). FA concentrations were expressed as percentages of total FAs by weight (wt%). The Omega-3 Index was defined as the sum of EPA and DHA, expressed as a percentage of the total FA content. The anti-inflammatory index was defined as the ratio of anti-inflammatory FAs (EPA, DHA) to pro-inflammatory FAs (C20:4n-6; arachidonic acid) (18).

The stearoyl-CoA desaturase (SCD) indexes were calculated using the product-substrate ratio, with SCD-18 as oleic acid/stearic acid and SCD-16 as palmitoleic acid/palmitic acid (19). The de novo lipogenesis index was calculated according to the following formula: palmitic acid/linoleic acid (19). To assess the degree of unsaturation of the FA pool, the double bond index (DBI) was calculated by summing the concentrations of FAs with one double bond, FAs with two double bonds multiplied by 2, FAs with three double bonds multiplied by 3, FAs with four double bonds multiplied by 4, FAs with five double bonds multiplied by 5, and FAs with six double bonds multiplied by 6, then this sum was divided by the total concentration of FAs.

The animals were fasted for 4 h with free access to water prior to the administration of 2 g/kg glucose by oral gavage. Tail vein blood samples were taken before administration and at 15, 40, 60, and 120 min post-administration, and blood glucose levels were measured using a Contour Next blood glucose meter (Ascensia Diabetes Care, Basel, Switzerland).

Human satellite cells were isolated from muscle biopsy samples of the musculus vastus lateralis (20). The isolation of satellite cells was performed based on the method of Wensaas et al. (21), with modifications described by Pettersen et al. (22). Human skeletal muscle biopsies were obtained after informed written consent and approval by the Regional Committee for Medical and Health Research Ethics South-East, Oslo, Norway (reference number: REK11959). The isolated skeletal muscle cells were cultured and proliferated in DMEM-GlutaMAX (5.5 mM glucose), supplemented with 10% FBS, HEPES (25 mM), gentamicin (50 ng/mL), penicillin (25 IU), streptomycin (25 μg/mL), amphotericin B (1.25 μg/mL), hEGF (10 ng/mL) from Thermo Fisher Scientific (Waltham, MA, US), dexamethasone (0.39 μg/mL), and 0.05% BSA from Sigma-Aldrich (St. Louis, MO, US). Differentiation of myoblasts into myotubes for 7 days was induced at 80–90% confluence by changing the medium to DMEM-GlutaMAX (5.5 mM glucose) supplemented with 2% FBS and 25 pM insulin (Actrapid^® Penfill^® 100 IE/mL from NovoNordisk (Bagsvaerd, Denmark)).

Human hepatoma cells, Huh7, purchased from ATCC (LGC Standards, Middlesex, UK), were expanded and maintained in Nunc™ Cell and Culture Treated Flasks with DMEM-GlutaMAX (5.5 mmoL/L glucose), supplemented with 10% FBS, HEPES (25 mmoL/L), penicillin (25 IU), streptomycin (25 μg/mL), and amphotericin B (1.25 μg/mL) at 37 °C in a humidified 5% CO₂ incubator until they reached 80–90% confluence.

To perform the substrate oxidation assays, approximately 12,000 Huh7 liver cells per well and 6,000 myoblasts per well were seeded in a 96-well Corning^® CellBIND^® tissue culture plate and grown until reaching 70–80% confluence. Myoblasts were further differentiated into myotubes for 7 days. Erucic acid and cetoleic acid were each dissolved in 0.1 M NaOH to a final concentration of 6 mM and subsequently conjugated with 2.4 mM fatty acid-free albumin (BSA) (ratio FA/BSA 2.5/1). The cells were treated with the erucic or cetoleic acid–BSA complexes for 24–48 h prior to the assays. Substrate oxidation assay was assessed by providing a radiolabeled (¹⁴C) substrate of interest and trapping the released ¹⁴CO₂, as described by Wensaas et al. (23). The Huh7 cells and myotubes were given the radiolabeled substrates, either [1-¹⁴C]oleic acid (0.5 μCi/mL, 100 μM) or D-[¹⁴C(U)]glucose (0.5 μCi/mL, 200 μM) from PerkinElmer (Boston, MA, US), in DPBS (with MgCl₂ and CaCl₂) supplemented with 10 mM HEPES and 10 μM BSA (both from Thermo Fisher Scientific). L-carnitine (1 mM) (Sigma Aldrich) was included in the assay medium for oleic acid oxidation. Respective amounts of the non-radiolabeled substrate were added to obtain the final concentrations of oleic acid. Following trapping, both the produced ¹⁴CO₂ and cell-associated (CA) radioactivity were measured using a 2,450 MicroBeta² liquid scintillation counter (PerkinElmer). Protein concentration in each well was determined using the Bio-Rad protein assay kit, allowing the normalization of ¹⁴CO₂ and CA data to cellular protein content. Complete substrate oxidation was represented by the measurement of ¹⁴CO₂, while uptake was calculated as the sum of ¹⁴CO₂ and CA. Safety handling of radioactive materials was performed according to regulations by the Norwegian Radiation and Nuclear Safety Authority (DSA).

Statistical analyses were performed using GraphPad Prism^® version 10.0. Data were presented as means ± standard deviations (SDs). For the in vivo experiment (n = 8 per group), the differences between the control group and the herring oil group receiving the CETO3^® supplement were assessed using unpaired two-tailed Student’s t-tests assuming equal variances. Unpaired two-tailed Student’s t-tests were also used for the in vitro experiments on the oxidation of oleic acid and glucose in myotubes (n = 10 for control; n = 5 for erucic acid and cetoleic acid) and Huh7 cells (n = 10 for control; n = 5 for erucic acid and cetoleic acid). A p-value of <0.05 was considered statistically significant.

To assess mechanisms that may contribute to cardiovascular disease risk, we investigated the mobilization of n-9 and n-11 fatty acids and their association with SCD-16 and SCD-18 activity. The consumption of CETO3^® changed the FA composition both in the liver and plasma (Supplementary Table 1) and increased the mobilization of n-9 long-chain MUFAs, reflected by C20:1n-9, C22:1n-9, C24:1n-9, and n-11 long-chain MUFAs, such as C20:1n-11 and C22:1n-11, both in the rat liver and plasma (p < 0.05; Figures 1A,B). It has been reported that n-11 long-chain MUFAs can only be derived from diets, whereas n-9 long-chain MUFAs can be de novo synthesized (24). The changes in the FA (C16:0/C18:2n-6) ratio increased both in the plasma and liver (p < 0.05), suggesting that the increased relative proportions of n-9 long-chain MUFAs in plasma are mediated by diet and de novo lipogenesis through the action of FA elongases on C18:1n-9 (Figure 2A). The relative plasma proportion of C18:1n-9 was reduced after the administration of the herring oil (p < 0.05; Supplementary Table 1).

It has been reported that long-chain MUFAs can cause transient lipidosis in some organs, which may be influenced by hepatic SCD activity (25). High hepatic SCD-16 activity, indicated by the C16:1n-7/C16:0 ratio, combined with high SCD-18 activity, indicated by the C18:1n-9/C18:0 ratio, in addition to a low C18:0/C16:0 ratio, has been associated with the progression of triglyceride mobilization in the liver (26). Interestingly, the hepatic C16:1n-7/C16:0 and C18:1n-9/C18:0 ratios, serving as proxies for assessing the activities of SCD-16 and SCD-18, and the C18:0/C16:0 ratio were not changed after CETO3^® supplementation (p > 0.05; Figures 2B–D).

Increased long-chain MUFAs from herring oil consumption were accompanied by reduced plasma levels of TG (−55%, p < 0.05) and total FA (−58%, p < 0.05) in the rats (Figure 3). The interference with cholesterol metabolism was confirmed, as the levels of total cholesterol (T. Chol), LDL, and non-HDL cholesterol were reduced by −41, −45%, and −56% (p < 0.05), respectively (Figure 3). In contrast, HDL levels were not changed (p > 0.05), while a significantly increased HDL/LDL cholesterol ratio (46%, p < 0.05) was observed following herring oil supplementation (Figure 3). Hence, CETO3^® treatment can reduce plasma lipids in rats.

Weight gain and the masses of fat tissues, liver, brain, heart, kidney, and testis were not significantly changed in the rats following herring oil administration (p > 0.05; Supplementary Figure 1). Moreover, plasma safety hematology parameters were not affected by CETO3^® treatment, except for an increased red cell distribution width (RDW) observed in the herring oil group (p < 0.05; Supplementary Table 3). In addition, glucose tolerance was not affected by herring oil supplementation (p > 0.05; Figure 4).

Furthermore, we aimed to investigate whether long-chain MUFAs can affect energy metabolism in cultured human myotubes and liver cells. In the myotubes, we observed increased uptake and oxidation of oleic acid with n-11 FAs (p < 0.05), whereas n-9 FAs only increased uptake (p < 0.05; Figures 5A,B). Interestingly, n-9 FAs also increased the uptake and oxidation of glucose in the myotubes (p < 0.05; Figures 5C,D). However, the uptake and oxidation of both glucose and oleic acid were not changed by n-9 and n-11 FAs in the cultured human liver cells (p > 0.05), except for an increased oxidation of OA by n-9 MUFAs (p > 0.05; Figures 5E–H).

To investigate the effects of long-chain MUFAs on energy metabolism in vivo, we utilized a rat model. We found that the levels of long-chain saturated fatty acids (C22:0, C23:0, and C24:0) decreased in both the liver and plasma (p < 0.05; Figures 6A,B), which may be due to the increased activity of peroxisomal beta-oxidation following CETO3^® supplementation.

Certain FAs and their ratios are important risk markers related to various diseases within the cardiometabolic syndrome, and reduced hepatic elongation of FA is related to fatty liver (18). In the present study, after herring oil supplementation, changes in hepatic elongases resulted in increased levels of n-3 PUFAs and reduced n-6 PUFAs in both plasma and liver, an increased n-3/n-6 ratio, a higher Omega-3 Index, and an elevated double bond index (p < 0.05; Figures 7A–E), accompanied by increased levels of both EPA and DHA (p < 0.05; Figures 8B,C). In general, CETO3^® treatment increased the relative proportion of EPA and DHA and reduced the relative proportion of n-6 FAs, except for C22:5n-6 (Supplementary Table 1). The anti-inflammatory FA indexes were strongly increased in both the liver and plasma (Figure 8A). This is mostly attributed to the increased proportion of EPA and DHA and the reduced proportion of arachidonic acid (p < 0.05; Figures 8B–D).