# Furan fatty acids supplementation in obese mice reverses hepatic steatosis and protects against cartilage degradation

Mathys Chaslin, Georges Maroun, Erwann Durand, Béatrice Bonafos, Said Assou, Jatuporn Chaiyut, Laurent Vaysse, Vincent Ferrer, Siriluck Liengprayoon, Thomas Brioche, et al.

► To cite this version:

Mathys Chaslin, Georges Maroun, Erwann Durand, Béatrice Bonafos, Said Assou, et al. Furan fatty acids supplementation in obese mice reverses hepatic steatosis and protects against cartilage degradation. Biomedicine and Pharmacotherapy, 2025, 187, 13p. 10.1016/j.biopha.2025.118072. hal-05051861

HAL Id: hal-05051861

https://hal.inrae.fr/hal-05051861v1

Submitted on 30 Apr 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# Furan fatty acids supplementation in obese mice reverses hepatic steatosis and protects against cartilage degradation

Mathys Chaslin<sup>a</sup>, Georges Maroun<sup>b</sup>, Erwann Durand<sup>c,d</sup>, Béatrice Bonafos<sup>a</sup>, Said Assou<sup>b,d</sup>, Jatuporn Chaiyut<sup>e</sup><sup>D</sup>, Laurent Vaysse<sup>f,g</sup>, Vincent Ferrer<sup>e,f,g</sup><sup>D</sup>, Siriluck Liengprayoon<sup>e</sup>, Thomas Brioche<sup>a</sup>, Laurence Pessemesse<sup>a</sup>, Mélanie Macart<sup>a</sup>, Christelle Bertrand-Gaday<sup>a,D</sup>, Yves-Marie Pers<sup>b,h</sup>, Charles Coudray<sup>a</sup>, Jean-Marc Brondello<sup>b,D</sup>, François Casas<sup>a,*f</sup><sup>1</sup>, Christine Feillet-Coudray<sup>a,*f</sup><sup>1,D</sup>

## ARTICLE INFO

**Keywords:**

Furan fatty acids

Obesity

Liver steatosis

Skeletal muscle

Osteoarthritis

RNA-sequencing

## ABSTRACT

Obesity is a major global health problem associated with numerous metabolic dysfunctions, an increased risk of developing Metabolic Associated Steatotic Liver Disease (MASLD) and osteoarthritis. Recently, we demonstrated that in Diet-induced-Obesity (DIO) mouse model, preventive furan fatty acids (FuFA-F2) supplementation, a natural compounds found in many foods, reduced the onset of metabolic disorders and increased muscle mass. Here, we aimed to determine whether a short FuFA-F2 supplementation is capable of providing beneficial health effects in obese mice, notably by reversing metabolic disorders and limiting cartilage degradation. 6-month-old obese C57BI/6 J mice were fed for four additional weeks on a high-fat and high-sucrose (HFHS) diet, supplemented or not with FuFA-F2 (40 mg/day/kg of body weight). Liver triglyceride content and histologic analysis revealed that 4 weeks of FuFA-F2 supplementation fully reversed hepatic steatosis in obese mice. Liver RNA-sequencing analysis highlighted that FuFA-F2 partly reversed the gene expression signature induced by the HFHS diet and favorably changed the expression of many genes known to be involved in the development of hepatic steatosis such as *Pcsk9*, *Stard4*, *Insig1* and *Sulf2*. We also found that FuFA-F2 supplementation increased skeletal muscle mass and protected against cartilage degradation and synovitis induced by obesity. Our findings demonstrated that FuFA-F2 supplementation for 4 weeks in obese mice was enough to reverse the development of MASLD, promote an increase in skeletal muscle mass and protect against cartilage degradation induced by the HFHS diet. This study highlights that nutritional supplementation with FuFA-F2 could be an effective approach to treat obesity-related disorders.

## 1. Introduction

The increase in obesity has become a major global health problem [1] and is associated with numerous metabolic dysfunctions including insulin resistance and the risk of developing metabolic syndrome [2].

Obesity is associated with an increased risk of Metabolic Associated Steatotic Liver Disease (MASLD), formerly known as non-alcoholic fatty liver disease (NAFLD), characterized by inflammation and accumulation of intrahepatic triglyceride content [3,4]. Obesity is also one of the significant risk factors for developing osteoarthritis (OA) [5].

<footnote><sup>a</sup> DMEM, Univ Montpellier, INRAE, Montpellier, France</footnote>

<footnote><sup>b</sup> Institute of Regenerative Medicine and Biotherapies (IRMB), Univ Montpellier, INSERM, Montpellier, France</footnote>

<footnote><sup>c</sup> Qualisud, Univ Montpellier, CIRAD, Montpellier, France</footnote>

<footnote><sup>d</sup> Qualisud, Univ Montpellier, Avignon Université, CIRAD, Institut Agro, IRD, Université de La Réunion, Montpellier, France</footnote>

<footnote><sup>e</sup> Kasetsart agricultural and agro-industrial product improvement institute, Kasetsart University, Bangkok, Thailand</footnote>

<footnote><sup>f</sup> CIRAD, UPR BioWooEB, Montpellier, France</footnote>

<footnote><sup>g</sup> BioWooEB, Univ Montpellier, CIRAD, Montpellier, France</footnote>

<footnote><sup>h</sup> Montpellier University Hospital, Clinical immunology and osteoarticular diseases therapeutic Unit, Lapeyronie, Montpellier, France</footnote>

<footnote>* Corresponding authors.</footnote>

<footnote>E-mail addresses: francois.casas@inrae.fr (F. Casas), christine.coudray@inrae.fr (C. Feillet-Coudray).</footnote>

<footnote n="1">¹ These authors contributed equally to this work.</footnote>
Osteoarthritis is a common chronic degenerative joint disease that is responsible for joint pain, loss of mobility and can lead to disability [6]. Due to its mass and function, skeletal muscle is essential for energy metabolism and is the predominant site of glucose uptake, both insulin-dependent and non-insulin-dependent. [7]. Muscles which are connected to the joints, also play a major role in joint function during movement and in maintaining joint stability [8]. Maintaining its function is therefore central to limit the risk of developing a metabolic syndrome and prevent the risk of OA progression associated with obesity.

Furan fatty acids (FuFAs) are natural compounds synthesized in plants [9] and microorganisms [10,11], and present in many foods [12]. Fish is a particularly important food source of FuFAs, due to its high consumption of algae and marine microorganisms rich in FuFAs [13,14], and FuFAs represent up to 1 % of the fatty acids found in commercial fish oils [15]. FuFAs are also found in butter and dairy products [16,17] and in cereals, vegetables and fruit [13]. Soya contains FuFAs, while levels cannot be quantified in olives, sesame, walnuts, grape seeds and sunflower [18]. Interestingly, these fatty acids, in the form of FuFA-F2 (10, 13-epoxy-11-methyl-octadecan-10,12-dienoic acid), are particularly abundant in the latex of PB235 clone of *Hevea brasiliensis* used to produce the natural rubber [19,20]. In Germany, the average daily intake of FuFAs is estimated at 10–25 mg from various sources (fish, milk, soybean oil, etc...) [17]. In populations with a high consumption of seafood and fermented dairy products, such as the Mediterranean population, it is reasonable to assume that this daily FuFAs intake is higher.

FuFAs have various biological effects that highlight their value as bioactive components for human health [21]. In particular, due to the electron-rich furan ring, FuFAs have antioxidant properties [22–24]. In the presence of reactive oxygen and nitrogen species (RONS), its ring can open and form an unstable and highly reactive dioxoene [25]. *In vitro*, FuFAs have been shown to increase the expression of adiponectin, an adipokine known for its anti-inflammatory and insulin sensitive effects in 3T3-L1 adipocyte cells [26]. *In vivo*, in a rat model of arthritis, FuFAs have a greater anti-inflammatory activity than eicosapentaenoic acid (EPA) [27]. Furthermore, while many studies demonstrate that the polyunsaturated fatty acids (n-3 PUFAs) contained in fish contribute to the prevention of cardiovascular disease (CVD) [28,29], some studies suggest that FuFAs may also be involved in the beneficial effect of fish consumption on CVD [14]. In addition, *in vitro* studies show that FuFAs inhibit platelet aggregation induced by arachidonic acid [30], reduce the oxidation of low-density lipoproteins (LDL) [23] and inhibit the progression of non-enzymatic lipid peroxidation [31]. Thus, in rat, after fish consumption, FuFAs are incorporated into blood phospholipids, coming into intimate contact with endothelial cells where LDL oxidation occurs [32], and may protect against the development of atherosclerosis. Recently, using FuFA-F2, we demonstrated that this fatty acid increased protein content in C2C12 myotubes *in vitro* [33]. In addition, we showed that oral gavage with FuFA-F2 for 3 weeks was sufficient to increase muscle mass and promote a more oxidative muscle metabolism in 3 month-old C57Bl/6 J mice. Furthermore, we showed that 3 months of preventive nutritional supplementation with FuFA-F2 in diet-induced obese (DIO) mice increased muscle mass and reduced the development of metabolic disorders [34]. Moreover, FuFA-F2 supplementation was associated with improved body composition, increased resting energy expenditure, better insulin sensitivity, and the prevention of hepatic steatosis. These studies highlighted that FuFA-F2 partly reproduces the beneficial effects of physical activity. Finally, the observation that FuFA-F6 demonstrated anti-inflammatory activity in a rat model of adjuvant-induced arthritis [27], suggests that FuFA-F2 supplementation could have a favorable effect on the development of obesity-related OA.

Obesity is associated with numerous metabolic dysfunctions and to an increase risk of developing OA. While the liver is a key organ that performs a wide range of biochemical functions necessary for maintaining metabolic homeostasis in the body, skeletal muscle is essential for energy metabolism and is the predominant site of insulin-mediated

glucose uptake in the post prandial state [7]. The aim of this work was therefore to determine whether short FuFA-F2 supplementation (4 weeks) is able to provide beneficial health effects, notably by reversing metabolic disorders and hepatic steatosis, by having a favorable effect on muscle mass, and by limiting cartilage degradation induced by several months of an obesogenic diet in mice. Fig. 1.

## 2. Results

### 2.1. Influence of 4-week FuFA-F2 supplementation on the overall phenotype of already obese mice

Nutritional supplementation with FuFA-F2 over a short 4-week period was carried out on already obese six-month-old mice. In animals fed the HFHS and supplemented with FuFA-F2, we observed no significant differences in final body weight or lean and fat mass compared with mice fed the HFHS diet (Fig. 2A). There was no specific effect of FuFA-F2 on food intake or IL-6 plasma levels, whatever the diet (Fig. 2BC). Furthermore, while plasma cholesterol levels were significantly increased with the HFHS diet compared with controls, no specific effect of FuFA-F2 was observed on plasma triglyceride, cholesterol and free fatty acid levels (Fig. 2D). Regarding the basal metabolism, oxygen consumption and carbon dioxide production using Oxymax-CLAMS system, we found that oxygen consumption, which reflects metabolic expenditure was similar whatever the group (Fig. 2E). In addition, analysis of Respiratory Exchange Ratio (RER) revealed that, as expected, HFHS diet favored the oxidation of lipids, which was reflected by a lower RER value in HFHS-fed mice compared to controls (Fig. 2F). FuFA-F2 supplementation did not further affect RER compared to HFHS alone (Fig. 2F).

### 2.2. Obese mice supplemented with FuFA-F2 for 4 weeks were no longer insulin-resistant

In the fasted state we showed that plasma glucose, insulin levels and HOMA-IR index were increased in HFHS diet-fed mice with no significant effect of FuFA-F2 supplementation (Fig. 3A-C). The fasting blood glucose values we found in HFHS diet-fed mice correspond to prediabetes [35]. Next, we perform insulin sensitivity test. In control animals, insulin caused an important drop in blood glucose levels (Fig. 3D). As expected, in HFHS diet-fed mice we observed a less pronounced fall in blood glucose levels, indicative of insulin resistance (area under the curve –32 % vs Control, p < 0.05) (Fig. 3D). Interestingly, FuFA-F2 supplementation induced a less pronounced reduction in blood glucose than that observed with the HFHS diet alone. Moreover, the difference with the control group was no longer statistically significant, suggesting that FuFA-F2 improved insulin sensitivity (Fig. 3D).

### 2.3. FuFA-F2 supplementation for 4 weeks reversed liver steatosis of obese mice

We found that liver weight was increased by the HFHS diet (Control: 1.10 g, HFHS: 1.32 g, HFHS + FuFAF2: 1.33 g, p < 0.05) (Fig. 4AC). However, plasma ALAT and ASAT activities (Alanine and Aspartate aminotransferase), biochemical markers of hepatic fibrosis, were not modified regardless of the diet (Fig. 4B). Interestingly, whereas we observed a significant increase of liver triglycerides in HFHS group indicative of hepatic steatosis, we found that FuFA-F2 supplementation restored normal levels (Control: 41.1 mg/g, HFHS: 56.4 mg/g, HFHS + FuFAF2: 38.1 mg/g, p < 0.05) (Fig. 4C). Liver cholesterol and free fatty acid levels remained unchanged whatever the groups (Fig. 4C). Measurement of total lipids in the liver showed a similar result (Fig. 4D). Furthermore, in line with triglyceride and total lipids analysis, histologic analysis revealed the presence of lipid droplets in the liver of HFHS group compared to controls where none was observed (Fig. 4E). However, in FuFA-F2-supplemented mice the presence of these lipid droplets
<figure><img src="image_2.png" /><figcaption>Fig. 1. Design of the study. Our experiments were performed on three groups of ten male C57BL/6 J mice (Janvier Labs). Ten mice, aged 6 months on a control diet and weighing about 30 g. Twenty mice aged 6 months and fed a high-fat diet for 20 weeks and weighing about 41 g. These 20 obese mice were randomized, according to their initial weight, into two groups of ten animals. The animals were fed for a further 4 weeks on one of 3 semi-purified diets:1)- control diet, 2)- HFHS diet, 3)- HFHS + FuFA-F2 (0.048 % FuFA-F2/kg diet). Whole-body composition was measured every weeks using an EchoMRI-700 analyzer. Mice oxygen consumption and carbon dioxide production were measured using a Comprehensive Lab Animal Monitoring System the last weeks of the study. Insulin tolerance test were performed the last weeks of the study.</figcaption></figure>

<figure><img src="image_3.png" /><figcaption>Fig. 2. Mice characteristics. (A) Body weight, lean and fat mass. (B) Food intake. (C) Plasma IL-6. (D) Plasma triglycerides, cholesterol and free fatty acids. (E) Oxygen consumption in mice detailed during 24 hours and mean (VO²). (F) Respiratory exchange ratio (RER) in mice detailed during 24 hours and mean. For each mouse, VO² and VCO² were measured 111 times in about 24 h. The respiratory exchange ratio (RER) was calculated as the volume of CO² vs. volume of oxygen (VCO²/VO²) ratio. Mean VO2 and RER values are mean of 8 mice for each group considering all the cycles. For all other parameters, n = 10 animals per group. Results were expressed as means ± SD. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

was nearly undetectable (Fig. 4E). These data indicate that FuFA-F2 supplementation reversed liver steatosis induced by obesogenic diet. Surprisingly, liver gene expression of key players in glucose and lipogenesis homeostasis revealed no difference whatever the groups (Fig. 4F).

## 2.4. FuFA-F2 supplementation for 4 weeks slightly improved mitochondrial respiratory chain activity and promoted antioxidant defense

We next investigated the mitochondrial function and oxidative stress parameters in the liver. While, citrate synthase activity (CS), commonly

used as a marker of the number of mitochondria, remained unchanged regardless of the diet (Fig. 5A), we found a significant decrease of the activities of the complex I and II of the mitochondrial respiratory chain with the HFHS diet (Control: 66 mU/mg, HFHS: 47.9 mU/mg/g, p < 0.05) (Fig. 5B). Interestingly, we found that FuFA-F2 supplementation in obese mice induced a less pronounced reduction of complex I and II activities than that observed with the HFHS group. Moreover, the difference with the control group was no longer significant, suggesting that FuFA-F2 supplementation tended to restore mitochondrial activity (Fig. 5B). In addition, beta-HAD activity increased in obese mice without effect of FuFA-F2 supplementation (Control: 389 nmol/mg, HFHS: 463
<figure><img src="image_4.png" /><figcaption>**Fig. 3.** Glucose homeostasis parameters. (A) Glycemia in fast state. (B) Plasma insulin level in fast state. (C) HOMA-IR. (D) Insulin tolerance test. Results were expressed as means ± S.D., n = 8–10 animals per group. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

nmol/mg, HFHS + FuFA-F2: 484 nmol/mg, p < 0.05) (Fig. 5C).

Regarding oxidative stress markers, we observed a decrease of TBARS levels (a marker of lipid peroxidation) in obese mice (Control: 261 nmol/g, HFHS: 150 nmol/g, HFHS + FuFA-F2: 144 nmol/g, p < 0.05) whereas -SH levels remained unchanged (Fig. 5D). In addition, we observed that in obese mice that GSH content increased (Control: 5626 nmol/g, HFHS: 6307 nmol/g, HFHS + FuFA-F2: 6766 nmol/g, p < 0.05) whereas it decreased for GS=SG (Control: 26,7 nmol/g, HFHS: 14.5 nmol/g, HFHS + FuFA-F2: 14.4 nmol/g, p < 0.05) (Fig. 5E). Moreover, we found that FuFA-F2 supplementation induced a slight increase of SOD and GPx enzymatic activities (for SOD: Control: 19.5 U/mg, HFHS: 19.7 U/mg, HFHS + FuFA-F2: 21.8 U/mg; for GPx: Control: 2.25 U/mg, HFHS: 2.40 U/mg, HFHS + FuFA-F2: 2.57 U/mg, p < 0.05) (Fig. 5F). This set of data indicates that a 4-weeks FuFA-F2 supplementation on obese mice improved mitochondrial respiratory chain activity and promoted antioxidant defense, however slightly.

## 2.5. FuFA-F2 supplementation for 4 weeks partly reverted the gene expression signature induced by HFHS diet

To identify the gene set associated with steatosis and unravel the molecular mechanisms by which FuFA-F2 supplementation reversed hepatic steatosis, we analyzed the liver transcriptomes profiles of mice (Fig. 6A). In the liver of control C57Bl/6 J mice, RNA-sequencing analysis identified the expression of 14446 genes.

Among these genes, 1408 (9.7%) were differentially expressed in the HFHS group compared to control mice (724 upregulated, 684 downregulated) (Fig. 6B). However, we found that only 489 genes (3.3%) were differentially expressed in the HFHS + FuFA-F2 group compared with the control group (236 upregulated, 253 downregulated) (Fig. 6C). This result highlighted that FuFA-F2 supplementation reduced the transcriptional deregulation induced by the HFHS diet. Last, we found that 302 genes (116 upregulated, 186 downregulated) were differentially expressed between the HFHS and HFHS + FuFA-F2 groups (Fig. 6D). When we compared the HFHS + FuFA-F2 group with the control group or the HFHS group for a diet independent specific effect of

FuFA-F2, we identified only 33 differentially expressed genes (18 upregulated, 15 downregulated) (Fig. 6E-F).

Pathway analysis and interrogation of the differentially expressed genes (gene ontology) led us to identify the main biological processes, molecular functions and cellular components affected by FuFA-F2 supplementation compared with HFHS diet alone (Supp Fig. 2). As expected, we found that FuFA-F2 affected the processes involved in cholesterol and steroid biosynthesis, and the pathways involved in lipid transport and metabolism. Among the most modified molecular functions, we identified aromatase activity, oxidoreductase activity and lipid transporter activity. Furthermore, all subcellular compartments appeared to be impacted by FuFA-F2, including peroxisomes, endoplasmic reticulum, mitochondria and cytosol (Supp Fig. 2). In addition, WikiPathways indicated that the main pathways affected were: cholesterol biosynthesis and metabolism, fatty acid biosynthesis and Type II Interferon Signaling Pathways (Supp Fig. 2).

Comparison between the 292 genes differentially expressed between the HFHS and HFHS +FuFA groups enabled us to identify several FuFA-F2 target genes involved in the establishment of hepatic steatosis: 4 upregulated (Pcsk9, Stard4, Insig1, Foxa3)[36–39] and 4 downregulated (Irf7, Stat1, Cd36 and Sulf2)[40–43] (Fig. 6G). We further employed RT-q-PCR to validate the expression profile in our liver samples. Although the pattern of gene expression was similar, only 4 out of the 8 selected genes were found to display the same expression profile by both RT-q-PCR and RNA-seq approach (Fig. 6G-H): Pcsk9, Stard4, Insig1 and Sulf2. Interestingly, among these genes, Pcsk9, Insig1 and Sulf2 are part of the list of 33 genes differentially expressed with FuFA-F2 supplementation.

## 2.6. Obese mice supplemented with FuFA-F2 for 4 weeks had increased muscle mass

We found that 4 weeks of food supplementation with FuFA-F2 increased quadriceps weight (the heaviest skeletal muscle in mice) (218 mg for HFHS group vs 230 for HFHS + FuFA-F2, p < 0.05) (Fig. 7A). In addition, muscle cholesterol, triglycerides and free fatty
<figure><img src="image_5.png" /><figcaption>Fig. 4. Liver phenotype. (A) Liver weight. (B) Plasma ALAT and ASAT activities. (C) Hepatic content of triglycerides, cholesterol and free fatty acids. (D) Total lipids content in percentage. (E) H&E staining of liver, pictures are representative from each group. Scale bar, 100 µm. (F) Relative liver protein expression of key players in glucose and lipogenesis homeostasis. Results were expressed as means ± S.D., n = 10 animals per group. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

acids content remained unchanged regardless of the diet (Fig. 4B). Next, we examined the cross-sectional area of the tibialis muscle although the weight of this small postural muscle did not appear to be affected by FuFA-F2 supplementation (Fig. 7A). However, this muscle is the most practical for analyzing the whole surface and observing changes in muscle fibers. Morphometric analysis of tibialis indicated that neither the HFHS diet nor FuFA-F2 supplementation had any impact on total fiber number (Fig. 7C). However, while we found an increase in mean cross-sectional area (CSA) in the HFHS group compared to controls (1628 µm² for control vs 1724 µm² for HFHS, p < 0.05), we observed that 4-week FuFA-F2 supplementation abolished this effect of the obesogenic diet (Fig. 7D). Furthermore, analysis of the number of fibers in each range of area also showed that the increase in CSA in the HFHS group was due to a significant increase in larger fibers (area greater than 2500 µm²) (684 fibers for control vs 919 fibers for HFHS, p < 0.05) (Fig. 7E-F). Again, we found that FuFA-F2 supplementation restored the number of larger fibers to the level of controls (684 fibers for control vs 709 fibers for HFHS + FuFA-F2) (Fig. 7E-F). In addition, we studied by western-blot skeletal muscle expression of the phosphorylated form of ACC (Acetyl-CoA carboxylase, involved in the biosynthesis of fatty acids), FAS (fatty acid synthase, an enzyme that catalyzes fatty acid synthesis), glucose transporter GLUT4 (a key player in the insulin-induced glucose uptake by muscle), PGC1α (a master regulator of mitochondrial activity), SREBP-1 (Sterol regulatory element-binding

protein 1, regulates genes related to lipid and cholesterol production) and PPARα (Peroxisome proliferator activated receptor alpha, is a transcription factor regulated by free fatty acids, and is a major regulator of lipid metabolism). We observed that in obese mice expression of p-ACC and FAS decreased (for p-ACC, Control: 100, HFHS: 60, p < 0.05; for FAS, Control: 100, HFHS: 52, p < 0.05) whereas it increased for SREBP1 (Control: 100, HFHS: 435, p < 0.05) (Fig. 7G). In addition, we showed that FuFA-F2 supplementation significantly accentuated the drop in p-ACC compared with the HFHS group (HFHS: 60, HFHS + FuFA-F2: 32, p < 0.05) (Fig. 7G).

## 2.7. FuFA-F2 supplementation for 4 weeks had no effect on mitochondrial function and oxidative stress in skeletal muscle of obese mice

We further measured the mitochondrial respiratory chain activities that determine the metabolic properties of skeletal muscle. CS activity increased in both groups of obese mice compared to controls (Control: 443 mU/mg, HFHS: 505 mU/mg, HFHS + FuFA-F2: 545 mU/mg, p < 0.05) (Fig. 8A). In addition, the activities of mitochondrial respiratory chain complexes remained unchanged (Fig. 8B). 3-Hydroxyacyl-CoA dehydrogenase (beta-HAD) is an enzyme involved in the mitochondrial β oxidation. Its activity increased in obese mice without effect of FuFA-F2 supplementation (Control: 92 mU/mg, HFHS: 136 mU/mg, HFHS + FuFA-F2: 137 mU/mg, p < 0.05) (Fig. 8C). Regarding
<figure><img src="image_6.png" /><figcaption>**Fig. 5.** Mitochondrial activity and liver oxidative stress parameters. (A) Citrate synthase activity. (B) Mitochondrial respiratory chain activities. (C) β-HAD activity. (D) TBARS and -SH values. (E) GSH and GS=SG content. (F) Catalase, SOD and GPx activities. Results were expressed as means ± S.D., n = 10 animals per group. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>**Fig. 6.** Identification of downstream targets of FuFA-F2 in liver. (A) The transcriptomes of liver from control, HFHS and HFHS + FuFA-F2 were compared by next-generation RNA sequencing (RNA-Seq). (B) Venn diagram showing the number of co-expressed genes and differentially expressed genes between livers from control and HFHS mice. (C) Venn diagram showing the number of co-expressed genes and differentially expressed genes between livers from control and HFHS + FuFA-F2 mice. (D) Venn diagram showing the number of co-expressed genes and differentially expressed genes between liver from HFHS and HFHS + FuFA-F2 mice. (E) Venn diagram showing the number of co-expressed genes and differentially expressed genes between the genes differentially expressed in both HFHS and HFHS + FuFA-F2 liver compared with control mice. (F) List of the 33 genes up or down regulated in a similar way in liver in HFHS + FuFA-F2 compared to control and HFHS. Expression levels of up-regulated and down-regulated genes are represented in green or red colors, respectively. (G) Custom gene-set analysis of 8 genes differentially expressed between liver from HFHS and HFHS + FuFA-F2. (H) Validation by Q-PCR of the 8 genes found to be differentially expressed between liver from HFHS and HFHS + FuFA-F2 mice. Groups were tested by Student's T-test. The limit of statistical significance was set at p < 0.05.</figcaption></figure>

markers of oxidative stress, TBARS, -SH levels, GSH content and antioxidant enzyme activities (catalase, SOD and GPx) remained unchanged regardless of diet (Fig. 8D-F).

## 2.8. FuFA-F2 supplementation for 4 weeks protected against articular cartilage degradation score and synovitis

Because obesity is also one of the most significant risk factors with

aging to develop osteoarthritis (OA), we sought to determine the impact of FuFA-F2 supplementation on the progression of this still incurable common rheumatic disease [44]. OA is characterized by a loss in whole joint homeostasis leading to cartilage breakdown, synovial inflammation and sub-chondral remodeling [44]. We analyzed the whole joint structure of control, HFHS and HFHS + FuFA-F2 treated mice. Cartilage degradation and synovitis assessed through histological scoring following Safranin-O Fast Green staining (Fig. 9A-B) [44]. Sub-chondral
<figure><img src="image_8.png" /><figcaption>**Fig. 7.** Muscle phenotype. (A) Quadriceps, gastrocnemius and tibialis muscles weight. (B) Quadriceps muscle content of triglycerides, cholesterol and free fatty acids measured on tissue homogenate by enzymatic method. (C) Total fibers number in Tibialis muscles. (D) Mean fibers area in Tibialis muscles (µm²). (E) Fiber size distribution in Tibialis muscles. (F) Number of fibers with CSA greater than 2500 µm². (G) Quantification of relative protein expression in quadriceps muscles extracts. Proteins were quantified with Image Lab™. Touch Software. Results were expressed as means ± S.D., n = 10 animals per group. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

bone structure alterations were evaluated by microCT analysis (Fig. 9C). In line with previous reports, we confirmed that a few months on the HFHS diet are sufficient to induce a significant OA phenotype in animal joints. Specially, we observed a significant increase in both cartilage degradation revealed by OARSI Scoring (Control: 1.6, HFHS: 3.3, p < 0.05) (Fig. 9A) and synovial hypertrophy revealed by synovitis scoring (Control: 0.23, HFHS: 1.26, p < 0.05) (Fig. 9B). Furthermore, sub-chondral bone remodeling was highlighted by an increase in Bone volume (BV) (Control: 0.11 mm³, HFHS: 0.12 mm³, p < 0.05) and thickness (Control: 0.115 mm, HFHS: 0.13, p < 0.05) (Fig. 9C-D) compared to the control group. In addition we observed a reduction of trabecular number (Control: 3.2 mm⁻¹, HFHS: 3 mm⁻¹, p < 0.05) (Fig. 9E). Remarkably, obese mice supplemented with FuFA-F2 were protected against cartilage degradation and synovial inflammation as evidenced by OARSI and synovitis scoring which were not significantly different from that of the control group (Fig. 9B). However, no significant impact of FuFA-F2 supplementation on subchondral bone sclerosis was observed (Fig. 9D-E).

### 3. Discussion

Recently, using a diet-induced obesity (DIO) mouse model, we demonstrated that 3 months of preventive FuFA-F2 supplementation reduced body fat, increased muscle mass, improved insulin sensitivity and prevented hepatic steatosis [34]. Furthermore, FuFA-F2 supplementation in healthy mice has been shown to increase muscle mass [33] and may have a favorable effect on the development of obesity-related osteoarthritis [27]. As obesity is associated with numerous metabolic

dysfunctions and to an increased risk to develop arthritis, the aim of this work was to determine whether a short FuFA-F2 supplementation (4 weeks) could reverse metabolic disorders and hepatic steatosis, improve muscle mass, and limit cartilage degeneration induced by a prolonged obesogenic diet in mice.

After a short 4-week period of nutritional supplementation with FuFA-F2 (40 mg/day/kg of body weight), there was no significant difference in body weight, lean mass, or fat mass between the supplemented mice and those on HFHS diet. However, FuFA-F2 supplementation effectively reversed insulin resistance in the mice, which is consistent with findings from our earlier study, where 12 weeks of preventive FuFA-F2 supplementation improved insulin sensitivity [34].

The systemic low-grade inflammation induced by obesity that drives insulin resistance also contributes to musculoskeletal dysfunction. This chronic inflammation, characterized by elevated reactive oxygen species (ROS), adipokines (such as leptin), and inflammatory cytokines like IL-6, disrupts joint homeostasis by inhibiting chondrogenesis and promoting the production of catabolic enzymes that are markers of the OA phenotype [45]. In addition, recent evidence has shown that musculoskeletal homeostasis is actively controlled by paracrine functions of skeletal muscle. For example, increased production of the myokine irisin by stimulated muscle cells improves OA cartilage structure [46]. We have shown here that FuFA-F2 treatment not only increased muscle mass, but also significantly reduced articular cartilage degradation in the joints of treated animals. Such a remarkable effect on OA cartilage could be due to the combination of a systemic reduction of inflammation and adipokines production in the whole animal and a modulation of
<figure><img src="image_9.png" /><figcaption>**Fig. 8.** Mitochondrial activity and muscle oxidative stress parameters. (A) Citrate synthase activity. (B) Mitochondrial respiratory chain activities. (C) β-HAD activity. (D) TBARS and -SH values. (E) GSH content. (F) Catalase, SOD and GPx activities. Results were expressed as means ± S.D., n = 10 animals per group. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>**Fig. 9.** FUFA-F2 reduces cartilage degradation with no significant effect on bone structure. (A) SafraninO-fast green staining representing cartilage damage to the mice tibia and subchondral bone, scale bar 250 µm. (B) Osteoarthritis Research Society International (OARSI) and synovitis grading of mouse knee cartilage damage degree. (C) Three-dimensional micro-computed tomography (µCT) images of the mouse tibial epiphysis structure. (D) Quantitative analysis of structural parameters of the knee joint subchondral bone: bone volume, bone surface/bone volume (BS/BV) and thickness. (E) Quantitative analysis of structural parameters of knee joint epiphyseal bone: trabecular number and trabecular thickness. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters were significantly different.</figcaption></figure>

myokines secretion by skeletal muscle induced by FuFA-F2 supplementation. Further experiments using FuFA-F2 depleted blood transfer experiments in obese animals could be designed to better decipher how a

simple FuFA-F2 dietary intervention exerts widespread beneficial effects on multiple organs.

We also found that after 6 months on an obesogenic diet (5 months
on the supplier's diet + 1 month in our animal facility), the mice developed hepatic steatosis, as evidenced by liver triglyceride measurements and histological analyses. In contrast, 4-week FuFA-F2 supplementation of already obese mice was sufficient to reverse hepatic steatosis. Due to rising rates of obesity, metabolic syndrome and type 2 diabetes, the incidence of MASLD (formerly known as NAFLD) is set to grow very significantly over the next decade [47]. Therefore, to limit the progression of the disease towards fibrosis and cirrhosis, all people with MASLD must adapt their lifestyle and follow treatments for the underlying metabolic disorders. Necessary lifestyle changes include physical exercise, dietary adjustments, and weight loss. We showed that 4 weeks of FuFAF2 supplementation were sufficient to increase muscle mass in already obese mice, confirming the positive effect of FuFA-F2 supplementation on muscle mass previously observed in healthy mice and in a DIO mouse model when given preventively [33,34]. For the third time, our findings demonstrated that FuFA-F2 partially mimics the positive effects of physical activity by inducing an increase in muscle mass.

Numerous experimental studies have highlighted and characterized the mitochondrial dysfunctions that can occur during MASLD [70,72]. However, results are sometimes contradictory and may depend on the severity of obesity, the presence of associated diabetes or the stage of progression of liver disease. In our study, we observed isolated hepatic steatosis and no fibrosis. Interestingly, we found that FuFA-F2 supplementation in obese mice induced a less pronounced reduction of complex I and II activities of the mitochondrial respiratory chain than that observed with the HFHS group. In addition, regarding oxidative stress markers, our results showed that FuFA-F2 supplementation induced a slight increase of SOD and GPx enzymatic activities. This set of data indicates that a 4-weeks FuFA-F2 supplementation on obese mice slightly improved mitochondrial respiratory chain activity and promoted antioxidant defense. These results are in line with studies showing that n-3 long-chain polyunsaturated fatty acid and extra virgin olive oil supplementation reduces hepatic steatosis in mice fed a high-fat diet, by improving mitochondrial oxidative capacity and reducing oxidative stress [71,73].

Thus, FuFA-F2 could represent an effective and promising natural substance to fight against MASLD through its physical activity mimicking effect and through its direct action on the liver, potentially reducing the need of pharmacological treatment. It is important to note that the Mediterranean diet, rich in oily fish, is naturally rich in FuFAs. These observations are in line with expert recommendations for fighting MAFLD, which encourage following the Mediterranean diet, reducing sedentary behavior and engaging in regular physical activity [75]. Interestingly, Tovar et al. showed by a nontargeted metabolite profiling approach that furan fatty acids present in a multifunctional diet were correlated with a reduction in cardiometabolic risk factors [74]. It would therefore be of interest to study the possible correlation between obesity, circulating furan fatty acid blood concentration and the risk of developing MAFLD in patients.

The RNA-sequencing analysis provided valuable information to understand the effect of FuFA-F2 supplementation on the liver. First, consistent with the observation that FuFA-F2 reversed hepatic steatosis, we demonstrated that it partly corrected the transcriptional deregulation induced by the HFHS diet, as indicated by the decrease in deregulated genes from 1408 to 489. In addition, RNA-Sequencing analysis identified several target genes known to be involved in the development of hepatic steatosis namely *Pcsk9*, *Stard4* and *Insig1* whose expression increase whose expression increased with FuFA-F2 supplementation, and *Sulf2* which expression decreased. The proprotein convertase subtilisin/kexin type 9 (PCSK9), predominantly synthesized in the liver, promotes the internalization and degradation of the hepatic LDL receptor, reducing the liver's ability to remove LDL from the circulation and leading to increased LDL plasma levels [48]. Anti-PCSK9 drugs such as evolocumab and alirocumab are thus indicated in the management of hypercholesterolemia and dyslipidemia [49,50]. However, PCSK9 can also bind to CD36 (a long-chain fatty acid transporter) and initiate its

degradation, thereby reducing fatty acids absorption and triglyceride accumulation in tissues [36]. Conversely, in *Pcsk9* (-/-) mice, the increase in hepatic CD36 correlates with an accumulation of triglycerides and lipid droplets. However, it is important to mention that long-term trials of PCSK9 inhibitors such as evolocumab and alirocumab have not shown any correlation with hepatic steatosis [49,50]. StAR-related lipid transfer protein 4 (STARD4) is a soluble protein involved in cholesterol transport [51]. STARD4 is known to increase the synthesis of bile acids and cholesterol esters in liver hepatocytes [52]. Interestingly, a study by Wei et al. shows that overexpression of STARD4 in the liver of mice fed a high-fat diet protects against hepatic steatosis [38]. Insulin-induced genes 1 (INSIG1) is an anchor protein of the endoplasmic reticulum that regulates sterol regulatory element-binding protein (SREBP) and 3-hydroxy-3-methylglutaryl-coenzyme A reductase (HMG-CoA) function [53,54]. Through these activities, Insig-1 plays a central role in cholesterol metabolism, lipogenesis, and glucose homeostasis in diverse tissues such as adipose tissue and liver [55]. Recently, Zhu et al. showed in *in vivo* and *in vitro* experiments that low abundance of INSIG1 contributed to SREBP-1c processing and hepatic steatosis in dairy cows whereas in hepatocytes treated with palmitate, overexpression of INSIG1 inhibited SREBP activation, up regulation of lipogenic genes and triglycerides synthesis [39]. Interestingly, Hernández-Rodas et al. have shown that supplementation with docosahexaenoic acid and extra-virgin olive oil prevents high-fat diet-induced hepatic steatosis in mice also through down-regulation of SREBP-1c and NF-kB (Hernandez-Rodas, Valenzuela et al., 2017). Sulfatase 2 (SULF2) is a protein of the extracellular matrix that desulfates heparan sulfate proteoglycans (HSPGs) which act as coreceptors for numerous heparinbinding growth factors and cytokines [56]. SULF2-induced desulfation of HSPGs enables the release of growth factors and cytokines from their storage sites to activate downstream signaling pathways [57,58]. In db/db mice, levels of hepatic SULF2 are markedly increased, leading to impaired clearance of very lowdensity lipoproteins [59]. Conversely, inhibition of SULF2 improves diabetic dyslipidemia [59]. In Sulf2(-/-) mice fed a high fat diet, Kim et al. showed a significant decrease of steatohepatitis, and hepatic fibrosis suggesting that targeting of SULF2 signaling may be a potential therapeutic mechanism in MASLD [41]. The significant drop in SULF2 expression in the livers of FuFA-F2-supplemented mice may have contributed to the disappearance of hepatic steatosis observed in our study. Based on this evidence, we hypothesize that FuFA-F2 supplementation reversed hepatic steatosis by increasing the expression of genes such as *Pcsk9*, *Stard4* and *Insig1* while repressing that of *Sulf2*, which in synergy would induce a reduction in triglyceride accumulation in the liver.

In conclusion, we have shown for the first time that a short FuFA-F2 supplementation (4 weeks) in a HFHS diet in already obese mice reversed hepatic steatosis, increased muscle mass and protected against cartilage degradation and synovitis. Our results suggest that FuFA-F2 supplementation or a healthier diet with FuFA-rich foods such as fish and dairy products (Mediterranean diet) may be an effective approach for treating obesity-related disorders such as MASLD, muscle wasting and osteoarthritis. Furthermore, it would be interesting to study the possible association between obesity, circulating levels of furan fatty acids and the risk of developing MAFLD in patients.

# 4. Methods

## 4.1. FuFA-F2 purification

Fresh latex from the PB235 *Hevea brasiliensis* clone was collected from a plantation of Chachoengsao

Rubber Research Center (CRRC), Chachoengsao, Thailand. Lipid extraction was performed by slowly adding the freshly harvested latex (25 l) dropwise into continuously stirred ethyl acetate (120 l, Univar, Illinois, USA) at a latex-to-solvent ratio of ~1:5 (v/v) in a 500 l stainless Steel Tank (B.E. Marubichi, Thailand). The coagulum, primarily
consisting of insoluble polyisoprene was removed and the extract was
left to decant for 2 hours. The upper layer, consisting of lipid-rich ethyl
acetate was collected, evaporated and filtered to remove precipitates
(Whatman No.1, England). Total lipid extract (~125 g, yield ~0.5 % w/
w) was further purified using flash chromatography (Biotage Selekt,
Sweden). For this, a lipid extract (30 g) was loaded onto a 350 g silica
gel (100 Å 60 µM, Biotage® Sfär Silica D - Duo 60 µM). The mobile
phase was made of hexane (100 % from 0 to 15 min) followed by a
gradient of hexane:diethyl ether (90:10, v/v) for 5 min (from 15 to
20 min) with a final 30 min plateau at the same ratio (from 20 to
50 min), all at a flow rate of 50 mL/min and a with collection volume of
50 mL/fraction. The quality of the separation was assessed by spotting
aliquots of each fraction onto a thin layer chromatography plate,
following the method described by Liengprayoon et al. for neutral lipids
[60]. The fractions with the highest trifuranoylglycerol (TGF) purity
were analyzed by GC-FID after derivatization to methyl esters, as pre-
viously described [60]. The FuFA-F2 methyl ester purity within all ob-
tained methyl esters, calculated from the GC-FID, was approximately
98 %.

4.2. *Sex as a biological variable*

Our study exclusively examined male mice. It is unknown whether
the findings are relevant for female mice.

4.3. *Animals and diets ethics statement*

Our experiments were performed on three groups of ten male C57BL/6 J mice (Janvier Labs): ten 6-month-old mice on a control diet and weighing about 30 g; twenty 6-month-old mice that were fed a high-fat diet for 20 weeks and weighing about 41 g, randomized, according to their initial weight, into two groups of ten animals. The animals were fed for a further 4 weeks on one of 3 semi-purified diets: 1)-control diet, 2)-HFHS diet,3)- HFHS + FuFA-F2 (0.048 % FuFA-F2/kg diet). The detailed composition of the diets was previously given [61]. The FuFA-F2 was incorporated into the diet after dissolution in the combined table oils (oil mixture of Carrefour). The dose of FuFA-F2 corresponds for a mouse to an intake of about 40 mg/day/kg of mice. Mice were housed (2 per cage) under controlled conditions of constant temperature (22°C +/-1°C), humidity (40-50 %) and standard dark cycle (19.30-07.30 h). Throughout the study, mice were given free access to tap water and to their respective diets. Mice body weight was monitored weekly and food consumption was determined every 2 days during the week or every 3 days on the week-end. Our institution's guidelines for the care and use of laboratory animals were followed and all experimental procedures were approved by the local ethical committee in Montpellier and French ministry of higher education and research, France (Reference APAFIS#31900-2021060417205673_V2). A detailed scheme of the study design is provided in Fig. 1.

4.4. *Body composition and metabolic analyses*

Mice's whole-body composition (fat and lean masses) was measured
every weeks using an EchoMRI-700 whole-body composition analyzer,
according to the manufacturer's instructions. Oxygen consumption and
carbon dioxide production were measured using a Comprehensive Lab
Animal Monitoring System (CLAMS) (Columbus Instruments). Mice
were housed in individual cages inside a controlled environment. Mice
were acclimatized individually in metabolic cages at 22 °C with *ad
libitum* access to food and water for 24-h, prior to a 24-h period of
automated recordings. Sampled air from individual cages was passed
through sensors to determine O₂ and CO₂ content. The respiratory ex-
change ratio (RER) was calculated as the volume of CO₂ vs. volume of
O₂ (VCO₂/VO₂) ratio [62].

4.5. *Insulin tolerance test (ITT)*

Insulin tolerance tests were performed in the last weeks of the study. ITT was assessed after 2 h fasting by administration of human insulin (0.75 U/kg) by intraperitoneal injection and blood glucose levels were monitored. Blood samples were collected from the tail vein at specified times. Glycemia was measured using a OneTouch Verio glucometer (Lifescan).

4.6. *Sampling and routine biochemical analyses*

At the end of the experiment, the animals were anaesthetized and
blood from 13 h-fasted mice was collected from the inferior vena cava
and distributed into heparinized tubes. Plasma was collected and stored
at -80°C until analysis. After cervical dislocation of mice, liver and
muscles were removed, rinsed with 0.9 % NaCl, weighed, cut into
several parts, when necessary, plunged into liquid nitrogen and then
kept at -80°C until analysis. Plasma levels of glucose, total cholesterol,
triglycerides and free fatty acids as well as enzymatic activity of alanine
aminotransferase (ALT) and aspartate aminotransferase (AST) were
measured at the ANEXPLO analysis platform (CHU RANGUEIL-BP
84225, France). Plasma levels of insulin and IL-6 were quantified
using ELISA kits (Merck Millipore and Crystal Chem respectively).

4.7. *Neutral lipids measurement*

Liver or muscle samples were homogenized in NaCl solution (9 g/L) and Triton X-100 (0.1 %) and free fatty acids, triglycerides and total cholesterol levels were quantified in the tissue homogenate by enzymatic methods (Wako-NEFA-C kit; Cholesterol CHOD-PAP SOBIODA kit and triglycerides LQ SOBIODA kit) [63].

4.8. *Histological study*

After euthanasia by cervical dislocation, liver samples, tibialis anterior muscles and knees joints were collected.

Livers were immediately fixed in 4 % paraformaldehyde at 4°C for
24 h then embedded in paraffin.

Tissue sections were cut at a thickness of 4 µm and stained with
Hematoxylin & Eosin (H&E).

Tibialis anterior muscles were freshly frozen in Tissue-Tek, and then
stored at -80°C. For immunostaining, the sections (10 µm thickness)
were permeabilized 30 min in PBS, 20 % horse serum, 0.1 % triton at
RT, and incubated for 1 h at 37°C with the anti-laminin (1/200, rabbit
polyclonal, Sigma L9393) in a solution of PBS, 1 % BSA. Sections were
washed in PBS 1X for 10 min 3 times and incubated with the secondary
antibody (Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Anti-
body, Alexa Fluor™ 488) for 1 h at 37°C. Sections were washed in PBS
1 × 2 times for 10 min, incubated 30 s with DAPI, and washed once in
PBS for 10 min and mounted with Aqueous Mounting Medium Perma-
Fluor™. For the morphometric analysis, muscle sections were scanned
using a NanoZoomer (Hamamatsu Photonics) with a 20X objective.
Image J free software was used to analyze and quantify the pictures for
each entire area.

For cartilage analysis, mouse knees were fixed in 4 % para-
formaldehyde at 4°C for 48 h, washed in PBS, and then processed for
routine histology. Knees were decalcified in 14 % EDTA/PBS for 3 weeks
and then paraffin-embedded. Tissue Section (5 µm) were rehydrated
through a gradient of ethanol and xylene. Sections were then stained
with Safranin-O/Fast Green to evaluate cartilage degradation. Cartilage
damage was analyzed using an arbitrary score from 0 to 30, based on the
OARSI cartilage OA grading system and modified by van den Berg for
the assessment of murine knee joints (grading scale of 0–6 for the
severity of cartilage destruction and of 0–5 for the extent of damaged
cartilage surface) [64]. To score synovium tissue inflammation, a
semi-quantitative scoring system was employed to evaluate synovitis,
with scores ranging from 0 (no thickening of the lining layer and no immune cell infiltration) to 3 (maximal observed thickening of the synovial lining layer and maximal immune cell infiltration) [65]. For bone parameters analysis, hind leg knees were dissected to remove soft tissues and scanned in a micro-CT scanner SkyScan 1176 (Bruker, Belgium, 0.5 mm aluminum filter, 45 kV, 500 µA, 18 µm resolution, 0.5° rotation angle). Images were reconstructed using CTAn v1.9, Nrecon v1.6 (Bruker, Belgium) and a three-dimensional (3D) model visualization software program (CTVol v2.0). Misalignment compensation, ring artifacts and beamhardening were adjusted to obtain the correct re-construction of each joint. Bone degradation was quantified in subchondral bone and in the epiphysis region of the medial and/or lateral plateau for each tibia (CTAn software, Bruker, Belgium). Reconstructed 3D images of joints were obtained using the Avizo software (Avizo Lite 9.3.0, FEI, France).

## 4.9. Tissue oxidative stress status and mitochondrial enzymatic activities

Long-established oxidative stress parameters (Thiobarbituric Acid Reactive Susbtances (TBARS), thiol groups (-SH), glutathione (GSH), glutathione disulfide (GS-SG), catalase, total superoxide dismutase (SOD), glutathione peroxidase (GPX)), citrate synthase and different mitochondrial respiratory complex activities were measured in liver and muscle as previously reported [66,67].

## 4.10. Protein isolation and western blotting analysis

Frozen liver or muscle samples were homogenized using an Ultra Turax homogenizer. Protein levels were assessed by Western blotting. Total proteins were measured using the Bio-Rad protein assay according to the manufacturer's instructions. Signals were revealed using a ClarityTM Western ECL Substrate Kit, and proteins were visualized by enhanced chemiluminescence using the ChemiDoc Touch Imaging System and quantified with Image Lab. Touch Software (version 5.2.1).

## 4.11. Antibodies

Primary antibodies used include: anti-GLUT2 (1/1000, Abcam ab54460); anti-GLUT4 (1/2000, Abcam ab654); anti-p-ACC (1/1000, Abcam ab92809), anti-FAS (1/1000, Cell signaling 3180); anti-PGC1α (1/1000, Abcam 54481); anti-CPT1 (1/1000, Abcam ab137550); Anti-α-Tubulin (1/6000, Sigma T5160); Anti-beta-Actin (1/400, Santa Cruz sc81178).

## 4.12. RNA-sequencing data analysis

Total RNA was extracted from the liver using Trizol® (n = 4 per group). The concentration of RNA was assessed using a NanoDrop spectrophotometer (Thermo Scientific), and the purity and integrity of the RNA samples were evaluated using the Agilent Bioanalyzer 2100 system (Agilent) using Agilent RNA 6000 Nano Kit. cDNA libraries were constructed using the Illumina Stranded mRNA Prep, Ligation Kit, with 200 ng total RNA. The protocol consisted of the following steps: polyA-selected RNA extraction, RNA fragmentation, random hexamer-primed reverse transcription, addition of an adenine nucleotide (A) to the 3' end of the fragments, pre-index anchors ligation to the double-stranded cDNA fragments, and addition of indexes and amplification of anchor-ligated DNA fragments by PCR. A 2x74nt paired-end sequencing was performed using the Illumina NextSeq550 system. The size distribution and integrity of the sequencing libraries were assessed using the Agilent Bioanalyzer 2100 system with the Agilent High Sensitivity DNA Kit. The construction of sequencing libraries for all samples, as well as the Illumina sequencing, were carried out at the Transcriptomics Core Facility of IRMB, CHU-INSERM-UM Montpellier. MultiQC was utilized to perform a comprehensive statistical analysis of the FastQC files, generating a consolidated report on the quality of the samples. The FastQC

analysis results indicated sufficient data quality and robustness of the sequencing experiment. Reads were mapped to the mouse genome (GRCm38, Mus musculus NCBI GRCm38) using the DRAGEN RNA App (DRAGEN v4.2.7, Illumina). The differential expression was analyzed with DESeq2 software [68,69]. Differentially expressed genes (DEGs) were filtered based on the threshold of significant differential expression, which was p < 0.05, log2Fold Change ≥ 2. The correlation between different samples was analyzed using the Pearson Correlation Coefficient (PCC) and Principal Component Analysis (PCA) (Supplemental Fig. 1).

## 4.13. Gene expression studies

Total RNAs were extracted from liver samples using Trizol®, and cDNAs were generated using the PrimeScript-1st strand cDNA Synthesis Kit (Takara Bio). Real-time PCR was performed using SYBR® Premix Ex Taq-II (Takara Bio) and Applied Biosystems Step-One Plus (Thermo Fisher Scientific). Gene expression was normalized to the expression of the housekeeping gene Rps9. Primer sequences are listed in Supplemental fig. 2.

## 4.14. Statistical analyses

All results are presented as means ± SD, or as percentages. Groups were tested for the effects of diet by a one-way ANOVA test, followed up by a Fisher's Least Significant Difference test. The limit of statistical significance was set at p < 0.05. The means with different letters are significantly different. Statistical analyses were performed using the StatView software.

## CRediT authorship contribution statement

**Liengprayoon Siriluck:** Formal analysis. **Brioche Thomas:** Formal analysis. **Pessemesse Laurence:** Investigation, Formal analysis. **Chaslin Mathys:** Investigation, Formal analysis. **Assou Said:** Investigation, Formal analysis. **Brondello Jean-Marc:** Investigation, Formal analysis. **François Casas:** Writing – original draft, Validation, Supervision, Methodology, Investigation, Funding acquisition, Formal analysis, Conceptualization. **Chaiyut Jatuporn:** Formal analysis. **Feillet-Coudray Christine:** Writing – original draft, Validation, Supervision, Methodology, Investigation, Funding acquisition, Formal analysis, Conceptualization. **Vaysse Laurent:** Formal analysis. **Ferrer Vincent:** Formal analysis. **Macart Mélanie:** Investigation. **Maroun Georges:** Investigation, Formal analysis. **Bertrand-Gaday Christelle:** Investigation, Formal analysis. **Durand Erwann:** Formal analysis. **Pers Yves-Marie:** Formal analysis. **Bonafos Béatrice:** Investigation, Formal analysis. **Coudray Charles:** Formal analysis.

## Animals and ethics statement

Our institution guidelines for the care and use of laboratory animals were observed and all experimental procedures were approved by the local ethical committee in Montpellier, France and the French Research Ministry (Reference APAFIS#31900-2021060417205673_V2).

## Author contributions

M.C., G.M., E.D., B.B., J.C., L.V., S.L., L.P., T.B, M.M., C.B-G., and F.C acquisition of data. E.D., S.A., YMP, C.C., J.M.B., F.C., and C.F-C. analysis and interpretation of data. C.F-C. and F.C. conception and design, wrote the paper. All the authors have read and approved the final version of this manuscript.
Declaration of Generative AI and AI-assisted technologies in the
writing process

During the preparation of this work the authors did not use AI or AI-
assisted technologies.

Funding

This work was supported by the French National Research Institute for Agriculture, Food and Environment (INRAE), by the French Lipid Nutrition Group, by French National Research Institute for Health and Medical Research (INSERM) and by the Key Initiative Food & Health of MUSE (Montpellier University of Excellence). Biotherapy Occitanie Cluster and Tremplin Montpellier Hospital Ph.D program awarded to G. M.

Declaration of Competing Interest

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

Acknowledgements

We thank Pierre Delobel for these helpful suggestions for morpho-
metric analysis of muscle sections. The authors also wish to thank the
Metamus DMeM facility (https://doi.org/10.15454/WYR2-8706),
which belongs to the Montpellier animal facilities network (RAM, Bio-
campus) for technical support and expertise for metabolism phenotyp-
ing, and Metamontp platform supported by the European fund for the
regional development of Occitanie (FEDER). The authors also thank the
"Réseau d'Histologie Expérimentale de Montpellier" (RHEM, Bio-
campus) histology facility for processing our liver samples and Genotoul
Anexplo for biochemical analysis of blood samples. We thank ECell
France and the RHEM platform for bone morphometric and histological
analysis.

Appendix A. Supporting information

Supplementary data associated with this article can be found in the
online version at doi:10.1016/j.biopha.2025.118072.

Data availability

Data will be made available on request.

References

[1] L.M. Jaacks, S. Vandevijvere, A. Pan, C.J. McGowan, C. Wallace, F. Imamura, et al., The obesity transition: stages of the global epidemic, Lancet Diabetes Endo 7 (3) (2019) 231-240.

[2] I. Lingvay, P. Sumithran, R.V. Cohen, C.W. Le Roux, Obesity management as a primary treatment goal for type 2 diabetes: time to reframe the conversation, Lancet 399 (10322) (2022) 394-405.

[3] E. Fabbrini, S. Sullivan, S. Klein, Obesity and nonalcoholic fatty liver disease: biochemical, metabolic, and clinical implications, Hepatology 51 (2) (2010) 679-689.

[4] T.G. Cotter, L. Dong, J. Holmen, R. Gilroy, J. Krong, M. Charlton, Nonalcoholic fatty liver disease: impact on healthcare resource utilization, liver transplantation and mortality in a large, integrated healthcare system, J. Gastroenterol. 55 (7) (2020) 722-730.

[5] J. Martel-Pelletier, A.J. Barr, F.M. Cicuttini, P.G. Conaghan, C. Cooper, M. B. Goldring, et al., Osteoarthritis, Nat. Rev. Dis. Prim. 2 (2016) 16072.

[6] D.J. Hunter, S. Bierma-Zeinstra, Osteoarthritis, Lancet 393 (10182) (2019) 1745-1759.

[7] R.A. DeFronzo, D. Tripathy, Skeletal muscle insulin resistance is the primary defect in type 2 diabetes, Diabetes Care 32 (2009) S157-S163.

[8] E. Shorter, A.J. Sannicandro, B. Poulet, K. Goljanek-Whysall, Skeletal muscle wasting and its relationship with osteoarthritis: a mini-review of mechanisms and current interventions, Curr. Rheuma Rep. 21 (8) (2019) 40.

[9] A. Batna, J. Scheinkonig, G. Spiteller, The occurrence of furan fatty acids in Isochrysis sp. and Phaeodactylum tricornutum, Biochim Biophys. Acta 1166 (2-3) (1993) 171-176.

[10] R.A.S. Lemke, S.M. Olson, K. Morse, S.D. Karlen, A. Higbee, E.T. Beebe, et al., A bacterial biosynthetic pathway for methylated furan fatty acids, J. Biol. Chem. 295 (29) (2020) 9786-9801.

[11] N. Shirasaka, K. Nishi, S. Shimizu, Biosynthesis of furan fatty acids (F-acids) by a marine bacterium, Shewanella putrefaciens, Biochim Biophys. Acta 1346 (3) (1997) 253-260.

[12] L. Xu, A.J. Sinclair, M. Faiza, D. Li, X. Han, H. Yin, et al., Furan fatty acids - beneficial or harmful to health? Prog. Lipid Res 68 (2017) 119-137.

[13] K. Hannemann, V. Puchta, E. Simon, H. Ziegler, G. Ziegler, G. Spiteller, The common occurrence of furan fatty acids in plants, Lipids 24 (4) (1989) 296-298.

[14] G. Spiteller, Furan fatty acids: occurrence, synthesis, and reactions. Are furan fatty acids responsible for the cardioprotective effects of a fish diet? Lipids 40 (8) (2005) 755-771.

[15] C.M. Scrimgeour, Quantitative analysis of furanoid fatty acids in crude and refined cod liver oil, J. Am. Oil Chem. Soc. 54 (5) (1977) 210-211.

[16] W. Vetter, S. Laure, C. Wendlinger, A. Mattes, A.W.T. Smith, D.W. Knight, Determination of furan fatty acids in food samples, 89(8): 1501-1508, J. Am. Oil Chem. Soc. 89 (8) (2012) 1501-1508.

[17] C. Wendlinger, W. Vetter, High concentrations of furan fatty acids in organic butter samples from the German market, J. Agric. Food Chem. 62 (34) (2014) 8740-8744.

[18] H.G. Wahl, H.M. Liebich, A. Hoffmann, Identification of fatty-acid methyl-esters as minor components of fish-oil with multidimensional Gc-Msd - new furan fatty-acids, HrcJ High. Res Chrom 17 (5) (1994) 308-311.

[19] H. Hasma, A. Subramaniam, The occurrence of a furanoid fatty acid inHevea brasiliensis latex, Lipids 13 (12) (1978) 905-907.

[20] S. Liengprayoon, K. Sriroth, E. Dubreucq, L. Vaysse, Glycolipid composition of Hevea brasiliensis latex, Phytochemistry 72 (14-15) (2011) 1902-1913.

[21] K. Alvarado, E. Durand, L. Vaysse, S. Liengprayoon, S. Gaillet, C. Coudray, et al., Potential beneficial effects of furan fatty acids, bioactive food lipids, Cah. Nutr. Diet. 56 (2) (2021) 11725.

[22] M.H. Masuchi Buscato, F. Muller, W. Vetter, J. Weiss, H. Salminen, Furan fatty acids in enriched omega-3 fish oil: oxidation kinetics with and without added monomethyl furan fatty acid as potential natural antioxidant, Food Chem. 327 (2020) 127087.

[23] Y. Okada, M. Kaneko, H. Okajima, Hydroxyl radical scavenging activity of naturally occurring furan fatty acids, Biol. Pharm. Bull. 19 (12) (1996) 1607-1610.

[24] Y. Okada, H. Okamjima, M. Terauchi, H. Konishi, I.M. Liu, H. Watanabe, Inhibitory effects of naturally occurring furan fatty acids on hemolysis of erythrocytes induced by singlet oxygen, Yakugaku Zasshi 110 (9) (1990) 665-672.

[25] A. Batna, G. Spiteller, Oxidation of furan fatty acids by soybean lipoxygenase-1 in the presence of linoleic acid, Chem. Phys. Lipids 70 (2) (1994) 179-185.

[26] J. Lauvai, A.K. Becker, K. Lehnert, M. Schumacher, B. Hieronimus, W. Vetter, et al., The furan fatty acid 9M5 acts as a partial ligand to peroxisome proliferator-activated receptor gamma and enhances adipogenesis in 3T3-L1 preadipocytes, Lipids 54 (5) (2019) 277-288.

[27] T. Wakimoto, H. Kondo, H. Nii, K. Kimura, Y. Egami, Y. Oka, et al., Furan fatty acid as an antiinflammatory component from the green-lipped mussel Perna canaliculus, Proc. Natl. Acad. Sci. USA 108 (42) (2011) 17533-17537.

[28] A. Alhassan, J. Young, M.E.J. Lean, J. Lara, Consumption of fish and vascular risk factors: a systematic review and meta-analysis of intervention studies, Atherosclerosis 266 (2017) 87-94.

[29] D. Mozzaffarian, J.H. Wu, Omega-3 fatty acids and cardiovascular disease: effects on risk factors, molecular pathways, and clinical events, J. Am. Coll. Cardiol. 58 (20) (2011) 2047-2067.

[30] G. Graff, J.L. Gellerman, D.M. Sand, H. Schlenk, Inhibition of blood-platelet aggregation by dioxo-ene compounds, Biochim. Et. Biophys. Acta 799 (2) (1984) 143-150.

[31] C.T. Fuchs, G. Spiteller, 9-(3,4-dimethyl-5-pentyl-furan-2-yl) nonanoic acid and 9-(3,4dimethyl-5-propyl-furan-2-yl) nonanoic acid: new naturally occurring peroxidase inhibitors, Z. Nat. C. 54 (11) (1999) 932-936.

[32] V. Puchta, G. Spiteller, H. Weidinger, F-Acids - a new component of the phospholipids of human-blood, Liebigs Ann. Chem. (1) (1988) 25-28.

[33] F. Pelletier, E. Durand, J. Chaiyut, C. Bronstein, L. Pessemesse, L. Vaysse, et al., Furan fatty acid extracted from Hevea brasiliensis latex increases muscle mass in mice, Biomed. Pharm. 166 (2023) 115330.

[34] L. Dore, E. Durand, B. Bonafos, J. Chaiyut, L. Vaysse, S. Liengprayoon, et al., Preventive nutritional supplementation with furan fatty acid in a DIO mouse model increases muscle mass and reduces metabolic disorders, Biomed. Pharm. 164 (2023) 114945.

[35] R.J. Fajardo, L. Karim, V.I. Calley, M.L. Bouxsein, A review of rodent models of type 2 diabetic skeletal fragility, J. Bone Min. Res 29 (5) (2014) 1025-1040.

[36] A. Demers, S. Samami, B. Lauzier, C. Des Rosiers, E.T. Ngo Sock, H. Ong, et al., PCSK9 induces CD36 degradation and affects long-chain fatty acid uptake and triglyceride metabolism in adipocytes and in mouse liver, Arterioscler. Thromb. Vasc. Biol. 35 (12) (2015) 2517-2525.

[37] R. Gopoju, J. Wang, X. Pan, S. Hu, L. Lin, A. Clark, et al., Hepatic FOXA3 overexpression prevents Western diet-induced obesity and MASH through TGR5, J. Lipid Res 65 (4) (2024) 100527.

[38] X. Wei, F. Yin, M. Wu, Q. Xie, X. Zhao, C. Zhu, et al., G protein-coupled receptor 35 attenuates nonalcoholic steatohepatitis by reprogramming cholesterol homeostasis in hepatocytes, Acta Pharm. Sin. B 13 (3) (2023) 1128-1144.
[39] Y. Zhu, L. Lei, X. Wang, Q. Jiang, J.J. Loor, F. Kong, et al., Low abundance of insulin-induced gene 1 contributes to SREBP-1c processing and hepatic steatosis in dairy cows with severe fatty liver, J. Dairy Sci. 106 (8) (2023) 5626-5635.

[40] M. Grohmann, F. Wiede, G.T. Dodd, E.N. Gurzov, G.J. Ooi, T. Butt, et al., Obesity drives STAT1-dependent NASH and STAT-3-dependent HCC, Cell 175 (5) (2018) 1289-1306, e20.

[41] T.H. Kim, B.A. Banini, F.Z. Asumda, N.A. Campbell, C. Hu, C.D. Moser, et al., Knockout of sulfatase 2 is associated with decreased steatohepatitis and fibrosis in a mouse model of nonalcoholic fatty liver disease, Am. J. Physiol. Gastrointest. Liver Physiol. 319 (3) (2020) G333-G344.

[42] P. Rada, A. Gonzalez-Rodriguez, C. Garcia-Monzon, A.M. Valverde, Understanding lipotoxicity in NAFLD pathogenesis: is CD36 a key driver? Cell Death Dis. 11 (9) (2020) 802.

[43] X.A. Wang, R. Zhang, S. Zhang, S. Deng, D. Jiang, J. Zhong, et al., Interferon regulatory factor 7 deficiency prevents diet-induced obesity and insulin resistance, Am. J. Physiol. Endocrinol. Metab. 305 (4) (2013) E485-E495.

[44] Y. Liu, Z. Zhang, T. Li, H. Xu, H. Zhang, Senescence in osteoarthritis: from mechanism to potential treatment, Arthritis Res Ther. 24 (1) (2022) 174.

[45] C. Zhang, Y. Lin, C.H. Yan, W. Zhang, Adipokine signaling pathways in osteoarthritis, Front Bioeng. Biotechnol. 10 (2022) 865370.

[46] K. Ning, Z. Wang, X.A. Zhang, Exercise-induced modulation of myokine irisin in bone and cartilage tissue-positive effects on osteoarthritis: a narrative review, Front Aging Neurosci. 14 (2022) 934406.

[47] M.L. Teng, C.H. Ng, D.Q. Huang, K.E. Chan, D.J. Tan, W.H. Lim, et al., Global incidence and prevalence of nonalcoholic fatty liver disease, Clin. Mol. Hepatol. 29 (2023) S32-S42 (Suppl).

[48] K.N. Maxwell, E.A. Fisher, J.L. Breslow, Overexpression of PCSK9 accelerates the degradation of the LDLR in a post-endoplasmic reticulum compartment, Proc. Natl. Acad. Sci. USA 102 (6) (2005) 2069-2074.

[49] M. Farnier, G.K. Hovagh, G. Langslet, R. Dufour, M.T. Baccara-Dinet, C. Din-Bell, et al., Longterm safety and efficacy of alirocumab in patients with heterozygous familial hypercholesterolemia: an open-label extension of the ODYSSEY program, Atherosclerosis 278 (2018) 307-314.

[50] M.L. O'Donoghue, R.P. Giugliano, S.D. Wiviott, D. Atar, A. Keech, J.F. Kuder, et al., Long-term evolocumab in patients with established atherosclerotic cardiovascular disease, Circulation 146 (15) (2022) 1109-1119.

[51] B. Mesmin, N.H. Pipalia, F.W. Lund, T.F. Ramlall, A. Sokolov, D. Eliezer, et al., STARD4 abundance regulates sterol transport and sensing, Mol. Biol. Cell 22 (21) (2011) 4004-4015.

[52] D. Rodriguez-Agudo, S. Ren, E. Wong, D. Marques, K. Redford, G. Gil, et al., Intracellular cholesterol transporter StarD4 binds free cholesterol and increases cholesteryl ester formation, J. Lipid Res 49 (7) (2008) 1409-1419.

[53] N. Sever, T. Yang, M.S. Brown, J.L. Goldstein, R.A. DeBose-Boyd, Accelerated degradation of HMG CoA reductase mediated by binding of insig-1 to its sterol-sensing domain, Mol. Cell 11 (1) (2003) 25-33.

[54] T. Yang, P.J. Espenshade, M.E. Wright, D. Yabe, Y. Gong, R. Aebersold, et al., Crucial step in cholesterol homeostasis: sterols promote binding of SCAP to INSIG-1, a membrane protein that facilitates retention of SREBPs in ER, Cell 110 (4) (2002) 489-500.

[55] X.Y. Dong, S.Q. Tang, Insulin-induced gene: a new regulator in lipid metabolism, Peptides 31 (11) (2010) 2145-2150.

[56] I. Matsuo, C. Kimura-Yoshida, Extracellular distribution of diffusible growth factors controlled by heparan sulfate proteoglycans during mammalian embryogenesis, Philos. Trans. R. Soc. Lond. B Biol. Sci. 369 (1657) (2014).

[57] J.P. Lai, D.S. Sandhu, C. Yu, T. Han, C.D. Moser, K.K. Jackson, et al., Sulfatase 2 up-regulates glypican 3, promotes fibroblast growth factor signaling, and decreases survival in hepatocellular carcinoma, Hepatology 47 (4) (2008) 1211-1222.

[58] J.P. Lai, J.R. Thompson, D.S. Sandhu, L.R. Roberts, Heparin-degrading sulfatases in hepatocellular carcinoma: roles in pathogenesis and therapy targets, Future Oncol. 4 (6) (2008) 803-814.

[59] K. Chen, M.L. Liu, L. Schaffer, M. Li, G. Boden, X. Wu, et al., Type 2 diabetes in mice induces hepatic overexpression of sulfatase 2, a novel factor that suppresses uptake of remnant lipoproteins, Hepatology 52 (6) (2010) 1957-1967.

[60] S. Liengprayoon, J. Chaiyut, K. Sriroth, F. Bonfils, J. Sainte-Beuve, E. Dubreucq, et al., Lipid compositions of latex and sheet rubber from Hevea brasiliensis depend on clonal origin, Eur. J. Lipid Sci. Tech. 115 (9) (2013) 1021-1031.

[61] L. Dore, E. Durand, B. Bonafos, J. Chaiyut, L. Vaysse, S. Liengprayoon, et al., Preventive nutritional supplementation with furan fatty acid in a DIO mouse model increases muscle mass and reduces metabolic disorders, Biomed. Pharmacother. 164 (2023) 114945.

[62] P.M. Marvyn, R.M. Bradley, E.B. Mardian, K.A. Marks, R.E. Duncan, Data on oxygen consumption rate, respiratory exchange ratio, and movement in C57BL/6J female mice on the third day of consuming a high-fat diet, Data Brief. 7 (2016) 472-475.

[63] C. Coudray, G. Fouret, K. Lambert, C. Ferreri, J. Rieusset, A. Blachnio-Zabielska, et al., A mitochondrial-targeted ubiquinone modulates muscle lipid profile and improves mitochondrial respiration in obesogenic diet-fed rats, Brit J. Nutr. 115 (7) (2020) 1155-1166.

[64] W. Su, G. Liu, B. Mohajer, J. Wang, A. Shen, W. Zhang, et al., Senescent preosteoclast secretome promotes metabolic syndrome associated osteoarthritis through cyclooxygenase 2, Elife 11 (2022).

[65] O. Malaise, Y. Tachikart, M. Constantinides, M. Mumme, R. Ferreira-Lopez, S. Noack, et al., Mesenchymal stem cell senescence alleviates their intrinsic and seno-suppressive paracrine properties contributing to osteoarthritis development, Aging-Us 11 (20) (2019) 9128-9146.

[66] M. Benlebna, L. Balas, B. Bonafos, L. Pessemesse, G. Fouret, C. Vigor, et al., Long-term intake of 9-PAHPA or 9-OAHPA modulates favorably the basal metabolism and exerts an insulin sensitizing effect in obesogenic diet-fed mice, Eur. J. Nutr. (2020).

[67] M. Benlebna, L. Balas, B. Bonafos, L. Pessemesse, C. Vigor, J. Grober, et al., Long-term high intake of 9-PAHPA or 9-OAHPA increases basal metabolism and insulin sensitivity but disrupts liver homeostasis in healthy mice, J. Nutr. Biochem 79 (2020) 108361.

[68] M.I. Love, W. Huber, S. Anders, Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2, Genome Biol. 15 (12) (2014) 550.

[69] H. Varet, L. Brillet-Gueguen, J.Y. Coppee, M.A. Dillies, SARTools: a DESeq2- and EdgeR-Based R pipeline for comprehensive differential analysis of RNA-Seq data, PLoS One 11 (6) (2016) e0157022.

[70] K. Begrique, J. Massart, M.A. Robin, F. Bonnet, B. Fromenty, Mitochondrial adaptations and dysfunctions in nonalcoholic fatty liver disease, Hepatology 58 (4) (2013) 1497-1507.

[71] F. Echeverría, R. Valenzuela, A. Bustamante, D. Alvarez, M. Ortiz, A. Espinosa, P. Illesca, D. Gonzalez-Manan, L.A. Videla, High-fat diet induces mouse liver steatosis with a concomitant decline in energy metabolism: attenuation by eicosapentaenoic acid (EPA) or hydroxytyrosol (HT) supplementation and the additive effects upon EPA and HT co-administration, Food Funct. 10 (9) (2019) 6170-6183.

[72] B. Fromenty, M. Roden, Mitochondrial alterations in fatty liver diseases, J. Hepatol. 78 (2) (2023) 415-429.

[73] M.C. Hernandez-Rodas, R. Valenzuela, F. Echeverría, M.A. Rincon-Cervera, A. Espinosa, P. Illesca, P. Munoz, A. Corbari, N. Romero, D. Gonzalez-Manan, L. A. Videla, Supplementation with docosahexaenoic acid and extra virgin olive oil prevents liver steatosis induced by a high-fat diet in mice through PPAR-alpha and Nrf2 upregulation with concomitant SREBP-1c and NF-kB downregulation, Mol. Nutr. Food Res 61 (12) (2017).

[74] J. Tovar, V.D. de Mello, A. Nilsson, M. Johansson, J. Paananen, M. Lehtonen, K. Hanhineva, I. Bjorck, Reduction in cardiometabolic risk factors by a multifunctional diet is mediated via several branches of metabolism as evidenced by nontargeted metabolite profiling approach, Mol. Nutr. Food Res 61 (2) (2017).

[75] X.F. Zeng, K.A. Varady, X.D. Wang, G. Targher, C.D. Byrne, R. Tayyem, G. Latella, I. Bergheim, R. Valenzuela, J. George, C. Newberry, J.S. Zheng, E.S. George, C. W. Spearman, M.D. Kontogianni, D. Ristic-Medic, W.A.F. Peres, G.Y. Depboylu, W. Yang, X. Chen, F. Rosqvist, C.S. Mantzoros, L. Valenti, H. Yki-Jarvinen, A. Mosca, S. Sookoian, A. Misra, Y. Yilmaz, W. Kim, Y. Fouad, G. Sebastiani, V. W. Wong, F. Aberg, Y.J. Wong, P. Zhang, F.J. Bermudez-Silva, Y. Ni, M. Lupso-Platon, W.K. Chan, N. Mendez-Sanchez, R.J. de Knegt, S. Alam, S. Treeprasertsuk, L. Wang, M. Du, T. Zhang, M.L. Yu, H. Zhang, X. Qi, X. Liu, K. Pinyopornpanish, Y. C. Fan, K. Niu, J.C. Jimenez-Chillaron, M.H. Zheng, The role of dietary modification in the prevention and management of metabolic dysfunction-associated fatty liver disease: an international multidisciplinary expert consensus, Metabolism 161 (2024) 156028.