Metabolic syndrome (MS), characterized by three or more of five medical components (including obesity, hyperglycemia, dyslipidemia, hypertension, and insulin resistance), can enhance the risk of type 2 diabetes (T2DM) and cardiovascular disease (CVD) [1
]. Compared with people without MS, people suffering from MS are twice as likely to die and three times as likely to have a heart attack or stroke. The prevalence of MS is growing worldwide, especially in developing countries, which is mainly due to the changes in lifestyles and dietary patterns. It is estimated that about one in four people worldwide suffer from MS [2
]. The current national prevalence of MS among Chinese adults is 24.2 %, which is a sharp growth compared with the value of 9.8% calculated 10 years ago under the same standard of diagnosis [3
]. Thus, prevalence of MS has become a severe threat to modern society, and its preventative strategies are significant.
A large amount of data indicates that gut microbiota is essential for maintaining the metabolic homeostasis of the host [5
]. A wide variety of commensal microbes colonizes gut lumen. For example, the population quantity of microbes in the gut microbiota is approximately 10 times that of somatic cells in human body. These microbes participate in most metabolic activities in vivo. Recently, some beneficial species or genera of gut microbiota have been demonstrated to be negatively associated with the development of MS, such as Akkermansia
], and Lactobacillus
]. In contrast, over-proliferation of some pro-inflammatory or pathogenic gut microbiota, such as Erysipelotrichaceae
, and Streptococcaceae
, are associated with the development of obesity, systemic inflammation, and metabolic disorders in both humans and rodents [8
]. Thus, regulating diet-induced gut microbiota disruption has been presented as a potential intervention target for the prevention of MS and related diseases [11
While many potential risk factors for MS have been identified, dietary patterns definitely play an important role. The intake and profile of dietary fat plays an accepted role in CVD risk and the development of cardio-metabolic diseases such as those syndromes included in the diagnostic criteria of MS [12
]. It has been demonstrated that dietary fat intake is a dominant factor influencing the pattern of gut microbiota [6
], and the Firmicutes
ratio (F/B ratio) [15
]. The F/B ratio is positively correlated with lean phenotype/weight loss, which has been supported by several studies [16
]. Compared with diets enriched with saturated fatty acids, diets with high levels of unsaturated fatty acids have been associated with lower body weight gain, and lipid accumulation in liver. These effects have been confirmed to be related to the diet-induced changes of gut microbiota [17
]. For the past few years, dietary fructose intake has also come under scrutiny. Several epidemiologic surveys suggest an underlying link between consumption of fructose containing sugars such as high fructose corn syrup, and sucrose and risk factors for CVD, and diagnostic standards of MS [18
]. A meta-analysis including 15 studies also showed that fructose consumption from processed foods is one of the causes of some chronic disorders such as MS among healthy adults [21
]. The effect of fructose on development of MS is driving hepatic fat, which can induce insulin resistance [22
]. The specific effects of fructose on liver are particularly related to a vicious circle that starts with liver steatosis driving insulin resistance. Fructose derived advance glycation end-products may promotes inflammation by engaging receptor for advanced glycation end products (RAGE). Thus, rats or mice fed with a high-fructose-high-fat diet (HFHFD) are used as an animal model of MS.
Virgin olive oil is the main source of dietary fat at the core of Mediterranean diet. There is a widespread recognition on association between the regular consumption of virgin olive oil and a lower risk of MS [23
]. These beneficial biological activities have been attributed not only to the high monounsaturated fatty acid (MUFA) content but also to the minor bioactive phytochemicals [24
]. Recent research indicates the possibility that virgin olive oil may attenuate MS associated with modulation of intestinal microbiota [25
]. Similarly, HOPO is also rich in MUFA (providing up to 80% of the fatty acid composition, similar even higher level to olive oil) and minor bioactive phytochemicals, such as polyphenol, phytosterols, and vitamin E, etc. However, MS prevention and gut microbiota modulating of HOPO has never been studied.
Hence, in this study, we conducted a comparison of the effects of HOPO, and EVOO supplement on MS in HFHFD-fed rats. Then, to illustrate the possible mechanisms, the profile of the gut microbiota was analyzed by utilizing a 16S rRNA sequencing technique, and the biochemical indexes were also determined. We demonstrated that both HOPO and EVOO can attenuate the HFHFD-induced MS. Moreover, this study presents a new perception of gut microbiota modulation in the prevention of MS by dietary fats rich in MUFA.
2. Materials and Methods
HOPO provided by Luhua Group (Laiyang, China). EVOO purchased from Mueloliva Co. Ltd. (Córdoba, Spain). Fatty acid profiles of HOPO, and EVOO are shown in Table S1
. Fructose purchased from SIWANG SUGAR Co. Ltd. (Binzhou, China).
2.2. Animals and Treatment
The 48 6-week-old male SD rats were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China). After fed adaptively for 1 week, rats were randomly divided into 4 groups (n = 12) to receive respectively the following diets ad libitum for 12 weeks: (A) NC (normal control, normal chow diet + ordinary drinking water); (B) M (model, high-fat diet + drinking water contains 10% fructose); (C) HOPO (high oleic acid peanut oil diet, high-fat diet contains 10% HOPO + drinking water contains 10% fructose); and (D) EVOO (extra virgin olive oil diet, high-fat diet contains 10% EVOO + drinking water contains 10% fructose). Compositions of the diets are shown in Table S2
. All the diets were purchased from Trophic Animal Feed High-Tech Co. Ltd. (Nantong, China). The rats were kept in a well-ventilated room maintained at 23 ± 2 °C with 12-h light-dark cycles. Their body weight and food and water intake were recorded weekly. Fasting blood-glucose, oral glucose tolerance test (OGTT), and insulin tolerance test (ITT) were measured at 0, 4, 8 and 12 weeks. Protocols for animal studies were approved by the Institutional Animal Care and Use Committee of Beijing Vital River Laboratory Animal Technology Co., Ltd. (VR IACUC, Beijing, China, No. P2018036).
2.3. Insulin Resistance Assessment
Oral glucose tolerance test (OGTT): the rats were fasted for 12-h and orally infused with glucose (2 g/kg). The blood was collected from tail vein and its glucose level was measured with glucometer (Johnson and Johnson Investment, Co. Ltd., Shanghai, China) before (0 min) and after gavage (15, 30, 60, 90 and 120 min). The area under curve (AUC) was calculated to represent the glucose tolerance. AUC was calculated using the formula AUC = 0.25 × (G0 + G15)/2 + 0.25 × (G15 + G30)/2 + 0.5 × (G30 + G60)/2 + 0.5 × (G60 + G90)/2 + 0.5 × (G90 + G120)/2, where G0, G15, G30, G60, G90, and G120 were blood glucose levels at different time points.
Insulin tolerance test (ITT): the rats were fasted for 12-h and received intraperitoneal injection of insulin solution (0.75 U/kg). Blood glucose test and AUC value calculation were carried out as for OGTT. HOMA-IR was calculated using the formula HOMA-IR = (FPG × FINS)/22.5, where FPG and FINS were fasting blood-glucose level and fasting insulin level.
2.4. Serum Biochemical Analysis
At the end of experiment, the animals were sacrificed, and blood samples were collected. Serum samples were prepared by centrifugation (4 °C, 2000× g for 15 min). SerumTG, TC, low density lipoprotein (LDL), high density lipoprotein (HDL), insulin, free fatty acid (FFA), and TNF-a levels were analyzed by using kits from Nanjing Jiancheng Bioengineering Institute (Nanjing, China).
2.5. Histopathological Examination and TG Level in Liver Tissue
Freshly isolated liver tissues were embedded in optimal cutting temperature (OCT) compound and stored at −80 °C ultra-cold storage freezer after flash frozen. The liver tissues were cut into 8 μm sections on a cryostat, and fixed with 4% paraformaldehyde for 10 min. The sections were stained with oil red O for 12 min and re-dyed with hematoxylin for 4 min. The sections were observed under a light microscope (Primo star, Carl Zeiss Microscopy GmbH, Jena, Germany). TG level in liver tissue were analyzed by using kits from Nanjing Jiancheng Bioengineering Institute (Nanjing, China).
2.6. Gut Microbiota Analysis
After 12 weeks of experimental treatment, fresh fecal samples were collected in sterilized Eppendorf tubes and stored at −80 °C ultra-cold storage freezer after flash frozen. Ten rats were selected randomly from each group for gut microbiota analysis. According to the instructions, QIAamp DNA stool Mini Kit from Qiagen (Hilden, Germany) was used to extract bacterial genomic DNA from frozen fecal samples temporary stored at −80 °C for 24 h. The 16S rRNA gene comprising V3 and V4 regions was magnified by PCR using composite specific bacterial primers (Table S3
). Thermal cycling was as following: 95 °C for 5 min (1 cycle), 95 °C for 30 s/50 °C for 30 s/72 °C for 40 s (25 cycles), and a final extension at 72 °C for 7 min. High-throughput pyrosequencing of the PCR products was performed on an Illumina MiSeq platform by Biomarker Technologies Co, Ltd. (Beijing, China).
The raw-paired end reads from the original DNA fragments were merged using FLASH and assigned to each sample according to the unique barcodes. High-quality reads for bioinformatics analysis were performed, and all of the effective reads from each sample were clustered into operational taxonomic units (OTUs) based on a 97% sequence similarity according to UCLUST. After the OTU data are normalized (logarithm), the top 80 species are selected and drawn based on the R heatmap. For alpha diversity analysis, we rarified the OTUs to several metrics, including curves of OTU rank, rarefaction and Shannon, and calculated indexes of Shannon, Chao1, Simpson, and ACE. For β-diversity analysis, heatmap of RDA-identified key OTUs, principal component analysis (PCA) and nonmetric multidimensional scaling (NMDS) were performed using QIIME. The LDA effect size (LEfSe) analysis was performed for the quantitative analysis of biomarkers among each group. Briefly, LEfSe analysis, LDA threshold of >4, used the nonparametric factorial Kruskal-Wallis (KW) sum-rank test and then used the (unpaired) Wilcoxon rank-sum test to identify the most differently abundant taxa. Metastats analysis obtained p values by T-test of relative abundance data. The species causing the difference in the composition of the two groups were screened out by p values (p < 0.05).
2.7. Statistical Analysis
Results were expressed as means ± SD. The statistical analysis was performed using SPSS, version 20 (IBM, Armonk, NY, USA). Differences between groups were statistically analyzed using ANOVA followed by Duncan’s test and considered statistically with a level of p < 0.05.
MS has been regarded as one of the urgent worldwide public health problems of this century, which enhances the risk of developing CVD and T2DM [1
]. Dietary pattern is one of the most important factors in the pathogenesis of MS. The type and quantity of dietary fat have a significant impact on the components of MS [27
]. In the present study, we demonstrated that both HOPO and EVOO can attenuate the HFHFD-induced MS, which was associated with reshaping the profile of gut microbiota. The results showed that both HOPO and EVOO significantly reduced body weight gain, and significantly improved insulin sensitivity and HDL/LDL. Moreover, the 16S rRNA gene sequence data indicated that HOPO and EVOO prevented HFHFD-induced gut disorder, and enriched relative abundance of probiotics, such as Bifidobacterium
As one of the core ingredients of Mediterranean diet, olive oil is recognized as a healthy dietary oil, and EVOO has the highest quality among olive oils. The health benefits of EVOO have been attributed not only to the high MUFA content but also to the minor bioactive phytochemicals content. Oleic acid not only has basic nutritional functions, but also has many health functions, such as anti-inflammation, anti-hypertension, cardiovascular disease, and diabetes prevention [28
]. Additionally, as a MUFA, oleic acid has strong oxidation stability. Thus, high-oleic acid peanuts have strong oxidation stability, long shelf-life, better nutritional and health functions that are favored by consumers. Compared to olive oil, HOPO has a similar and even higher MUFA level, and is also rich in minor bioactive phytochemicals, such as phytosterols and polyphenol. In fact, as one of the most important oil crops in the world, peanuts are grown on all six continents except Antarctica, and is therefore more widely grown than olives [29
]. In the present study, we found that both HOPO and EVOO could improve serum lipids profile in HFHFD-induced MS rats, and protective effects of HOPO were better than EVOO. Due to the high production levels, moderate prices and attractive flavor, HOPO has great potential to be used as a health-promotive oil, especially in the countries and areas unsuitable for olive cultivation.
Accumulating experimental evidence has suggested that the human gut microbiota plays a fundamental role in the metabolic capacity for processing nutrients and host health [30
]. Recent research indicates the possibility that in attenuating MS effects of virgin olive oil the modulation of intestinal microbiota could be involved in [25
]. EVOO diet-fed mice had significant lower body weight than butter groups (p
< 0.01), although EVOO group had a higher food intake level. Serum insulin level in EVOO group was significantly higher than the butter group, while serum glucose level without significant difference between two groups [25
]. These results were in line with this study. As expected, supplementation of MUFA-rich oils-HOPO and EVOO could significantly suppress the HFHFD-induced body weight gain, insulin resistance, and gut disorder by modulating the gut microbiota. Bifidobacterium
is a gram-positive, anaerobe bacterium belonging to the family Bifidobacteriaceae
, which is one of the most important probiotics in human gut. Relative abundance reduction of Bifidobacterium
has been demonstrated to be associated with obesity and its related diseases [33
]. The possible protective mechanism is maintaining normal intestinal permeability and restraining inflammation caused by lipopolysaccharide. The relative abundance of Bifidobacterium
in HOPO and EVOO group was significantly higher than M group in this study. The family Lachnospiraceae
is supposed to facilitate lipopolysaccharide transfer from the intestinal tract to the blood [34
]. Previous studies have suggested that Lachnospiraceae
family is significantly increased in both obese nonalcoholic fatty liver disease and T2DM in humans [35
is a genus of the family Lachnospiraceae
, which was thought to take part in the development of glucose metabolism disturbances [37
]. In this study, relative abundance of Lachnospiraceae
in HOPO group and EVOO group was lower than M group, but only HOPO group with significant difference.
However, there are some gut microbiota results which are unlike some other studies. Many studies have shown that hosts with obesity, T2DM and high fat diet has a higher F/B ratio [15
]. The imbalance of F/B ratio might be a contributing factor to obesity and its relative metabolic disease. In this study, F/B ratio of M group was increased compared to NC group, but F/B ratio of HOPO group and EVOO group was higher than M group. A few studies indicated that F/B ratio was not necessarily related with obesity linked diseases [40
]. Duncan et al. found that there is no relationship between BMI or absolute weight loss and the relative proportions of Bacteroidetes and Firmicutes of colonic bacteria in obese or non-obese subjects [41
]. This correlation is controversial and needs further research so far. Moreover, Akkermansia
be regarded as one of major probiotics which can increase the thickness of intestinal mucin and improve the barrier function of intestinal mucosa, so as to inhibit the obesity caused by high-fat diet [42
]. However, relative abundance of Akkermansia
in NC group was significantly lower than M group, and there was no significant difference between HOPO group vs. M group and EVOO group vs, M group. The variation on Akkermansia
abundance of the same dietary intervention may depend on its baseline abundance level. Subjects in the caloric restricted diet group with a high baseline level of Akkermansia
had a decrease in abundance of Akkermansia,
while there was an increase in subjects with a low baseline level [7
]. The abundance changes of some specific species should not be simply judged as a possible mechanism by the past similar studies. Therefore, we are enlightened that the interactions within gut microbiota species and between gut microbiota and external factors are still insufficiently studied. Above all, we suppose that Bifidobacterium
is the key probiotic of rats fed with HOPO and EVOO. Lower Lachnospiraceae
relative abundance were also related to modulation of the gut microbiota.