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. 2024 Jan 3;17:25–35. doi: 10.1016/j.aninu.2023.12.006

Trimethylamine oxide supplementation differentially regulates fat deposition in liver, longissimus dorsi muscle and adipose tissue of growing-finishing pigs

Andong Zha a,c, Wanquan Li b, Jing Wang b, Ping Bai d, Ming Qi a,c, Peng Liao a, Bie Tan b,∗, Yulong Yin a,b,c,d
PMCID: PMC10920132  PMID: 38464952

Abstract

Trimethylamine oxide (TMAO) is a microbiota-derived metabolite, and numerous studies have shown that it could regulate fat metabolism in humans and mice. However, few studies have focused on the effects of TMAO on fat deposition in growing-finishing pigs. This study aimed to investigate the effect of TMAO on fat deposition and intestinal microbiota in growing-finishing pigs. Sixteen growing pigs were randomly divided into 2 groups and fed with a basal diet with 0 or 1 g/kg TMAO for 149 d. The intestinal microbial profiles, fat deposition indexes, and fatty acid profiles were measured. These results showed that TMAO supplementation had a tendency to decrease lean body mass (P < 0.1) and significantly increased backfat thickness (P < 0.05), but it did not affect growth performance. TMAO significantly increased total protein (TP) concentration, and reduced alkaline phosphatase (ALP) concentration in serum (P < 0.05). TMAO increased the α diversity of the ileal microbiota community (P < 0.05), and it did not affect the colonic microbial community. TMAO supplementation significantly increased acetate content in the ileum, and Proteobacteria and Escherichia–Shigella were significantly enriched in the TMAO group (P < 0.05). In addition, TMAO decreased fat content, as well as the ratio of linoleic acid, n-6 polyunsaturated fatty acids (PUFA), and PUFA in the liver (P < 0.05). On the contrary, TMAO increased intramuscular fat content of the longissimus dorsi muscle, whereas the C18:2n6c ratio was increased, and the n-6 PUFA:PUFA ratio was decreased (P < 0.05). In vitro, 1 mM TMAO treatment significantly upregulated the expression of FASN and SREBP1 in C2C12 cells (P < 0.05). Nevertheless, TMAO also increased adipocyte area and decreased the CPT-1B expression in subcutaneous fat (P < 0.05). Taken together, TMAO supplementation regulated ileal microbial composition and acetate production, and regulated fat distribution and fatty acid composition in growing-finishing pigs. These results provide new insights for understanding the role of TMAO in humans and animals.

Keywords: Growing-finishing pig, Trimethylamine oxide, Fat deposition, Fatty acid composition, Intestinal microbiota

1. Introduction

Fat deposition is an effective way to store energy in humans and animals, and is closely associated with long-term metabolic health. In growing-finishing pigs, fat deposition is also associated with meat quality. Recent studies have shown that fat deposition is not only associated with genetics, diet and lifestyle, but also significantly associated with gut microbes. Bäckhed et al. (2004) showed that fecal microbiota transplantation regulated fat deposition in germ-free mice. In addition, an increasing number of studies also showed that single species also regulate fat deposition. Akkermansia muciniphila improves obesity by promoting energy consumption and downregulates perilipin 2 protein expression in mice (Depommier et al., 2020). Parabacteroides distasonis increased the circulating succinate and lithocholic acid levels to decrease weight gain and fat storage in mice (Wang et al., 2019a). The colonization of Prevotella copri activated chronic inflammatory responses via the TLR4/mTOR pathway and significantly upregulated the expression of lipolysis genes to promote fat deposition in mice (Chen et al., 2021). Therefore, it is an effective way to regulate fat deposition and metabolic homeostasis by intestinal microbiota.

Trimethylamine oxide (TMAO) is an important microbiota-derived metabolite. Red meat and seafood intake is significantly associated with elevated serum TMAO levels (Abbasi, 2019; Hamaya et al., 2020; Wang et al., 2019b). Choline, L-carnitine, and betaine are decomposed into trimethylamine (TMA) by the intestinal microbiota community, and TMA is transformed into TMAO by hepatic flavin-containing monooxygenase 3 (FMO3) (Verhaar et al., 2020). The production of TMA depends on the cutC/cutD/cutA gene of intestinal microbiota (Rath et al., 2017; Roberts et al., 2018). Clinical trials have shown that cardiovascular disease is associated with high levels of TMAO in the blood (Abbasi, 2019). An increasing number of studies have shown that high levels of TMAO and TMAO precursors (carnitine, choline, betaine) are associated with a higher risk of cardiovascular disease (Fu et al., 2020; Komaroff, 2018; Tang et al., 2019). High levels of circulating TMAO are associated with an increased risk of diabetes and obesity (Dehghan et al., 2020; Zhuang et al., 2019). Trimethylamine oxide induces forkhead box O1 (Foxo1) expression by selectively activating the protein kinase R-like endoplasmic reticulum kinase to promote hyperglycemia (Chen et al., 2019). Dietary TMAO supplementation has been found to upregulate the expression of macrophage scavenger receptors (Wang et al., 2011). Long-term TMAO exposure was shown to reduce diet-induced glucose tolerance impairment and reduce adipocyte endoplasmic reticulum stress and lipolysis in mice. In addition, TMAO induces adipogenesis in palmitic acid treated HepG2 cells (Tan et al., 2019). However, limited studies have focused on the effects of TMAO on host fat deposition.

In our study, we performed a TMAO intervention trial in growing-finishing pigs and evaluated the effect of TMAO on fat accumulation, fatty acid composition, and intestinal microbiota.

2. Materials and methods

2.1. Animal ethics statement

The experiment was conducted (No. 20200505) in accordance with the guidelines of the Experimental Animal Ethics Committee of the Institute of Subtropical Agriculture (Changsha, Hunan Province, China). The experiment complied with the ARRIVE guidelines.

2.2. Animal experiment

A total of 16 growing pigs (26.00 ± 2.12 kg) with similar body weight and age were randomly divided into 2 groups (n = 8). The NC group were fed a basal diet and the TMAO group were fed the basal diet supplemented with 1 g/kg TMAO (Overland et al., 1999). All pigs had free access to feed and water. Before the experiment, the piggery was disinfected and all pigs were raised in separated pens. The experiment lasted 149 d. Feed intake and body weight were recorded during the experiment. After the experiment, the pigs were fasted for 24 h and weighed to calculate their average daily gain (ADG), average daily feed intake (ADFI), and feed conversion rate. The experiment was conducted in 2020 at the Institute of Subtropical Sciences, Chinese Academy of Sciences.

2.3. Chemical analysis of feed chemical compositions

The basal diet was formulated according to NRC (2012). The ingredients and chemical composition are shown in Table 1. Values of digestible energy were calculated from data provided by Feed Database in China (2020) (Tian et al., 2023). The crude protein (CP) contents in diet were measured using flow injection analysis (AA3, SEAL Analytical Instruments Co., Ltd, Germany) according to the method in DB43/T 2139-2021. The ether extract in diet was measured using a Soxhlet extractor according to method 920.39 (AOAC, 2006; Feng et al., 2023).

Table 1.

Ingredients and chemical composition of the basal diet (%, as-fed basis).

Ingredients Content Chemical composition2 Content
Stage 1
Corn meal 65.00 Digestible energy, MJ/kg 13.90
Soybean meal 25.00 Crude protein 16.75
Wheat bran 6.00 Ether extract 3.08
Premix1 4.00
Stage 2
Corn meal 65.00 Digestible energy, MJ/kg 13.79
Soybean meal 23.00 Crude protein 17.08
Wheat bran 8.00 Ether extract 2.99
Premix1 4.00
Stage 3
Corn meal 66.00 Digestible energy, MJ/kg 13.78
Soybean meal 22.00 Crude protein 16.43
Wheat bran 8.00 Ether extract 3.12
Premix1 4.00
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1

Per kilogram of premix contains 405.50 g limestone meal, 270.00 g rice bran, 150.00 g dicalcium phosphate, 80.00 g salt, 32.00 g lysine, 10.00 g choline, 5.20 g phytase, 16.00 g FeSO4·H2O, 8.00 g CuSO4·5H2O, 6.00 g ZnSO4·H2O, 4.00 g MnSO4·H2O, 1.00 g KI, 1.00 g Na2SeO3, 3.00 g multienzyme complex, 2.00 g feeding attractant, 1.00 g CoSO4·H2O, 1.00 g white sugar, 0.40 g Antioxidant-33 (active compound: ethoxy quinoline), vitamin A 24,375 IU, vitamin B1 7.5 mg, vitamin B2 18.75 μg, vitamin B6 9 mg, vitamin B12 75 mg, vitamin D3 7,500 IU, vitamin E 60 IU, vitamin K3 3 7.5 mg, biotin 0.45 mg, folic acid 1.0 mg, D-pantothenic acid 37.5 mg, and nicotinic acid 75 mg.

2

Values of digestible energy were calculated from data provided by Feed Database in China (2020). Crude protein and ether extracts were measured values.

2.4. Slaughter experiment

The slaughter experiment was performed according to the China National Standard (Ministry of Agriculture of the People's Republic of China, 2019, 2004). After removing the head, feet, tail and viscera (except kidneys), the carcass was weighed to calculate the slaughter rate. The remainder of the carcass was divided by skin, fat, muscle, and bone, then weighed to calculate lean body percentage, fat percentage, bone percentage, and skin percentage. The thickness of subcutaneous adipose tissue at the 6th to 7th ribs was measured as backfat thickness. In addition, the heart, liver, spleen, lung and kidney were also weighed. Carcass straight length and carcass oblique length were measured according to a previous study (Li et al., 2021b).

2.5. Serum biochemistry index

The serum biochemistry index was measured according to a previous study (Liu et al., 2015; Qi et al., 2022). Blood samples were centrifuged at 835 × g for 15 min, and the supernatant was taken for measurement. The serum albumin, total protein (TP), glutamic-oxaloacetic transaminase, glutamic-pyruvic transaminase, alkaline phosphatase (ALP), urea nitrogen, triglyceride, cholesterol, low-density lipoprotein, high-density lipoprotein, glucose, total bilirubin, creatinine, γ-glutamyl transpeptidase, D-lactate, uric acid, total bile acids, creatine kinase, lipase, complement C3, and complement C4 contents were determined using assay kits (Nanjing Jian Cheng Bioengineering Institute, Jiangsu, China) according to standard operating procedure.

2.6. Hematoxylin and eosin (H&E) staining

H&E staining was performed according to a previous method (He et al., 2022). Longissimus dorsi muscle, liver, and subcutaneous adipose tissue samples were fixed with 4% paraformaldehyde fix solution. Fixed samples were sheared, dehydrated, embedded, sliced, dyed, and sealed in accordance with standard operating procedure. Pathological sections were scanned using a 3D panoramic scanner (PANNORAMIC, 3DHISTECH (Hungary), Japan) (Skydsgaard et al., 2021).

2.7. Medium-long chain fatty acid profiles

The determination of medium-long-chain fatty acid profiles in longissimus dorsi muscle and liver were performed according to previous studies (Bai et al., 2019; Gao et al., 2020; Yin et al., 2017). The fat in the tissue samples was extracted with a mixture solution of benzene and petroleum. Extracted fat was rapidly methylated using 0.4 mol/L sodium hydroxide methanol solution. The supernatant was dried with anhydrous sodium sulfate and then was measured by HPLC (LC1290, Agilent, USA) according to the user's manual.

2.8. Real time fluorescence quantitative PCR

This experiment was conducted according to previous studies (Wang et al., 2016, 2021). The primer sequence is shown in Table 2. The tissue RNA was extracted using the Trizol method. Extracted DNA was tested by 1% agarose gel and a NanoDrop spectrophotometer (Thermo fisher scientific, Waltham, MA, USA) (1.8 < A260:A280 ratio < 2.1). Extracted DNA was reversely transcribed into cDNA by Evo M-MLV assay kits (Accurate Biology, Hunan, China). Then, the cDNA was used as a template to perform a PCR reaction in a fluorescence-based quantitative PCR instrument (Light Cycler 480II, Roche, Switzerland). GAPDH was used as the housekeeping gene. Data were analyzed by the 2−ΔΔCt method.

Table 2.

Primers used for RT-PCR.

Gene Primer sequences (5´ to 3´) Tm, °C
Pig primer
PPARα F: CGGGAAAGGCCAGCAAT 60
R: GGCCACGAGCGTCTTCTC
Elovl2 F: ATTCTTCACCACCAGCGAGG 60
R: TGCCTGGCTGTTATCACTCG
Elovl5 F: TACCACCATGCCACTATGCT 60
R: GACGTGGATGAAGCTGTTGA
Elovl6 F: TGAAATTGGGCAAACACTGTGT 60
R: AGGGCACTTTAAGCACCTCTACA
Ascl1 F: GAACAGGGTTGCTTTGCTTATT 60
R: GAGTTCGCCTTTGTTGATGATG
Scd F: AAGGAGCTGGTCAGTCGTTG 60
R: GCTTTCGAAGCTTTGTGCCA
FADS2 F: ACGGCCTTCATCCTTGCTAC 60
R: GTTGGCAGAGGCACCCTTTA
GAPDH F: ATGGTCCACATGGCCTCCAA 60
R: GAAGTCAGGAGATGCTCGGTG
FASD1 F: GTCACTGCCTGGCTCATTCT 60
R: AGGTGGTTCCACGTAGAGGT
ACC F: ATCCCTCCTTGCCTCTCCTA 60
R: ACTTCCCGTTCAGATTTCCG
FASN F: GCCGAGTACAGCGTCAACAACC 60
R: TGGTCCTTCTTCATCAGCGGGAT
CPT-1B F: ACTGTCTGGGCAAACCAAAC 60
R: CTTCTTGATGAGGCCTTTGC
LPL F: CTCGTGCTCAGATGCCCTAC 60
R: GGCAGGGTGAAAGGGATGTT
HSL F: GCAGCATCTTCTTCCGCACA 60
R: AGCCCTTGCGTAGAGTGACA
PPARγ F: TGACCATGGTTGACACCG 60
R: AAGCATGAACTCCATAGTGG
FABP4 F: TGGAAACTTGTCTCCAGTG 60
R: GGTACTTTCTGATCTAATGGTG
CD36 F: TGTGGATACTTGGAGGTGGG 60
R: TGCTGGTTGGAATACAGTGG
ATGL F: GCGAAAATGTCATCATAACC 60
R: ATGGTGCTCTTGAGTTCGT
SREBP1 F: GCACTTTCTGACCCGCTTCTTC 60
R: CTGCATGGCAACAGGCACCGA
C2C12 primer
GAPDH F: AAGAGGGATGCTGCCCTTAC 60
R: TACGGCCAAATCCGTTCACA
SREBP1 F: GCATGCCATGGGCAAGTAC 60
R: CCACATAGATCTCTGCCAGTGTTG
PGC1α F: AACCACACCCACAGGATCAGA 60
R: TCTTCGCTTTATTGCTCCATGA
ACC1 F: AAGTCCTTGGTCGGGAAGTATACA 60
R: ACTCCCTCAAAGTCATCACAAACA
FASN F: TGGTGAATTGTCTCCGAAAAGA 60
R: CACGTTCATCACGAGGTCATG
PPARγ F: ATGTCTCACAATGCCATCAGGTT 60
R: GCTCGCAGATCAGCAGACTCT
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PPARα = peroxisome proliferator activated receptor alpha; Elovl 2 = ELOVL fatty acid elongase 2; Elovl 5 = ELOVL fatty acid elongase 5; Elovl 6 = ELOVL fatty acid elongase 6; Ascl1 = achaete-scute family bHLH transcription factor 1; Scd = stearoyl-CoA desaturase; FADS2 = fatty acid desaturase 2; GAPDH = glyceraldehyde-3-phosphate dehydrogenase; FADS1 = fatty acid desaturase 1; Acc = acetyl-CoA carboxylase; FASN = fatty acid synthase; CPT-1B = carnitine palmitoyltransferase 1B; LPL = lipoprotein lipase; HSL = lipase E, hormone sensitive type; PPARγ = peroxisome proliferator activated receptor gamma; FABP4 = fatty acid binding protein 4; CD36 = CD36 molecule; ATGL = patatin-like phospholipase domain containing 2; SREBP1 = sterol regulatory element binding transcription factor 1; PGC1α = PPARG coactivator 1 alpha.

2.9. Determination of short chain fatty acids (SCFAs)

This work was performed according to a previous study (Li et al., 2019). Colonic contents and ileal contents were mixed with ultrapure water. Then the supernatant was reacted with 25% phosphoric acid solution for 3 h. Then the reaction mixture was tested using LC–MS (7890A, Agilent, USA).

2.10. DNA extraction and 16S rRNA sequencing

This work was conducted in accordance with a previous study (Li et al., 2021a). The bacterial DNA was extracted by the CTAB/SDS method, and assessed by an ultra-micro spectrophotometer (Nanodrop 2000, Thermo Fisher Scientific, USA) and agarose gel electrophoresis. The V3 to V4 region of bacterial 16S rDNA was sequenced by the Illumina Hiseq platform (515F (5′-barcode-GTGCCAGCMGCCGCGG-3′), 907R(5′-CCGTCAATTCMTTTRAGTTT-3′)). The data sequence was analyzed by the Quantitative Insights Into Microbial Ecology (QIIME) pipeline based on operational taxonomic units (OTUs). The sequence data is stored in the SRA database (PRJNA813690).

2.11. Determination of TMAO

This experiment was performed in accordance with a previous study (Jiang et al., 2021). Plasma TMAO was extracted using methanol. The supernatant was measured by UHPLC (1290 Infinity, Agilent, USA) and QTRAP (5500, AB SCIEX, USA).

2.12. Hydrolyzed amino acid profiles

The determination of hydrolyzed amino acid profiles in the longissimus dorsi muscle and liver was performed according to previous protocols (Bai et al., 2019; Long et al., 2016; Yin et al., 2017). Tissue samples were hydrolyzed with 6 mol/L hydrochloric acid solution at 110 °C for 22 h. After the filtrate was dried it was dissolved with 0.01 mol/L hydrochloric acid solution. Finally, the solution was tested in an amino acid analyzer (PF8700, Hitachi, Japan).

2.13. C2C12 cell culture

The C2C12 cell was cultured according to a previous study (Zhang et al., 2020). Mouse C2C12 cell line was purchased from American Type Culture Collection and cultured in Dulbecco's modified Eagle's medium containing 10% fetal bovine serum, 50 U/mL penicillin, and 50 μg/mL streptomycin at 37 °C in 5% CO2 atmosphere. C2C12 cells were seeded in 6 well plates to 70% to 80% confluency. Then, TMAO was added into the cells at a final concentration of 1 mM, 5 mM or 10 mM (Zou et al., 2022), and thereafter the cells were incubated for 48 h.

2.14. Data analysis

Experimental data was sorted by Excel 2019 (Microsoft, Redmond, WA, USA). Data were analyzed by Student's t-test using SPSS 22.0 (IBM, Armonk, NY, USA). Data are shown as mean ± SEM, and figures were plotted using Graphpad Prism 8 (La Jolla, CA, USA). The results were considered statistically significant at P < 0.05, and 0.05 ≤ P < 0.1 was considered a trend.

3. Results

3.1. TMAO increased fat deposition in growing-finishing pigs

The experimental design is shown in Fig. 1A. Dietary TMAO supplementation significantly increased the serum TMAO concentration (P < 0.05), and tended to decrease serum TMA content (P < 0.1) (Fig. 1B to C). Dietary TMAO supplementation did not change the ADG, ADFI, and feed conversation rate (Table 3), but tended to decrease the total lean body mass (P < 0.1), and increase the backfat thickness (P < 0.05) (Table 3). In addition, TMAO supplementation significantly increased serum contents of TP (Table 3).

Fig. 1.

Fig. 1

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Experimental design and bar graphs. (A) Experimental design. (B) Serum trimethylamine (TMA) contents. (C) Serum trimethylamine oxide (TMAO) contents. Data are shown as mean ± SEM. ∗∗P < 0.01. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet.

Table 3.

Effects of trimethylamine oxide (TMAO) on growth performance of finishing pigs.

Item NC TMAO P-value
ADG, g/d 572.11 ± 13.264 529.91 ± 29.421 0.21
ADFI, g/d 2108.96 ± 55.636 2060.20 ± 57.525 0.55
Feed conversion rate 3.69 ± 0.044 3.96 ± 0.195 0.21
Lean body mass, kg 49.46 ± 1.594 45.83 ± 1.284 <0.10
Backfat thickness, mm 18.30 ± 0.872b 20.89 ± 0.698a 0.04
Total protein, g/L 63.75 ± 1.828b 69.23 ± 1.726a <0.05
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ADFI = average daily food intake, ADG = average daily gain.

NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet.

Data are shown as mean ± SEM. Within a row, means without a common superscript differ at P < 0.05.

3.2. TMAO regulated ileal microbiota composition and increased ileal acetate concentration

TMAO is a microbiota-derived metabolite, but there are limited studies focus on its effect on intestinal microbiota composition. Therefore, we performed 16S rDNA sequencing of the ileal and colonic microbiota community. TMAO supplementation significantly increased the richness and evenness of ileal microbiota based on Shannon index (P < 0.05), but it did not change the β diversity of ileal microbiota (Fig. 2A to B). The ileal microbial community composition is shown in Fig. 2C. Linear discriminant analysis Effect Size analysis showed that Proteobacteria, and Escherichia–Shigella were significantly enriched in the ileal microbiota community of the TMAO group (LDA > 3, Fig. 2D). Moreover, carbohydrate degradation (superpathway of fucose and rhamnose degradation, D-glucarate degradation I, D-fructuronate degradation, D-galacturonate degradation I, fucose degradation, D-galactarate degradation I) was significantly enriched in the TMAO group (P < 0.05, Fig. 3). TMAO supplementation did not affect the diversity and richness of the colonic microbiota community (Fig. S1). TMAO had greater effect on the ileal microbial community than the colonic microbial community.

Fig. 2.

Fig. 2

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Trimethylamine oxide (TMAO) regulated ileal microbiota composition. (A) Ileal microbiota α diversity based on Simpson and Shannon indexes. (B) Ileal microbiota β diversity based on principal component analysis (PCA). (C) Ileal microbiota composition at phylum and genus level. (D) LEfSe analysis between NC group and TMAO group. Data are shown as mean ± SEM. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet.

Fig. 3.

Fig. 3

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Trimethylamine oxide (TMAO) regulated ileal microbiota function prediction. NAD = nicotinamide adenine dinucleotide. Data are shown as mean ± SEM. ∗P < 0.05. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet.

At the phylum level, TMAO increased the relative abundance of Proteobacteria (P < 0.05, Fig. 4A). At the genus level, TMAO significantly increased the relative abundance of Escherichia–Shigella (P < 0.05, Fig. 4B). What is more, TMAO supplementation significantly increased acetate content in the ileum (P < 0.05, Fig. 4C). Spearman analysis showed that ileal acetate concentration significantly correlated with the relative abundance of Escherichia–Shigella (P < 0.05, Fig. 4D).

Fig. 4.

Fig. 4

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Trimethylamine oxide (TMAO) regulated ileal acetate concentration in growing-finishing pigs. (A) The relative abundance of Proteobacteria. (B) The relative abundance of Escherichia–Shigella. (C) Ileal short chain fatty acid concentration. (D) Spearman analysis between ileal short chain fatty acid concentration and microbiota (genus level). Data are shown as mean ± SEM. ∗P < 0.05. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet.

3.3. TMAO reduced liver lipid accumulation and regulated fatty acid composition in liver

TMAO supplementation significantly decreased serum ALP levels (P < 0.05) (Fig. 5A). Representative H&E staining is shown in Fig. 5B. TMAO significantly decreased liver inflammation and histological score (P < 0.05), which indicates TMAO supplementation may contribute to alleviating liver injury (Fig. 5C). TMAO significantly decreased fat content in liver and significantly changed the medium-long chain fatty acid composition (P < 0.05, Fig. 5D to E). TMAO also significantly decreased polyunsaturated fatty acids (PUFA) and n-6 PUFA in liver (P < 0.05, Fig. 5F). TMAO decreased the relative abundance of C18:2n6c and increased the relative abundance of C20:0 in liver (P < 0.05, Fig. 5E). Furthermore, TMAO supplementation significantly decreased P-Ser, and Asp contents in liver (P < 0.05, Fig. 5F).

Fig. 5.

Fig. 5

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Trimethylamine oxide (TMAO) reduced inflammation and regulated fat deposition in the liver. (A) Serum alkaline phosphatase (ALP) content. (B) Representative image of hematoxylin and eosin (H&E) staining in liver. (C) Histological and inflammation scores. (D, E, and F) Fat content, fatty acid composition, and free amino acid composition in liver. Data are shown as mean ± SEM. ∗P < 0.05, ∗∗P < 0.01. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet. Saturated fatty acid (SFA) = C10:0 + C12:0 + C14:0 + C15:0 + C16:0 + C17:0 + C18:0 + C20:0 + C23:0; monounsaturated fatty acid (MUFA) = C14:1 + C16:1 + C17:1 + C18:1n9t + C18:1n9c + C20:1 + C24:1; polyunsaturated fatty acid (PUFA) = C18:2n6t + C18:2n6c + C18:3n6 + C18:3n3 + C20:3n6 + C20:4n6 + C20:5n3 + C22:6n6; n-3 PUFA = C18:3n3 + C20:5n3; n-6 PUFA = C18:2n6t + C18:2n6c + C18:3n6 + C20:3n6 + C20:4n6 + C22:6n6; UFA = unsaturated fatty acid; Asp = aspartate; P-Ser = phosphoserine.

3.4. TMAO increased intramuscular fat content and regulated fatty acid composition in longissimus dorsi muscle

As the H&E staining shows, TMAO did not affect the myofiber area in longissimus dorsi muscle (Fig. 6A to C, P < 0.05). TMAO significantly increased intramuscular fat content in longissimus dorsi muscle and it also significantly regulated medium-long chain fatty acid composition (P < 0.05, Fig. 6D to E). Specifically, TMAO significantly increased saturated fatty acid (SFA) percentage (C16:0, and C18:0) and n-3 PUFA percentage and decreased n-6 PUFA and unsaturated fatty acid (UFA) percentages in longissimus dorsi muscle (P < 0.05, Fig. 6E). TMAO increased C18:2n6C and C20:3n6 percentages in longissimus dorsi muscle (P < 0.05, Fig. 6E). TMAO significantly upregulated the mRNA expression of sterol-regulatory element binding protein 1 (SREBP1) and downregulated the expression of fatty acid desaturase 2 (FADS2) (P < 0.05, Fig. 6F). We also investigated the effect of TMAO on fat deposition in C2C12 cells, the 48 h TMAO treatment at 1 mM significantly increased the expression of SREBP1 and fatty acid synthase (FASN) (P < 0.05, Fig. 6G).

Fig. 6.

Fig. 6

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Trimethylamine oxide (TMAO) increased fat deposition and regulated fatty acid composition. (A) Representative H&E staining image in longissimus dorsi muscle. (B) Myofiber area of longissimus dorsi muscle. (C) Myofiber number/Microscope. (D) Intramuscular fat contents. SFA = C10:0 + C12:0 + C14:0 + C15:0 + C16:0 + C17:0 + C18:0 + C20:0 + C23:0; MUFA = C14:1 + C16:1 + C17:1 + C18:1n9t + C18:1n9c + C20:1 + C24:1; PUFA = C18:2n6t + C18:2n6c + C18:3n6 + C18:3n3 + C20:3n6 + C20:4n6 + C20:5n3 + C22:6n6; n-3 PUFA = C18:3n3 + C20:5n3; n-6 PUFA = C18:2n6t + C18:2n6c + C18:3n6 + C20:3n6 + C20:4n6 + C22:6n6. (E) Fatty acid composition in longissimus dorsi muscle. (F) The mRNA expression in longissimus dorsi muscle. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet. Elovl 2 = ELOVL fatty acid elongase 2; Elovl 5 = ELOVL fatty acid elongase 5; Elovl 6 = ELOVL fatty acid elongase 6; Ascl1 = achaete-scute family bHLH transcription factor 1; SCD = stearoyl-CoA desaturase; FADS2 = fatty acid desaturase 2; FASN = fatty acid synthase; CPT-1B = carnitine palmitoyltransferase 1B; SREBP1 = sterol regulatory element binding transcription factor 1; PGC1α = PPARG coactivator 1 alpha. (G) Effect of TMAO treatment on fat deposition in C2C12 cells (n = 3). PPARγ = peroxisome proliferator activated receptor gamma; SREBP1 = sterol regulatory element binding transcription factor 1; PGC1α = PPARG coactivator 1 alpha; ACC = acetyl-CoA carboxylase; FASN = fatty acid synthase. NC group, 0 mM TMAO treatment; TMAO1 group, 1 mM TMAO treatment; TMAO5 group, 5 mM TMAO treatment; TMAO10 group, 10 mM TMAO treatment. Data are shown as mean ± SEM. ∗P < 0.05, ∗∗P < 0.01.

3.5. TMAO increased adipocyte area in subcutaneous adipose tissue

Consistent with the previous results, TMAO significantly increased the mean adipocyte area in subcutaneous adipose tissue (P < 0.05, Fig. 7A to C), and TMAO supplementation also significantly downregulated the expression of carnitine palmitoyltransferase-1B (CPT-1B) (P < 0.05, Fig. 7D). TMAO did not affect medium-long chain fatty acid composition.

Fig. 7.

Fig. 7

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Trimethylamine oxide (TMAO) decreased lipolysis in subcutaneous adipose tissue. (A) H&E stained image. (B) Mean adipocyte area. (C) Adipocyte number. (D) The mRNA expression in subcutaneous adipose tissue. FAS = fatty acid synthase; ACC = acetyl-CoA carboxylase; HSL = lipase E, hormone sensitive type; ATGL = patatin-like phospholipase domain containing 2; FABP4 = fatty acid binding protein 4; LPL = lipoprotein lipase; CD36 = CD36 molecule; CPT-1B = carnitine palmitoyltransferase 1B; GAPDH = glyceraldehyde-3-phosphate dehydrogenase. Data are shown as mean ± SEM. ∗P < 0.05. NC group, basal diet; TMAO group, 1 g/kg TMAO supplementation in the basal diet.

4. Discussion

Fat deposition is a dynamic equilibrium process of anabolism and catabolism. Fat is synthesized from glycerol and fatty acids. Although most fatty acids come from de novo lipogenesis, fatty acids absorbed from the gut also contribute to fat synthesis in pig (O'hea and Leveille, 1969). In addition, carbohydrates, short-chain fatty acids, and keto acids are also used for lipogenesis and its products are mainly palmitic acid and stearic acid (Nürnberg et al., 1998). Lipogenesis is involved in the cytoplasm and mitochondria. The cytoplasm uses acetyl CoA to produce palmitic acid, while the mitochondria extend palmitic acid and desaturate saturated fatty acids (Ameer et al., 2014). Lipogenesis is modulated by FASN and acetyl-CoA carboxylase lipids are mobilized for energy supply during energy deficit, such as fasting. Lipolysis is mainly carried out in the mitochondria and modulated by HSL, CPT1, LPL, and ATGL enzymes. Fatty acids produced by lipolysis are oxidized in the mitochondria, and CPT1 is a key rate-limiting enzyme in β-oxidation (Morigny et al., 2021).

Accumulated studies have proven that the intestinal microbiota plays an important role in fat deposition and energy homeostasis (Chen et al., 2021; Turnbaugh et al., 2009). Existing literature suggests that TMAO is an important intestinal microbiota-derived metabolite which is closely associated with a series of diseases, such as obesity and metabolic syndrome. Ihubacter massiliensis also contributes to TMAO production in animal models (Wu et al., 2020). There are few studies on the effect of TMAO on the intestinal microbiota and our studies showed that dietary TMAO supplementation increased the richness and diversity of ileal microbiota. TMAO did not affect the α diversity and β diversity of the colonic microbiota. Moreover, TMAO had a limited effect on colonic microbiota function capacity (data not shown). Yoo et al. (2021) reported that a high-fat diet increased circulating TMAO levels by promoting the relative abundance of Escherichia coli, and our study also showed that TMAO increased the relative abundance of Escherichia–Shigella (Yoo et al., 2021). In addition, KEGG analysis showed that carbohydrate degradation was significantly enriched in the TMAO group; these results indicated that TMAO increased the relative abundance of Escherichia–Shigella to enhance carbohydrate degradation and acetate production in the ileum. Intragastric administration of acetic acid significantly decreased hepatic lipid accumulation in high-fat-fed mice and upregulated PPARα, UCP2, and CPT-1 in HepG2 cells (Kondo et al., 2009). Yamashita et al. (2009) showed that acetate treatment increased the mRNA expression of myoglobin (Myog) and Glut4 in the abdominal muscle. Nevertheless, a previous study also reported that acetate can promote the adipogenic differentiation of IM-BAT cells and significantly upregulates the mRNA expression of PPARγ in IM-BAT cells, which also promotes the differentiation of 3T3-L1 cells by upregulating the mRNA expression of PPARγ2 via activated G-protein coupled receptor 43 (GPR43) (Hong et al., 2005; Hu et al., 2016), but this result needs to be demonstrated in vitro.

Alkaline phosphatase (ALP) mainly exists in hepatocytes, and increased serum ALP levels are an important biomarker for acute liver injury. Our study showed that TMAO supplementation decreased serum ALP contents. Further, H&E staining showed that TMAO supplementation significantly decreased the inflammation score and histological score in the liver. Previous studies also showed that injection of TMAO significantly alleviated liver fibrosis in CCl4-challenged mice (Zhou et al., 2022), and oral 120 mg/kg TMAO administration significantly alleviated high-fat high-cholesterol diet-induced steatohepatitis in rats (Zhao et al., 2019). Therefore, we speculated that TMAO has a beneficial effect on liver injury. Moreover, TMAO supplementation decreased the fat content in the liver and this is consistent with a previous study of Zhao et al. (2019). There is also some literature suggesting that TMAO increases fat deposition in the liver (Tan et al., 2019b), however we speculate that this may be due to the dose and animal model being different. In our study, TMAO decreased C18:2n6c (linoleic acid) abundance in the liver. Our study showed that TMAO improved liver injury, decreased fat deposition, and decreased PUFA abundance in the liver. A meta-analysis showed that increased microbiota-derived TMAO increases the risk of obesity (Dehghan et al., 2020). In agreement with a previous study (Dehghan et al., 2020), our results have also shown that TMAO significantly increased backfat thickness in pigs. CPT1 is a key rate-limiting enzyme in β-oxidation, and is located in the mitochondrial outer membrane. It catalyzes the conversion of long chain fatty acids to acetyl-carnitine (Schlaepfer and Joshi, 2020; Yoon et al., 2020). Our results showed that TMAO downregulated the mRNA expression of CPT-1B in subcutaneous adipose fat. These results indicated TMAO may inhibit lipolysis to promote fat deposition in subcutaneous adipose tissue in pigs. Intramuscular fat content is an important factor in determining meat quality and our results showed that TMAO significantly increased intramuscular fat content in longissimus dorsi muscle. SREBP1 is a nuclear transcription factor which is directly involved in fatty acid biosynthesis, triglyceride synthesis, and glucose metabolism (Guo et al., 2014; Hagen et al., 2010). SREBP1 was significantly upregulated in the TMAO group, indicating that TMAO may upregulated SREBP1 to increase intramuscular fat content in longissimus dorsi muscle. Mammals do not have Δ 15 desaturase and Δ 12 desaturase and cannot synthesize linoleic acid and linolenic acid. Linoleic acid (C18:2n6c) and linolenic acid (C18:3n3) are essential fatty acids. Our results also showed that TMAO significantly increased the percentages of gamma linoleic acids C18:2n6, C22:3n3, and C20:3n6, and decreased the percentage of C20:4n6 in longissimus dorsi muscle. FADS2 is a key gene in the polyunsaturated fatty acid anabolism pathway, which can convert linoleic acid and α-linolenic acid into γ-linolenic acid and stearidonic acid (Glaser et al., 2010; Zhang et al., 2016). The mRNA expression of FADS2 was significantly downregulated in the TMAO group, indicating that TMAO increased the percentage of C18:2n6 via FADS2 in longissimus dorsi muscle, however these results need to be further verified in vitro.

5. Conclusion

Taken together, our study demonstrated that TMAO modulated ileal microbiota composition and increased ileal acetate concentration by increasing the relative abundance of Escherichia–Shigella. TMAO also regulated fat distribution in different tissues, which increased fat deposition in longissimus dorsi muscle and subcutaneous adipose tissue, and decreased fat deposition in the liver. In addition, TMAO also decreased the percentage of C18:2n6c in the liver, and increased the percentage of C18:2n6c in longissimus dorsi muscle. In conclusion, our results indicated that dietary TMAO supplementation significantly regulated fat distribution in pigs.

Author contributions

Andong Zha and Wanquan Li: Writing–Original draft preparation, Conducting the research; Jing Wang and Ming Qi: Investigation; Ping Bai, Peng Liao, Bie Tan and Yulong Yin: Research design, Funding acquisition.

Declaration of competing interest

We declare that we have no financial and personal relationships with other people or organizations that can inappropriately influence our work, and there is no professional or other personal interest of any nature or kind in any product, service and/or company that could be construed as influencing the content of this paper.

Acknowledgments

This research was supported by the National Natural Science Foundation of China (U20A2054), the earmarked fund for China Agriculture Research System (CARS-35), and the Science and Technology Major Project of Yunnan Province (202202AE090032), and Large-scale Healthy Breeding Technology Research and Industrialization Demonstration of pig (202102AE090046).

Footnotes

Peer review under responsibility of Chinese Association of Animal Science and Veterinary Medicine.

Supplementary data to this article can be found online at https://doi.org/10.1016/j.aninu.2023.12.006.

Appendix supplementary data

The following is the Supplementary data to this article.

Multimedia component 1
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