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Objective: A high intake of fat in the diet plays a crucial role in promoting obesity and obesity-related pathologies, and especially visceral obesity is closely associated with obesity-related complications. Because adipose tissue is anatomically associated with lymph nodes, the secondary lymphoid organ, we hypothesized that fat tissue-derived factors may influence the cellularity of lymphoid tissue embedded in fat.
Methods and Procedures: Mesenteric and inguinal lymph nodes were isolated from obese mice fed a high-fat diet and control mice fed a regular diet. T-cell population, activation state, and the extent of apoptosis were determined by flow cytometric analysis or terminal deoxynucleotidyl transferase biotin-dUTP nick end labeling (TUNEL) assay.
Results: The weight of mesenteric lymph nodes and the total number of lymphoid cells in the obese mice significantly decreased compared with those in the control mice; however, no change was observed in the weight of inguinal lymph nodes. The numbers of CD4+ and CD8+ T cells in the mesenteric lymph nodes of obese mice significantly decreased compared with those of the control. Enhanced T-cell activation and apoptosis were observed in the mesenteric lymph node cells of the obese mice. The treatment of lymph node cells with free fatty acids, oxidative stress, and chylomicrons, which are obesity-related factors, resulted in lymph node T-cell activation and apoptosis.
Discussion: These results suggest that visceral fat accumulation with a high-fat diet can cause the atrophy of mesenteric lymph nodes by enhancing activation-induced lymphoid cell apoptosis. Dietary fat-induced visceral obesity may be crucial for obesity-related immune dysfunction.
Visceral obesity is an important determinant of the risk of developing obesity-related pathologies, such as insulin resistance, type 2 diabetes, and atherosclerosis. Visceral fat exhibits more enhanced lipolytic activity and inflammatory phenotypes than other fat depots (1,2). Thus, the vicious metabolic and inflammatory phenotypes of visceral fat are more closely implicated in the development of obesity-related complications than other fat depots. A high intake of fat in the diet is also associated with the obesity-related pathologies, independent of obesity and body fat localization (3). Obesity, apart from causing metabolic complications, impairs the immune system. For example, obese patients or animals are more susceptible to infectious diseases as well as several types of cancer (4,5). However, it is unclear whether the vicious phenotypes of visceral fat are associated with immune dysfunction in obesity.
Lymph nodes, one of the secondary lymphoid organs packed with lymphocytes, macrophages, and dendritic cells, are the major sites for antigen delivery to mount immune responses. Interestingly, most major lymph nodes are embedded in adipose tissue. It has been postulated that adipose tissue surrounding lymph nodes supplies immune cells with the energy to induce immediate and effective responses of the adjacent lymph node to a foreign antigen (6,7,8,9), indicating that metabolic cross talk exists between tissues. Adipose tissue functions not only as an energy storage organ but also as a secretory organ producing various biologically active molecules called as adipocytokines (10), thus, fat tissue-derived factors may also be crucial for the cross talk between adipose tissue and lymphoid cells. Because mesenteric fat tissue, which is a representative visceral fat depot with special vicious phenotypes, is anatomically associated with mesenteric lymph nodes, mesenteric adipose tissue-derived factors may directly affect the mesenteric lymphoid cell system by a local interaction or cross talk, thereby affecting the immune system in obesity.
In this study, we tested the hypothesis that the vicious phenotypes of the obese visceral fat tissue surrounding mesenteric lymph nodes influence lymphoid cellularity. Our data suggest that visceral fat tissue-derived factors are crucial for the induction of activation-induced lymphoid cell death, leading to the atrophy of mesenteric lymph nodes of obese mice.
Methods and Procedures
Male C57BL/6 mice at 8 weeks of age were purchased from ORIENT (Pusan, Korea). All the mice were maintained under specific pathogen-free condition in the animal facility of the Immunomodulation Research Center (University of Ulsan, Ulsan, Korea). The mice were given free access to autoclaved water and irradiated pellet food. The mice were fed a high-fat diet (45% calories from lard and soybean oil; Research Diets, New Brunswick, NJ; 21.22% energy as fat; 48.48% as carbohydrate; 17.01% as protein, and 0.15% cholesterol) or a standard pellet diet (13% calories from soybean oil; Harlan Teklad, Madison, WI) for 3 months. Body weight was measured every 3 days.
Preparation of lymph node cells
Mice were killed under CO2 anesthesia. Inguinal lymph nodes were carefully removed from inguinal adipose tissues. After entering the abdominal cavity, the ileocecal region was inspected, and mesenteric chain with four nodes were removed from the adjacent mesentery before resecting the right colon. Lymph nodes were aseptically removed, placed in cold phosphate-buffered saline, and immediately homogenized. The cells were filtered through a nylon mesh to remove tissue debris. Erythrocytes were lysed in a lysing buffer (1.7 mmol/l Tris and 0.14 mol/l ammonium chloride). Pooled (3–5 mice/experiment) lymph node cells were suspended in RPMI-1640 supplemented with 10% heat-inactivated, fetal bovine serum, 20 mmol/l L-glutamine, 10 mg/l penicillin-streptomycin, (Gibco BRL, Grand Island, NY) and 2 mg/l gentamicin (Gibco BRL, Grand Island, NY), and were then counted.
Flow cytometric analysis (FACS)
Cells isolated from lymph nodes were incubated with Fcγ receptor blocking antibodies (2.4G2) for 10 min on ice, and the cells were double stained with phycoerythrin-conjugated anti-CD3, anti-CD4, anti-F4/80, anti-CD11b, anti-Dx5a or anti-CD11c, anti-Foxp3 or anti-B220 antibody, fluorescein isothiocyanate-conjugated anti-CD4, anti-CD8, anti-CD11b, anti-Gr1 or anti-CD3 or Cy-conjugated anti-TcRαβ, anti-TcRγδ. For intracellular staining, cells were fixed and permeabilized with Cytofix/Cytoperm (BD Pharmingen, San Diego, CA) for 15 min at room temperature (RT). Then, the cells were stained with monoclonal antibody Foxp3. The cells were also stained with monoclonal antibodies for CD4, CD8, CD25, CD44, CD62L, and Annexin V. After incubation with the antibodies, the cells were washed with fluorescence-activated cell sorting (FACS) buffer and analyzed on a FACSCalibur (BD Biosciences, San Jose, CA) with CellQuest software (BD Biosciences).
Mesenteric lymph nodes were isolated from the obese and control mice, fixed for 24 h in 10% formalin, and embedded in paraffin. Eight-micrometer thick sections were obtained, mounted on two glass slides, and stained with hematoxylin-eosin.
For the in situ detection of DNA fragmentation in frozen tissue sections, terminal deoxynucleotidyl transferase biotin-dUTP nick end labeling (TUNEL) assay was performed using the TACS TdT kit (R&D, Minneapolis, MN), following the manufacturer's instructions. Briefly, sequential 8-μm thick tissue sections were fixed on precleaned slides (Fisher Scientific) and left to dry at RT. Subsequently, the sections were dehydrated. Protein digestion was performed by incubating the sections in 20 μg/ml proteinase K (Worthington, Lakewood) for 15 min at RT. Endogenous peroxidase was inactivated with 3% H2O2 in distilled water for 5 min at RT. The labeling mixture containing biotinylated dUTP in TdT enzyme buffer was added to the sections and incubated at 37 °C in a humified chamber for 1 h. After stopping the enzymatic reaction, sections were rinsed with phosphate-buffered saline, covered with antidigoxigenin peroxidase conjugate, and incubated for 30 min at RT in a humidified chamber. Then, the sections were incubated in Tris-buffered saline with 0.05% diaminobenzidine plus 3% H2O2 until a color reaction was observed. Finally, the sections were washed, counterstained in methyl green, dehydrated, and mounted in a mounting solution. Nuclei that were stained dark brown were considered to be apoptotic cells when visualized at a magnification of ×200 using a light microscope.
Preparation of chylomicrons
Blood was collected in EDTA-treated evacuated tubes and plasma was prepared immediately by centrifugation (910 × g, 4 °C, 10 min). Chylomicrons were then isolated from 5 ml plasma layered under 5 ml NaCl (9 g/l) by ultracentrifugation (120,000 × g, 30 min, 10 °C). The chylomicron layer was removed carefully using a pipette. Triacylglycerols (TG) was assayed in chylomicrons using enzymatic colorimetric methods with commercial kit (A-san pham., Hwa Sung, Korea).
Preparation of fatty acid-albumin complexes
Palmitate sodium salt (C 16:0) and bovine serum albumin were purchased from Sigma-Aldrich. Palmitate was combined with fatty acid-free and low endotoxin bovine serum albumin at a molar ratio of 10:1 (fatty acid: albumin) in RPMI-1640 medium containing 10% heat-inactivated, fetal bovine serum at 50 °C for 6 h for a final palmitate concentration of 0.5–2 mmol/l as described previously (11). Fatty acid-albumin complex solution was freshly prepared prior to each experiment.
Lymph node cell culture
Lymphocytes were plated at a density of 5 × 106 cells/ml in RPMI-1640 medium. T cells were treated with free fatty acid (palmitate 0.5–2 mmol/l), hydrogen peroxide (H2O2, 10–50 μmol/l), monocyte chemoattractant protein-1 (MCP-1, 10–100 ng/ml), chylomicrons (1–100 mg% TG). After incubating for different time periods (12–48 h), the cells were collected and used for FACS or MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay.
MTT assay was used to measure lymphocyte viability. MTT is a pale yellow substrate that produces a dark blue formazan product when incubated with viable cells. MTT rings are cleaved in active mitochondria, and this process occurs only in viable cells. After the supernatant was removed, the cells were incubated at 37 °C with MTT (0.05 mg/ml) for 4 h and absorbance was then measured at 570 nm using an enzyme-linked immunosorbent assay reader (Molecular Devices, Sunnyvale, CA).
Measurement of free fatty acid
Adipose tissue (200 mg of frozen tissue) was homogenized in 400 μl of 20 mmol/l Tris-HCl buffer (pH 7.4). Free fatty acid contents in fat samples were determined in the adipose tissue homogenate by the colorimetric determination of nonesterified fatty acids using the NEFA kit (Wako Chemicals, Osaka, Japan).
Determination of reactive oxygen species generation
Stromal vascular fractions (SVFs) were obtained from the mesenteric and subcutaneous adipose tissues of the obese and nonobese mice. The adipose tissue-derived SVF cells (50,000 cells/sample) were used for the measurement of reactive oxygen species (ROS) production by FACS. The SVF cells were incubated with 2 μmol/l of oxidation sensitive dye 2′, 7′-chlorodihydrofluorescein diacetate (DCFDA; Molecular Probe; Sigma-Aldrich) and were incubated at 37 °C for 30 min. Samples were analyzed by FACSCalibur (BD Biosciences, San Jose, CA).
Measurement of malondialdehyde and 4-hydroxy-2(E)-nonenal levels
Measurement of malondialdehyde (MDA) and 4-hydroxy-2(E)-nonenal (4-HNE) are end products derived from the peroxidation of polyunsaturated fatty acids and related esters. The measurement of these aldehydes provides a convenient index of lipid peroxidation. In this study, we evaluated the adipose tissues ROS levels by measuring these products of lipid peroxidation according to manufacturer's instructions. Briefly, adipose tissue was homogenized in 2 vol of ice-cooled 20 mmol/l Tris-HCl buffer (pH 7.4). The supernatant of this homogenate was used to determine the lipid peroxide concentration, which was determined using a spectrophotometric assay kit (Lipid peroxidation assay kit; Calbiochem, La Jolla, CA).
All experiments were repeated at least three times. The results were expressed as means ± s.e.m. Statistical analysis was performed using ANOVA and Duncan's multiple range test. Differences were considered to be significant at P < 0.05.
Tissue weight, number of lymphoid cell, and the extent of apoptosis in mesenteric lymph nodes of obese mice
Body weights were significantly higher for the obese mice (38.55 ± 0.52 g) fed a high-fat diet than for the control mice (27.42 ± 1.188 g) fed a regular diet, and the adiposity in the obese mice (mesenteric fat tissue: 1.04 ± 0.12 g; epididymal fat: 2.29 ± 0.07 g; renal fat: 0.83 ± 0.03 g; subcutaneous fat: 1.45 ± 0.14 g) significantly (P < 0.001) increased compared with that in the control (mesenteric fat: 0.2 ± 0.03 g; epididymal fat: 0.55 ± 0.05 g; renal fat: 0.18 ± 0.02 g; subcutaneous fat: 0.31 ± 0.03 g) (Figure 1a). The weight of mesenteric lymph nodes of obese mice significantly (P < 0.005) decreased compared with that of the control (Figure 1b). The total number of lymphoid cells in the mesenteric lymph nodes of the obese mice also significantly (P < 0.005) decreased compared with those of the control; there were no differences in this parameter in the inguinal lymph nodes (Figure 1c). In addition, a cross-section of the lymph nodes showed numerous follicles in the cortex, some of which contain germinal centers, and the central medulla, and no morphological abnormality was observed in the mesenteric lymph nodes of the obese mice despite the size reduction of the lymph nodes (Figure 1d). To determine whether apoptotic cell death is associated with the atrophy of mesenteric lymph nodes, TUNEL assay was performed. TUNEL-positive cells that were stained with brown were detected in the mesenteric lymph nodes of the obese mice, but not in those of the control mice (Figure 1e). These results indicate that lymphoid cell death results in the atrophy of mesenteric lymph nodes in obese mice fed a high-fat diet.
Characterization of lymphocytes in lymph nodes of obese and nonobese mice
To characterize lymphocytes subsets in lymph nodes, we performed FACS. The percentages of CD4+ cells (29 ± 2.5% in obese, 33 ± 2.9% in nonobese) and CD8+ cells (21 ± 2.0% in obese, 25 ± 2.5% in nonobese) in the mesenteric lymph nodes of the obese mice were not significantly different from those in the nonobese mice (data not shown). In consistent with previous report (11,12), lymph node lymphocytes consisted of a majority of T cells (50 ± 5.1% in obese, 57 ± 5.2% in nonobese) and B cells (25 ± 2.9% in obese, 27 ± 4.6% in nonobese) with a fewer T lymphocytes, natural killer cells, and natural killer T cells (data not shown). When we normalized the number of cell per gram of lymph nodes, the number of immune cells (e.g., T cells, macrophages, natural killer cells, dendritic cells, and B cells) in the obese mice significantly (P < 0.05) decreased compared with those in the control (Table 1). The number of CD4 or CD8 positive T cells was 36%, 43% lower in the mesenteric lymph nodes compared with those in the control. The number of antigen presenting cells (dendritic cells and macrophages) was 30%, 37% lower in the mesenteric lymph nodes of the obese mice compared with those in the control. Interestingly, the number of CD4+Foxp3+ population (0.050 ± 0.011 in the obese, 0.083 ± 0.022 in the nonobese mice) was decreased in the mesenteric lymph nodes of the obese mice compared with those in the nonobese control (Table 1). Moreover, the number of CD4+CD25+ T-cell population was decreased in the obese compared with those in the nonobese mice, while CD8+CD25+ T-cell populations were rarely detected (Table 1). These results indicate that regulatory T-cell numbers in the mesenteric lymph nodes decreased in the obese mice compared with the nonobese control.
Table 1. Immune cell profile in lymph nodes from obese and nonobese mice
T-cell activation and apoptosis in mesenteric lymph nodes of obese mice
To determine whether T-cell activation results in T-cell apoptosis in mesenteric lymph nodes, the levels of CD62L and CD44 expression, which are T-cell activation markers, were measured by FACS. The percentage of CD4+ expressing CD62Llow or CD44high in mesenteric lymph node cells of obese mice increased compared with that of the control (Figure 2a). The percentage of CD8+ expressing CD62Llow or CD44high also in the mesenteric lymph node cells of obese mice increased compared with that of the control (Figure 2a). Moreover, we determined the expression levels of Annexin V, which is an apoptosis marker, in the mesenteric lymph node lymphocytes of the obese and control mice. The percentage of CD4+ or CD8+ and Annexin V double positive cells increased in the mesenteric lymph node lymphocytes of obese mice (Figure 2b), indicating that activation-induced T-cell death occurs in mesenteric lymph node cells.
Effect of obesity-related factors on activation-induced lymphoid T-cell apoptosis
To determine whether obesity-related factors such as free fatty acids induce mesenteric lymph node cell death, we examined the effect of free fatty acids on lymph node cell viability by MTT assay. The cells were treated with or without various concentrations of palmitate for different time periods (12 h, 24 h, 48 h), and then cell viability was measured. As shown in Figure 3a, a dose- and time-dependent reduction in viability of mesenteric lymph node cells was observed. Oxidative stress has been reported to have a proapoptotic effect (12). Mesenteric lymph node cell viability decreased dose- and time-dependently in the presence of H2O2 (Figure 3a), which generates ROS. Because MCP-1 in mesenteric adipose tissue plays a key role in enhanced inflammatory response in obese visceral fat tissue (2), we examined whether MCP-1 induces a reduction in mesenteric lymph node cell survival. MCP-1 showed no effect on mesenteric lymph node cell viability (data not shown). In addition, dietary fat absorbed mostly as chylomicrons into lymph system after triglycerides are resynthesized. We observed that the exposure of mesenteric lymph node cells to chylomicrons resulted in the lymphoid cell death (Figure 3b). FACS also revealed that palmitate and H2O2 treatment increased the percentages of CD4+/CD62L−, CD8+/CD62L− cells, CD4+/Annexin V+ cells, and CD8+/Annexin V+ cells in the obese mice compared with those in the control (Figure 3c), indicating that T-cell activation and apoptosis occurred in the mesenteric lymph node T cells of the obese mice. These results indicate that obesity-related factors such as free fatty acids and oxidative stress as well as a high-fat diet cause lymphoid T-cell apoptosis.
To determine whether the Fas/FasL pathway is involved in lymphocyte apoptosis, mesenteric lymph node lymphocytes isolated from the nonobese mice were incubated with anti-FasL with or without palmitate for 48 h. As shown in Figure 3d, palmitate-induced cell death was significantly inhibited by the anti-FasL antibody; however, the nonimmune isotype of hamster IgG had no effect, indicating that FasL is involved in free fatty acid-induced cell death in obese mice.
Oxidative and metabolic status in mesenteric lymph nodes of obese mice
Our in vitro data revealed that obesity-related factors such as free fatty acids and oxidative stress cause lymphoid T-cell apoptosis. We further examined the oxidative (ROS, MDA and 4-HNE levels) and metabolic status (release of free fatty acids) of the adipose tissues surrounding the mesenteric and inguinal lymph nodes in obese and/or control mice. As shown in Figure 4a, the concentrations of free fatty acid were higher in the mesenteric adipose tissue than those in the subcutaneous adipose tissue of obese mice. We also measured ROS production in adipose tissue-derived SVF. The production of ROS of mesenteric adipose tissue SVF from the obese mice was significantly greater than that of the control SVF, while there was no difference in the subcutaneous SVF (Figure 4b). The total numbers of adipose tissue SVF markedly increased in the obese mice (mesenteric: 5.6 × 105 cells/tissue g; subcutaneous: 4.0 × 105 cells/tissue g) compared with those in the control (mesenteric: 3.9 × 105 cells/tissue g; subcutaneous: 3.3 × 105 cells/tissue g). Therefore, oxidative stress may increase in the mesenteric adipose tissue of the obese mice. MDA and 4-HNE are end products derived from peroxidation of polyunsaturated fatty acids and related ester. The levels of MDA and 4-HNE also significantly increased in the mesenteric adipose tissue from the obese mice compared with those from the nonobese control mice (Figure 4c). In addition, the levels of MDA and 4-HNE tended to be higher in the obese mesenteric adipose tissue than in the obese subcutaneous adipose tissue.
This study demonstrates that visceral fat accumulation causes the atrophy of mesenteric lymph nodes in obese mice by inducing T-cell activation-mediated apoptosis.
Adipose tissue plays an important role in providing energy to lymphoid cells in immune defense (9). Interestingly, we observed for the first time that the weight of mesenteric lymph nodes surrounding mesenteric fat tissue and the total number of lymphoid cells significantly decreased in the obese mice fed a high-fat diet compared with the control fed a regular diet; however, no difference in either parameter was observed in inguinal lymph nodes embedded in subcutaneous fat tissue. Apoptotic bodies were detected in the mesenteric lymph nodes from the obese mice, indicating that the atrophy of mesenteric lymph nodes results from the apoptosis in mesenteric lymph nodes. It has been shown that visceral adipose tissue elicits highly lipolytic and inflammatory phenotypes (1,2), and is considered to be a source of high concentrations of free fatty acids and inflammatory adipocytokines in obesity. In this regard, the increase in the amounts of mesenteric adipose tissue-derived factors with increasing fat mass may affect the microenvironment of the mesenteric lymphoid tissue embedded in fat tissue, leading to lymphoid cell death.
Apoptosis, or programmed cell death, contributes to the homeostasis of multicellular organisms. Apoptosis resulting from T-cell activation referred to as activation-induced T-cell death, is crucial for the maintenance of T-cell homeostasis. Studies have shown that obesity impairs immunity, causing lymphopenia in obese human (13,14). T-cell populations and their functions were reduced in human obesity, and this was related to the elevated tumor necrosis factor-α production. Furthermore, the T-cell dysfunction could be recovered by adequate weight reduction (14). Several lines of evidence suggest that environmental factors, such as nonlymphoid-derived substances and cytokines, modulate activated T-cell apoptosis. For example, various mediators (e.g., free fatty acids, inflammatory cytokines, and oxidative stress) can trigger apoptosis in antigen-presenting cells (15,16,17,18) and induce activation-induced lymphocyte apoptosis (19,20,21). It is noteworthy that all the mediators increase in obese adipose tissues, particularly visceral fat tissue. Two adhesion molecules, CD62L (L-selectin) and CD44 (homing-associated cell adhesion molecule), are associated with T-cell activation. Naive T cells exhibit a CD62LhighCD44low phenotype, whereas activated T cells exhibit a CD62LlowCD44high phenotype (22). Our observation in the expressions of T-cell activation markers, such as CD62L and/or CD44, clearly showed that T cells from the mesenteric lymph nodes are partially activated in obese mice. We also found that the extent of apoptosis of the mesenteric lymph node T cells of the obese mice increased three- to fourfold compared with that of the control. These results indicate that activation-induced T-cell death occurs in the mesenteric lymphoid cells of obese mice.
Inflammatory cytokines, free fatty acids, and oxidative stress, which are representative vicious phenotypes of obese visceral adipose tissue (23,24), have been reported to induce T-cell activation, leading to apoptosis (19,20,21), suggesting that obesity-related factors contribute to the activation-induced death of lymphoid cells adjacent to fat tissue. Mesenteric adipose tissues have special vicious phenotypes including macrophage accumulation and activation (2), inflammatory cytokine/chemokine gene expression (2,25,26), and lipolysis sensitivity (27,28). Regarding lipolytic activity, visceral adipocytes, compared with subcutaneous fat cells, are more sensitive to catecholamine-induced lipolysis, and less sensitive to insulin's antilipolytic effects (1). Moreover, recent study has shown that the ratio of visceral to subcutaneous expression of hormone sensitive lipase and lipoprotein lipase genes increased twofold in obese dogs fed a high-fat diet, suggesting enhanced lipolytic activity in the visceral fat depot relative to subcutaneous fat (29). Indeed, we observed that the levels of free fatty acid in the mesenteric adipose tissue significantly increased in the obese mice compared with those in the nonobese mice, presumably reflecting the enhanced lipolytic activity in the tissue. Because free fatty acid can trigger apoptosis in many cell types including lymphocytes and antigen-presenting cells (17,21), mesenteric adipose tissue-derived free fatty acid may be associated with fewer lymphoid cells including lymphocytes, macrophages, and dendritic cells in the mesenteric lymph node of obese mice. Indeed, our in vitro study showed that the exposure of mesenteric lymph node cells to obesity-related factors, such as free fatty acids and oxidative stress, results in activation-induced T-cell apoptosis. Activated T cells undergo apoptosis through Fas (CD95) or FasL (CD95L), which is considered to be a critical mechanism in inflammatory processes (30). We observed that activation-induced T-cell death induced by obesity-related factors, such as free fatty acids or oxidative stress, is partially abrogated by the an anti-CD95L monoclonal antibody. We also observed site-specific differences in the oxidative (ROS production and MDA/4-HNE) and metabolic status (release of free fatty acids) of the adipose tissues surrounding the mesenteric and inguinal lymph nodes in obese animals. These findings suggest that the site-specific vicious phenotypes of mesenteric adipose tissue may contribute to mesenteric lymph node cellularity and atrophy in obese mice by inducing activation-induced lymphocyte apoptosis.
Recent study has shown that a high-fat diet changes intestinal microbiota and induces the death of Gram-negative bacteria in gut, causing endotoxemia, and the metabolic endotoxemia can initiate obesity and insulin resistance (31). It has been shown that lipopolysaccharide induces apoptosis in lymphocytes in vivo, and lipopolysaccharide-induced apoptosis increases in lymphocytes both in the cortex of the thymus and in the germinal centers and paracortical areas of mesenteric lymph nodes (32,33). Mesenteric lymph nodes are gut-associated secondary lymphoid organs where soluble dietary antigens are presented to naive T cells (34). Therefore, the atrophy of mesenteric lymph nodes, in our observation, may be associated with the metabolic endotoxemia induced by the high-fat diet. In addition, most dietary fat is transported from the gut into mesenteric lymph, principally as triacylglycerol-rich chylomicron composed of lipids and several different apolipoproteins (35). We observed that the exposure of mesenteric lymph node cells to chylomicron resulted in the lymphoid cell death. It has also been shown that the very low-density lipoprotein + chylomicron fraction of the plasma inhibited lymphocyte proliferation in vitro (36). These results indicate that the high-fat diet per se may also contribute to the lymph node atrophy.
It is noteworthy that regulatory T-cell population decreased in the mesenteric lymph nodes of obese mice. Regulatory T cells are crucial for the maintenance of immunological self-tolerance and immune homeostasis by suppressing aberrant or excessive immune responses, and thus are implicated in autoimmune diseases and allergy (37,38,39). Deletion or mutation of transcription factor Foxp3 interferes with the generation and/or function of CD4+CD25+ regulatory T cells (40,41). A decrease in the total number of circulating CD4+CD25+ regulatory T cells and their suppressive activity have been found in patients suffering from autoimmune diseases, such as type I diabetes, multiple sclerosis, autoimmune hepatitis, and systemic lupus erythematosus (42). Obesity, apart from causing metabolic complications, impairs the immune system. For example, obese patients or animals are more susceptible to infectious diseases as well as allergic diseases, such as asthma and atopy (43). From this point, it may be speculated that regulatory T cells play a role in obesity-related immune dysfunctions. Further studies are necessary to address this intriguing phenomenon.
In conclusion, our data demonstrate that a high-fat diet-induced mesenteric adipose tissue accumulation causes the atrophy of the mesenteric lymph nodes of obese mice. Visceral fat-derived factors such as free fatty acids and oxidative stress can induce activation-induced T-cell death in mesenteric lymph nodes. Thus, dietary fat-induced visceral fat accumulation may play a role in immune dysfunction in obesity.
This work was supported by the Science Research Center Fund to Immunomodulation Research Center at the University of Ulsan from the Korea Science and Engineering Foundation (KOSEF) and the Korean Ministry of Science and Technology, and partially supported by a grant from KOSEF (R01-2005-000-10408-0). S.-C.L. was supported by Brain Korea 21 Program project. T.K. was supported by the Research and Development Program for New Bio-industry Initiatives and Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sport, Science and Technology of Japan (15081205).