Galectin-3 Deficiency Accelerates High-Fat Diet–Induced Obesity and Amplifies Inflammation in Adipose Tissue and Pancreatic Islets

Male Gal-3–deficient (LGALS3^−/−) mice on the C57BL/6 background and their littermate controls, wild-type (WT) C57BL/6 mice (6 weeks of age), obtained from the University of California Davis (Davis, CA; by courtesy of D.K. Hsu and F.T. Liu) were fed either a low-fat diet (LFD; 3% fat) or HFD (60% fat) obtained from Mucedola (Milano, Italy) for 11 or 18 weeks. All animal procedures were approved by the ethical committee (04/11) of the Faculty of Medical Sciences, University of Kragujevac.

Body weights and fasting blood glucose levels were measured once every 2 weeks. Mice were fasted for 4 h, and glucose levels (mmol/L) were determined using the Accu-Chek Performa glucometer (Roche Diagnostics, Mannheim, Germany). Serum concentrations of lipids and HbA_1c were measured using the Olympus AU600 Chemistry Immuno Analyzer (Olympus, Tokyo, Japan) and fasting insulin in sera using the Rat/Mouse Insulin ELISA Kit (Millipore Corporation, Billerica, MA). Homeostasis model assessment of insulin resistance (HOMA-IR) was calculated as previously described (20).

Total visceral adipose tissue (VAT) was minced, placed in PBS containing 1 mg/mL collagenase type II and 2% BSA (Sigma-Aldrich, St. Louis, MO), incubated for 1 h in a water bath at 37°C, and passed through a 40-µm nylon cell strainer (BD Biosciences, San Jose, CA) to enrich stromal vascular fraction cells.

Dissected pancreata were pooled, minced, and digested using 2 mg/mL collagenase type V in Hanks’ balanced salt solution (Sigma-Аldrich) with 10% FCS for 15 min in a water bath at 37°C (21), and digests were passed through a 40-µm cell strainer.

Spleens were excised and single-cell suspensions were obtained by mechanical disruption and lysis of RBCs.

Cells were labeled with fluorochrome-conjugated monoclonal antibodies: anti-mouse CD3, CD4, CD8, CD44, CD62L, CD279 (PD-1), CD11b, interferon-γ (IFN-γ), IL-17 (BD Biosciences), NK1.1, F4/80, CD206, and CD11c antibodies (BioLegend, San Diego, CA). For intracellular staining, cells were activated with PMA/ionomycin and processed as previously described (22). Primary rabbit anti-mouse NLRP3, IL-1β, and NF-κB (phospho-S536) antibody (1 μg/mL; Abcam, Cambridge, U.K.) and secondary goat anti-rabbit antibody labeled with PE-Cy5.5 (1/400; Invitrogen, Carlsbad, CA) were used. Cells were analyzed with the FACSCalibur Flow Cytometer (BD Biosciences), and analysis was conducted with FlowJo (Tree Star).

Pancreatic tissue cryostat sections were stained with hematoxylin-eosin. Histological analysis of the distribution of inflammatory cell infiltrate in pancreatic islets was performed in blinded fashion by two independent observers. The images were captured with a light microscope (BX51; Olympus) equipped with a digital camera. Insulitis was graded and a mean insulitis score was calculated (23).

Immunofluorescent staining of pancreatic tissue cryosections (5 μm) was performed using rabbit anti-mouse NLRP3 (1:200), IL-1β (1:200), IL-18 (1:200), RAGE (1:400), and AGE (1:200) antibodies (Abcam), followed by incubation with goat anti-rabbit IgG antibody PE-Cy 5.5, followed by FITC-conjugated anti-mouse insulin antibody (1:400; Abcam). The sections were mounted with ProLong Gold antifade reagent with DAPI (Invitrogen) and analyzed at ×40 magnification using the Nikon Eclipse Ti-E inverted research microscope equipped with NIS-Elements Imaging software (Nikon Instruments Inc., Melville, NY). Only brightness and contrast were adjusted.

Pancreatic tissue cryosections were incubated with biotinylated F4/80 antibody (Invitrogen) followed by visualization using the Mouse-Specific HRP/DAB Detection IHC Kit (Abcam). Next, they were incubated with rabbit anti-mouse NLRP3, IL-1β, AGE, RAGE, and Gal-3, visualized by rabbit-specific conjugate (Expose Rb-Specific HRP/AEC Detection IHC Kit; Abcam), photomicrographed with a digital camera mounted on light microscope (Olympus BX51), digitized, and analyzed (24). Results are expressed as percent positive staining in the specified region.

Macrophages were collected from the peritoneal cavity of mice under sterile conditions and cultured in complete Dulbecco’s modified Eagle’s medium supplemented with 10% FCS at 37°C in a 5% CO₂ incubator. After pretreatment with lipopolysaccharide (LPS) (100 ng/mL) for 4 h, cells were stimulated with BSA alone or palmitate-BSA (100 μmol/L), glucose (22 mmol/L), and H₂O₂ (10 µmol/L) (all from Sigma-Aldrich) for 24 h. Where indicated, cells were preincubated with the caspase-1 inhibitor Z-YVAD-FMK (10 μmol/L; Bachem AG, Bubendorf, Switzerland).

For the determination of caspase-1 activity, we used the Caspase-1 Colorimetric Kit (R&D Systems) according to the manufacturer's recommendations.

Peritoneal macrophages were seeded on six-well plates (1 × 10⁶ cells/well), washed once with 2 mL small interfering RNA (siRNA) transfection medium, transfected with 60 pmol NLRP3 siRNA or 60 pmol negative control siRNA, and incubated for 7 h according to the manufacturer’s instructions (Santa Cruz Biotechnology, Santa Cruz, CA).

Pancreatic tissue lysates were prepared in a solution containing 62.5 mmol/L Tris-HCl, pH 6.8, 2% weight/volume (wt/vol) SDS, 10% glycerol, 50 mmol/L dithiothreitol, 0.01% wt/vol bromophenol blue, 1 mmol/L phenylmethylsulphonyl fluoride, 1 μg/mL aprotonin, and 2 mmol/L EDTA, and proteins from VAT were isolated with TRIzol reagent (Genosys, Woodlands, TX) and further processed (25). Rabbit polyclonal IgG antibodies to ASC (1:500), NLRP3 (1:500), IL-1β (1:2500), caspase-1 (1:500), NF-κB p65 (phospho-S536) (1:500), and NF-κB p65 (1:500) (all from Abcam) and mouse anti-mouse IgG1 β-actin antibody (1:500) (Sigma-Aldrich) as internal control were used, followed by donkey anti-rabbit horseradish peroxidase at 1:10,000 or sheep anti-mouse horseradish peroxidase (1:2,500) (GE Healthcare, Buckinghamshire, U.K.). Detection was performed by chemiluminescence (ECL; GE Healthcare), and photographs were made by X-ray films (Kodak). Scion Image Alpha 4.0.3.2 (Scion Corporation, Frederick, MD) was used to calculate protein expression.

Cytokine levels were determined using mouse duo sets for IL-1β/IL-1F2, IL-6, IL-4, IL-13, IL-10, IFN-γ, IL-17, and CRP (R&D Systems, Minneapolis, MN, USA) according to the manufacturer's recommendations.

All data are presented as means ± SEM. Statistical significance was determined by ANOVA (one-way ANOVA and Bonferroni post hoc test), independent-sample Student t test, and, where appropriate, Kruskal-Wallis and Mann-Whitney U test. Relationships between variables were assessed using Pearson correlation. Statistical significance was assumed at P < 0.05. Statistical analyses were performed using SPSS 13.0.

Body weight (Fig. 1A) and total VAT weight (Fig. 1B) were significantly higher in HFD-fed LGALS3^−/− mice compared with other experimental groups at both 11 and 18 weeks, and significantly greater in WT mice on HFD versus LFD at 18 weeks (Fig. 1A and B). There was no difference in food intake between LGALS3^−/− and WT mice (Fig. 1A). Fasting blood glucose levels and insulinemia and HbA_1c were significantly increased in HFD-fed LGALS3^−/− mice compared with other experimental groups at both 11 and 18 weeks, whereas in WT mice on HFD versus LFD at 18 weeks, the differences did not reach statistical significance (Fig. 1C and D). Hyperglycemia was noticed in LGALS3^−/− mice fed LFD at 18 weeks (Fig. 1C). Of note, the significantly increased body weight (26.25 ± 0.66 vs. 22.19 ± 0.67 g, P = 0.010) and fasting glycemia (8.92 ± 0.47 vs. 6.82 ± 0.31 mmol/L, P = 0.004) were already noticed after 3 weeks of HFD in LGALS3^−/− mice compared with diet-matched WT mice. HbA_1c (%) and fasting glycemia significantly correlated in LGALS3^−/− mice at 18 weeks (Fig. 1C). HOMA-IR was significantly higher in HFD-fed LGALS3^−/− mice compared with other experimental groups at 11 and 18 weeks (Fig. 1E). Total, HDL, and non-HDL cholesterol serum levels increased with HFD but were not significantly different between the genotypes, and serum levels of triglycerides were similar between experimental groups at 11 weeks (data not shown).

Significantly increased CD3⁺ T cells and CD3⁺NK1.1⁺ NKT cells were found in VAT in HFD-fed LGALS3^−/− mice (Fig. 2A) with markedly increased CD3⁺NK1.1⁺ IFN-γ⁺ type 1 NKT cells, and CD3⁺IFN-γ⁺ type 1 T cells were significantly higher compared with LFD-fed mice of both genotypes at 11 weeks (Fig. 2B). The incidence of CD4⁺ or CD8⁺ T-naïve, memory or effector populations and CD3⁺IL-17⁺ cells was not different between groups (data not shown). CD4⁺CD25⁺FoxP3⁺ Tregs were significantly lower in the VAT and spleens (P = 0.071) of obese LGALS3^−/− mice compared with other experimental groups (Fig. 2C and D). CD4⁺PD-1⁺ cells were significantly increased in the VAT and spleens with HFD in LGALS3^−/− mice (Fig. 2E).

We next monitored the recruitment of macrophages into adipose tissue after 11 weeks on HFD to look for potential mechanisms of glucose metabolism abnormalities in obese LGALS3^−/− mice. Proinflammatory F4/80⁺CD11c⁺CD206⁺ macrophages significantly increased in adipose tissue of LGALS3^−/− mice fed HFD compared with other experimental groups (Fig. 3A). HFD increased splenic F4/80⁺CD11b⁺CD11c⁺ bone marrow–derived dendritic cells (BMDCs) in mice of both genotypes (data not shown). After 18 weeks on HFD, adipose tissue BMDCs were significantly higher in LGALS3^−/− compared with WT mice (72.63 ± 3.91 vs. 38.04 ± 10.54, P = 0.006). Alternatively, activated F4/80⁺CD11c⁻CD206⁺ M2 macrophages were markedly reduced in VAT from both LGALS3^−/− and WT mice fed HFD compared with LFD-fed WT mice at 11 weeks (Fig. 2B), and further decreased in LGALS3^−/− mice fed HFD for 18 weeks, significantly lower compared with diet-matched WT mice (0.93 ± 0.093 vs. 7.66 ± 2.99, P = 0.046).

In LGALS3^−/− mice, proinflammatory F4/80⁺CD11c⁺CD206⁺ adipose tissue macrophages significantly correlated with body weight (r = 0.919, P = 0.001) and fasting blood glucose levels (r = 0.912, P = 0.002) (Fig. 3C), whereas Tregs inversely correlated with body weight (r = 0.633, P = 0.067) and fasting glycemia (r = 0.819, P = 0.007) (Fig. 3D) in contrast to WT mice (data not shown).

Serum cytokine levels were measured to evaluate the systemic inflammatory profile in studied mice. CRP, IL-6, and IL-4 levels significantly increased in the sera of LGALS3^−/− mice with HFD, whereas IL-13 and IL-10 levels significantly decreased compared with diet-matched WT mice at 11 weeks (Fig. 4A). After 18 weeks on HFD, IL-4 and IL-1β levels were significantly higher, whereas IL-10 levels remained significantly lower in LGALS3^−/− mice (Fig. 4B), with no significant difference in the serum levels of IFN-γ (Fig. 4A and B) and IL-17 (data not shown) compared with WT mice.

Given the fact that LGALS3^−/− mice exhibited obesity-related abnormalities of glucose metabolism, we performed histological analysis of the distribution of inflammatory cell infiltrate in pancreatic islets from LGALS3^−/− and WT mice fed HFD or LFD for 11 weeks (Fig. 5A). We found significantly increased average insulitis as expressed by the insulitis score in HFD-fed LGALS3^−/− mice compared with other experimental groups (Fig. 5B). The percentage of total intact islets and those with perivascular/periductal and peri-insulitis was 58% in LGALS3^−/− mice compared with 89% in WT mice, both on HFD (P < 0.001). The percentages of islets with mild or severe insulitis were 24 and 17%, respectively, in HFD-fed LGALS3^−/− mice compared with 10% of islets with mild insulitis and none with severe insulitis in WT mice fed HFD (both P < 0.001). Of note, the percentages of islets with mild or severe insulitis were 17 and 5% in LFD-fed LGALS3^−/− mice compared with 2% of islets with mild insulitis (P < 0.05) and none with severe insulitis in WT mice fed LFD (P < 0.05). The percentage of islets with mild insulitis was significantly higher in HFD-fed WT mice compared with LFD-fed WT mice (P < 0.05).

Flow cytometric analysis of mononuclear cells isolated from digested pancreata of obese LGALS3^−/− mice revealed that the majority of cells were F4/80⁺CD11b⁺ myeloid cells, among which the proinflammatory F4/80⁺CD11c⁺CD11b⁺ BMDCs were the predominant cell type (Fig. 5C).

We next sought to assess NLRP3 inflammasome and IL-1β expression in pancreatic islets of mice fed HFD for 11 weeks. Immunofluorescence and morphometric immunohistochemical data showed increased NLRP3 inflammasome expression in islets of HFD-fed LGALS3^−/− mice (Fig. 6A, top) compared with HFD-fed WT mice (Fig. 6A, bottom) with a significantly higher percentage of NLRP3 inflammasome–positive area in islets of obese LGALS3^−/− mice (Fig. 6B) and prominent infiltration of F4/80⁺ macrophages (Fig. 6C). Also, increased IL-1β expression in islets from obese LGALS3^−/− mice (Fig. 6D, top) compared with HFD-fed WT mice (Fig. 6D, bottom) was found with a significantly higher percentage of IL-1β–positive area in islets of obese LGALS3^−/− mice (Fig. 6E and F). IL-18 was not detected (data not shown). Furthermore, we show significantly higher percentages of NLRP3 inflammasome– and IL-1β–expressing macrophages derived from digested pancreatic tissue (Fig. 6G) and VAT (Fig. 6H) from HFD-fed LGALS3^−/− mice compared with diet-matched WT controls.

Western blot analyses showed a similar expression of NLRP3 inflammasome in pancreata among experimental groups and significantly increased expression of ASC in HFD-fed LGALS3^−/− mice compared with LFD-fed LGALS3^−/− and WT mice (both P < 0.05). Procaspase-1 was significantly increased in HFD-fed LGALS3^−/− mice compared with other experimental groups (all P < 0.05) and mature caspase-1 in comparison with LFD-fed LGALS3^−/− mice (P < 0.05). ASC expression was significantly higher in HFD-fed WT mice compared with LFD-fed LGALS3^−/− mice (P < 0.05) (Fig. 6I).

In VAT, a significant increase of NLRP3 inflammasome expression was found in HFD-fed LGALS3^−/− mice compared with LFD-fed LGALS3^−/− and WT mice (both P < 0.05) and increased ASC and procaspase-1 in comparison with LFD-fed LGALS3^−/− mice (both P < 0.05). Mature caspase-1 protein expression was significantly higher in obese LGALS3^−/− mice compared with other experimental groups (all P < 0.05). NLRP3 and ASC expression were significantly lower in LFD-fed LGALS3^−/− mice compared with HFD or LFD-fed WT mice (all P < 0.05). Procaspase-1 was significantly lower in LFD-fed LGALS3^−/− mice compared with HFD-fed WT mice (P < 0.05) (Fig. 6J).

As Gal-3 has a role in the uptake and removal of metabolic compounds with proinflammatory properties related to diabetes complications, we examined the expression of AGE and RAGE in pancreatic islets. Immunofluorescence and immunohistochemical (IHC) data showed increased accumulation of AGE (Fig. 7A and B) and an increased percentage of RAGE-positive area in islets in LGALS3^−/− mice fed HFD for 11 weeks compared with diet-matched WT mice (Fig. 7C and D).

We show increased expression of Gal-3 protein in the pancreatic islets of HFD-fed WT mice compared with LFD-fed WT mice at 11 weeks (Fig. 7E).

We demonstrate the inherent genotype differences in the cellular response of LGALS3^−/− mice vs. WT mice as LPS (100 ng/mL), palmitate-BSA (100 µmol/L), and LPS plus palmitate induced significantly higher IL-1β and IL-6 production in LGALS3^−/− peritoneal macrophages (Fig. 8A). IL-1β production was significantly reduced in the presence of the caspase-1 inhibitor Z-YVAD-FMK (10 µmol/L), whereas IL-6 production was not affected (Fig. 8A). No significant difference in IL-1β production between LGALS3^−/− and WT mice was observed when cells were stimulated with high glucose or H₂O₂ with and without priming with LPS (data not shown).

In vitro stimulation of cells with LPS or palmitate significantly increased percentages of NLRP3 inflammasome–expressing F4/80⁺ macrophages (Fig. 8B) and significantly increased caspase-1 activity (Fig. 8C) in LGALS3^−/− compared with WT macrophages. Transfection of LGALS3^−/− peritoneal macrophages with NLRP3 siRNA resulted in a significant reduction of the NLRP3 inflammasome expression compared with cells transfected with control siRNA (15.6 vs. 3.7%) (Fig. 8D). Transfection with NLRP3 siRNA, but not with control siRNA, led to a significant decrease in IL-1β production by LGALS3^−/− macrophages stimulated with palmitate alone and LPS plus palmitate (Fig. 8D).

We next tested for the differences in NF-κB activation in LGALS3^−/− vs. WT mice by using an antibody against the NF-κB p65 subunit phosphorylated at the amino acid residue serine 536, an activated form of NF-κB that is induced in response to a variety of proinflammatory stimuli, including LPS. Figure 8E represents the percentage of phospho–NF-κB p65–expressing peritoneal macrophages in response to LPS (1 μg/mL) with and without palmitate (100 µmol/L). A significant increase of phospho–NF-κB p65–expressing F4/80⁺ macrophages in LGALS3^−/− compared with WT cells in response to LPS and LPS plus palmitate was observed (Fig. 8E).

Western blotting showed significantly increased expression of phospho–NF-κB p65 in pancreata of LGALS3^−/− fed HFD for 18 weeks compared with other experimental groups (all P < 0.05) (Fig. 8F). Phospho/total ratio of NF-κB p65 was the highest in the pancreata of HFD-fed LGALS3^−/− mice (0.816) compared with HFD-fed WT mice (0.413) and LFD-fed LGALS3^−/− mice (0.120) or WT animals (0.345). In VAT, phospho–NF-κB p65 expression was significantly higher in HFD-fed LGALS3^−/− mice compared with LFD-fed LGALS3^−/− mice (P < 0.05) (Fig. 8F). Phospho/total ratio of NF-κB p65 was the highest in the VAT of HFD-fed LGALS3^−/− mice (0.842) compared with HFD-fed WT mice (0.469) and LFD-fed LGALS3^−/− mice (0.370) or WT animals (0.272).

This study provides evidence that knockdown of Gal-3 results in accelerated HFD-induced obesity and diabetes, as indicated by the significantly higher body weight and total VAT weight, hyperglycemia, increased HOMA-IR, severe insulitis, and pronounced infiltration of pancreatic islets with proinflammatory macrophages. Ongoing inflammation in largely expanded adipose tissue in obese LGALS3^−/− mice was characterized by the increased incidence of type 1 CD3⁺ T and CD3⁺NK1.1⁺ NKT lymphocytes expressing IFN-γ. Th1 effector cells have a major role in obesity-associated chronic inflammation (26), and NKT cell depletion prevents diet-induced metaflammation and glucose intolerance (27). Reduced adipose tissue and splenic Tregs in obese LGALS3^−/− mice are in line with the data of depleted adipose tissue Tregs during obesity (28). HFD increased the incidence of activated CD4⁺PD-1⁺ T cells in VAT and spleens in LGALS3^−/− mice, possibly due to enhanced TCR-mediated signaling in T cells in Gal-3 deficiency (10,29).

F4/80⁺CD11c⁺CD206⁺ macrophages, increased in the VAT of obese LGALS3^−/− mice, were recently described as a novel macrophage subset to facilitate adipose tissue inflammation and insulin resistance in obese human subjects (30). Eighteen weeks of HFD in the VAT of LGALS3^−/− mice increased proinflammatory F4/80⁺CD11b⁺CD11c⁺ BMDCs, cells that have a crucial role in obesity-induced inflammation (31). Markedly reduced alternatively activated M2 macrophages in the VAT of LGALS3^−/− mice are in line with the reported data of Gal-3 involvement in the polarization of M2 macrophages (32,33).

Accelerated HFD-induced obesity in Gal-3–deficient mice was associated with systemic inflammation. Increased IL-6 and IL-1β induced by HFD in LGALS3^−/− mice are in line with the data of increased production of IL-1β and IL-6 in Gal-3–deficient macrophages during infection (34). Increased systemic IL-4 might indicate enhanced T-cell function during nutrient excess in LGALS3^−/− mice as Gal-3–deficient CD4⁺ T cells were shown to produce increased amounts of IFN-γ and IL-4 after T-cell receptor engagement (35). Lack of Gal-3 on dendritic cells favors lower IL-10 production and increased IFN-γ by allogeneic T cells (36). Reduced levels of IL-13 and IL-10 in the sera of obese LGALS3^−/− mice might contribute to amplified obesity-induced inflammation. IL-10 has a protective role in type 2 diabetes by increasing insulin sensitivity in skeletal muscle (37).

The most striking finding in our study was the presence of severe insulitis in obese LGALS3^−/− mice, but also in lean LGALS3^−/− mice. Enhanced expression of Gal-3 protects rat pancreatic β-cells from the cytotoxic effect of IL-1β (18), and Gal-3 is highly expressed in islet endothelial cells in obesity-induced diabetes in mice (19), but its role in diabetes progression still remains to be elucidated. Moreover, we show upregulation of Gal-3 protein in pancreatic islets in HFD-fed WT mice. F4/80⁺CD11c⁺CD11b⁺ BMDCs were the most abundant cell type in islet infiltrates in obese LGALS3^−/− mice, and higher expression of NLRP3 inflammasome and IL-1β in F4/80⁺ macrophages reflects ongoing inflammation in islets of HFD-fed LGALS3^−/− mice. Apart from macrophages, β-cells could produce IL-1β when exposed to proinflammatory metabolic signals, possibly through NLRP3 inflammasome activation (38). Most recently, type 2 diabetes has been considered an autoinflammatory disease with the central role for NLRP3-ASC inflammasome–mediated IL-1β production (39,40). We did not find a significant increase in protein content of NLRP3 inflammasome in whole-tissue lysates of pancreas and VAT in HFD-fed LGALS3^−/− mice compared with HFD-fed WT mice. However, a trend toward increased ASC and NLRP3 inflammasome protein content was found in pancreas and VAT, respectively. The increased mature caspase-1 protein expression found in the VAT of obese LGALS3^−/− mice might indicate increased NLRP3 inflammasome activation in adipose tissue in Gal-3 deficiency. In fact, LGALS3^−/− macrophages produced increased amounts of IL-1β, which was caspase-1 dependent, and had increased NLRP3 inflammasome expression and higher caspase-1 activity in response to LPS and/or palmitate compared with WT cells. In addition, silencing of NLRP3 by siRNA attenuated IL-1β production by LGALS3^−/− macrophages, suggestive of the enhanced NLRP3 inflammasome activation in Gal-3–deficient mice. The most recent study shows that NLRP3 expression is increased by TLR agonists, including LPS, in murine macrophages in an NF-κB–dependent manner (41). Moreover, LGALS3^−/− macrophages produced increased amounts of IL-1β and IL-6 in response to LPS stimulation, which is line with the evidence of markedly elevated LPS-induced proinflammatory cytokines in Gal-3–deficient macrophages since Gal-3 binds LPS and prevents its activity (42). HFD strongly increases intestinal permeability and increases blood levels of endotoxin (43). Endotoxin, free fatty acids, and AGE, through TLRs and RAGE, activate NF-κB and lead to IL-1β and tumor necrosis factor-α production, thus promoting insulin resistance (39,44). LGALS3^−/− macrophages had increased expression of phospho–NF-κB p65 in response to LPS, which complements the reported data (45). Increased pancreatic and also VAT phospho–NF-κB p65 expression could be related to increased accumulation of AGE and upregulation of RAGE in islets of LGALS3^−/− mice fed HFD. These findings indicate that additional NF-kB–mediated proinflammatory pathways (46) might operate in amplified obesity-induced inflammation in LGALS3^−/− mice.

Collectively, the amplified obesity-induced inflammation in adipose tissue and pancreatic islets in LGALS3^−/− mice suggest a protective role for Gal-3 in obesity and type 2 diabetes, which could be of therapeutic relevance.

This work was supported by grants from the Serbian Ministry of Science and Technological Development (175071 and 175069) (Belgrade, Serbia).

No potential conflicts of interest relevant to this article were reported.

N.N.P. and J.M.P. researched data and wrote the manuscript. I.P.J., G.D.R., M.Z.M., and I.G.N. researched data. N.S.Z. and A.L.D. researched data and contributed to discussion. N.N.A. reviewed and edited the manuscript. M.L.L. contributed to discussion and reviewed and edited the manuscript. M.L.L. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

We would like to thank Milan Milojevic, Dragana Djurdjevic, and Ivana Milovanovic (Center for Molecular Medicine and Stem Cell Research) and Dragana Djacic (University of Kragujevac) for technical assistance.