Purple corn extract induces long-lasting reprogramming and M2 phenotypic switch of adipose tissue macrophages in obese mice

Eight-week old male C57BL/6J mice (n = 64 Charles River, Calco, Italy), were used and maintained at 23 ± 3 °C, 12 h light cycle (08.00–20.00). The sample size was determined a priori using a power analysis in order to reach 80% power at an alpha-level of 0.10 to detect a strong effect size considering effects reported in previous research [10, 11, 14]. Mice were randomly divided into 3 groups and assigned to either control diet (20% protein, 70% carbohydrate, and 10% fat; 3.85 kcal/g; #D12450B, Research Diets; New Brunswick, NJ, USA) or HF diet (20% protein, 35% carbohydrate and 45% fat; 4.73 kcal/g; #D12451, Research Diets) for 12 weeks along with water or RED extract supplied in drinking water at an ACN concentration of 290 mg/Kg body weight/day. Diets and drinks were replaced weekly and daily respectively, to prevent oxidization. Mice were weighed weekly and daily, food and water/RED extract intakes were recorded throughout the study. The RED extract was produced by SVEBA Srl (Appiano Gentile, Italy) to a final concentration of 40 mg/g of ACNs, as previously described [11]. After 12 weeks, liver, spleen, and epididymal adipose tissue were carefully removed and weighed. All procedures involving animals and their care conformed to institutional guidelines in compliance with national (D.L. N.116, G.U., suppl. 40, 18-2-1992; D.L. N. 26, G.U. 4-3-2014) and international law and policies (EEC Council Directive 2010/63/EU, OJ L 276/33, 22-09-2010; NIH Guide for the Care and Use of Laboratory Animals, US National Research Council-2011) and approved by the University of Milan Animal Welfare Body and by the Italian Minister of Health (Project number 7/2013). All efforts were made to minimize animal suffering and to reduce the number of animals used.

Powdered plant material (100 mg of corn cob) was extracted twice with 400 µl of 80% MeOH with 2% formic acid. Prior to the injection 100 µl of extract were mixed with 25 µl of solvent A (water with 0.5% formic acid). After centrifugation 5 µl of mixture were injected per sample. Phenylpropanoids were separated by reverse-phase HPLC (Acquity UPLC™ with a CSH Phenyl-Hexyl, 1.7 µm, 2.1 × 100 mm column; Waters, Eschborn, Germany) at 25 °C. At a flow rate of 0.5 ml/min, the following gradient was applied: initial 2% solvent B (acetonitrile with 0.5% formic acid); 0.6 min, 6% B; 8 min, 20% B; 10 min, 40% B; 11 min, 95% B; 13 min, 95% B; 13.5 min, 2% B. Eluted substances were detected with a photodiode array detector (PDA 2996, Waters). Absorbance spectra were recorded at a frequency of 10 s⁻¹ between 210 and 600 nm, with a bandwidth of 1.2 nm, and chromatograms were extracted from the PDA data at 535 nm. The outlet of the PDA detector was coupled online with an LCT Premier Time-of-Flight (TOF) mass spectrometer (Waters) equipped with an electrospray ionization (ESI) and modular LockSpray interface. The capillary voltage was 2.2 kV and the source temperature was 100 °C. Spectra were recorded in positive-ion W-mode between mass and charge ratios of 100 and 1000. Mass calibration was performed in the range of 100–1000 m/z with phosphoric acid. Leucine enkephalin (2 ng/μl in 50% (v/v) aqueous acetonitrile) was employed during lockspray operation as internal mass reference for automated accurate mass measurement. Data were analyzed using Waters Mass Lynx software. Tentative annotations in the analytical chromatographic runs are based on detected protonated molecular mass, in source fragment masses relating to the anthocyanidin aglycones and the retention time order (Additional file 1: Figure S1, Additional file 2: Table S1). Isolation and mass spectrometric identification of the major anthocyanins from purple cob powder was realized as previously described [11].

For the glucose tolerance test (GTT), mice were fasted for 16 h, followed by an intraperitoneal glucose injection (2 g/kg body weight). Blood glucose was measured by tail bleeding using the One-Touch AccuChek Glucometer (Roche, Monza, Italy) at indicated times (0, 30, 60, 120 min).

For determination of non-esterified fatty acids (NEFA), plasma aliquots (50 μl) were added to a screw-capped vial sealed with a Teflon septum together with 12.5 µg of pentadecanoic and 12.5 µg of hexadecanoic acid as internal standards, butylated hydroxytoluene (BHT), K3-EDTA and 600 µl of methanol. After vigorous vortexing and centrifugation, the methanol phase was collected and evaporated under a gentle stream of nitrogen and converted into trimethylsilyl ethers with BSTFA with TCS 1% (Pierce).

Gas chromatography mass spectrometry (GC–MS) analysis was performed on an Elite column (30 m × 0.32 mm id × 0.25 mm film; Perkin Elmer, USA) and injection was performed in splitless mode and using helium (1 ml/min) as a carrier gas. The temperature program was as follows: initial temperature of 140 °C was held for 1 min, followed by a linear ramp of 10 °C/min to 290 °C, and thus held for 3 min. The mass spectrometer operates in full scan (from m/z 50 up to m/z 550) mode [17].

Peritoneal elicited macrophages (PEC) were derived from mice intraperitoneally (i.p.) injected with 1 ml thioglycollate (3% thioglycollate medium w/o dextrose; Becton–Dickinson, Franklin Lakes, NJ, USA). After 4 days, cells were collected by washing the peritoneal cavity with 5 ml of ice-cold sterile PBS, incubated in erythrocyte lysis buffer for 5 min, washed and cultured in a humidified incubator containing 5% CO₂ at 37 °C in 10% fetal calf serum (FCS)-RPMI 1640 medium for 24 h before treatment with extracts.

Epididymal white adipose tissue (WAT) and livers were minced into fine (< 10 mg) pieces, placed in HEPES buffered DMEM (Thermo Fisher, Walthan, MA, USA) supplemented with 10 mg/ml BSA (Sigma-Aldrich, Milano, Italy), and centrifuged at 1000×g for 10 min at room temperature (RT). An LPS-depleted collagenase Type II cocktail (Sigma-Aldrich) at a concentration of 0.03 mg/ml and 50 U/ml DNase I (Sigma-Aldrich) was added to the tissue suspension, and the samples were incubated at 37 °C on an orbital shaker (215 Hz) for 45 min. Samples were then passed through a sterile 100 µm nylon mesh (Corning, NY, USA) and centrifuged at 1000×g for 10 min. The pelleted cells from the WAT were collected as the SVF and the floating cells as the adipocyte-enriched fraction. Cells were resuspended in erythrocyte lysis buffer for 5 min and centrifuged at 500×g for 5 min before proceeding with the analysis.

Flow cytometry was performed using a BD FACSCantoII™ flow cytometer and analysis run with FACSDiva 6.1.1 software (Becton–Dickinson). Cells were stained with anti-mouse CD45-PB (#103125), CD11b-PE/Cy5 (#101209), F4/80-APC (#123115), F4/80-FITC (#123107), CD206-FITC (#141703), CD80-PE (#104707), SiglecF1-PE (#142403), Gr1-APC/Cy7 (#108423), CD3-APC (#100235), CD4-PE (#100407), CD8-PE/Cy5 (#100709) (BioLegend, San Diego, CA, USA) for 20 min at 4 °C, washed and resuspended in FACS Buffer (PBS, 5 mM EDTA and 0.2% BSA).

Intracellular staining for Ki67-AF488 (#561165, BD Biosciences) was performed by adding 3 ml cold 70% ethanol to the cell pellet and incubating at − 20 °C for 1 h, followed by staining with the antibody at RT for 30 min.

Once isolated, PEC or SVF from C57BL/6J were pre-treated with the extracts at a concentration of 125 μM of RED extract or cyanidin 3-glucoside (C3G) (Extrasynthese, Genay, France) for 24 h. The cells where then challenged with 100 ng/ml LPS for an additional 4 h for mRNA collection or 24 h for supernatant collection.

Total RNA was isolated from tissues and cells with Direct-Zol™ RNA MiniPrep kit (Zymo Research, Irvine, CA, USA). 1 μg RNA was reverse-transcribed with miScript II RT Kit (Qiagen, Hilden, Germany) and qRT-PCR performed with Fast Evagreen qPCR Master Mix (Bio-Rad, Segrate, Italy) in a CFX96 real-time PCR detection system (Bio-Rad). The expression of each specific mRNA was normalised against GAPDH and values were expressed as Fold Change on CTR + H₂O sample. Values represent mean ± SEM obtained from duplicate measurements performed on 4–6 biological replicates, as indicated in Figure legends (n = 4–6 mice/group). Primer sequences are summarized in Additional file 2: Table S2.

Samples were analysed for mouse IL-6, mouse IL-1β, and mouse TNF-α using Ready SET-Go! ELISA kit (Affymetrix, Santa Clara, CA, USA). The optical density (OD) of each well was analysed at 450 nm by a Microplate Reader (Tecan Infinite F200PRO).

Adipose tissues were lysed in a buffer containing 50 mM Tris–HCl, pH 7.2, 0.1% sodium deoxycholate, 1% Triton X-100, 5 mM EDTA, 5 mM EGTA, 150 mM NaCl, 40 mM NaF, 2.175 mM NaVO₄, 0.1% SDS, 0.1% aprotinin, and 1 mM PMSF. 30 μg of protein underwent SDS–PAGE following transfer on nitrocellulose membranes. Bands were detected using Pierce™ ECL Western Blotting Substrate (ThermoFisher Scientific). Antibodies against Anti-NF-kB p65 (Abcam, ab86299) and α-Tubulin (Sigma, T9026) were used. Densitometry quantification was performed with ImageJ Software (NIH) and expressed as ratio of specific protein to α-tubulin.

After fixing in 4% paraformaldehyde overnight, the fat and livers were dehydrated and embedded in paraffin. 5 μm sections were deparaffinised and incubated for 40 min in 2% HCl containing 10% potassium ferrocyanide, washed in 0.1 M phosphate buffer, followed by 0.025% 3,3′-DAB-4HCl (DAB; Sigma-Aldrich) and 0.005% H₂O₂ in a 0.1 M phosphate buffer for 40 min and/or counterstained with haematoxylin and eosin (H&E; Carl Roth, Karslruhe, Germany) and observed under a light microscope (Leica DM6000B, Wetzlar, Germany). For quantitative analysis of adipocyte area, five images per section were randomly acquired from each sample, and the cross-sectional area of each adipocyte was measured using ImageJ software (NIH).

C57BL/6J male mice were maintained on CTR + H₂O, HF + H₂O or HF + RED for 12 weeks. At the time of harvest, the weight of both HF groups was significantly greater than that of the CTR + H₂O (Fig. 1a). No differences were noticed between the two HF groups. Liquid intake was comparable among the three groups throughout the entire experiment (Additional file 1: Figure S2A). Food intake was slightly lower in the two HF groups compared with the control group (Additional file 1: Figure S2B), but it did not differ between the HF + H₂O and HF + RED groups. Energy intake was constant throughout the experiment and feed efficiency was greater in the two HF groups compared to CTR + H₂O (Additional file 2: Table S3). WAT weight was significantly greater in the two HF groups than in the control (2.8 ± 0.28 g, 7.5% of total body mass) and, although not statistically significant, greater in the HF + H₂O (6.117 ± 0.45 g, 13.2% of total body mass) compared to HF + RED (5.104 ± 0.28 g, 10.6% total body mass) (Fig. 1b). Histological examination of the WAT revealed a marked reduction in adipocyte cell size area in the HF + RED group (2735 ± 119 μm) compared to HF + H₂0 (4203 ± 183 μm) (Fig. 1c, d), suggesting adipocyte hypertrophy likely accounts for the increase in total adiposity. Blood glucose levels were recorded at time 0, 30, 60, and 120 min after injection of glucose to evaluate changes in glucose tolerance (Fig. 1e). In the HF + H₂O group the relative area under the curve (AUC) was increased by 36.8% (p = 0.0048), while the HF + RED group showed an increase of the 32.8% (p = 0.1981; Fig. 1f) relatively to the control group. Total and LDL/VLDL cholesterol levels were increased in both HF groups compared to CTR + H₂O and significantly reduced in HF + RED compared to HF + H₂O, whereas HDL cholesterol was significantly increased in HF + RED compared to both CTR + H₂O and HF + H₂O (Fig. 1g). No difference in total NEFA was observed in the three groups, despite a modest reduction in miristic, palmitic, linoleic, oleic and stearic acids in HF + RED compared to HF + H₂O (Additional file 2: Table S4). Serum ALT activity was significantly greater in the two HF groups than in CTR + H₂O, with a modest, but still not significant, reduction in HF + RED compared to HF + H₂O (Fig. 1h). Liver weight did not change among the three groups (data not shown), however HF + H₂O mice liver sections stained with H&E displayed abundant steatosis compared to the CTR + H₂O as well as HF + RED, indicating that the purple corn extract might inhibit fat accumulation in hepatocytes (Fig. 1i).

The SVF from epididymal WAT was stained with antibodies to assess leukocyte populations by flow cytometry. Macrophages were defined as F4/80⁺/CD11b⁺/Siglec-F⁻/Gr-1⁻ cells (Fig. 2a). Notably, CD3⁺ T cells accounted for approximately 24% of the CD45⁺ cells in both control and HF + RED groups, while set around 30% in the HF + H₂O group (Fig. 2b). Moreover, despite the overall prevalence of CD4⁺ T cells in all three groups (Fig. 2c), the CD4⁺ T cell fraction was larger in CTR + H₂O (52%) and HF + RED (50%) groups than in the HF + H₂O group (44%), whereas 25% of the CD3⁺ T cells in the HF + H₂O group were CD8⁺ , against the 20% in HF + RED and 16% in CTR + H₂O (Fig. 2d) groups, resulting in a slightly improved CD8/CD4 cell ratio of the HF + RED group compared to HF + H₂O group (Additional file 1: Figure S3).

Macrophage percentage was slightly higher in the HF + H₂O group (42%) compared to CTR + H₂O (35%) and HF + RED (34.4%) (Fig. 2e). Eosinophil content in the AT diminishes in obesity, as confirmed in both HF + H₂O (25.2%), and HF + RED (26%) groups, compared to CTR + H₂O (35%) (Fig. 2f). Neutrophil content in AT was higher in the HF + H₂O group compared to CTR + H₂O and HF + RED groups (Fig. 2g), although the difference was not significant.

F4/80 and CD11b were used to identify resident hepatic Kupffer cells (Fig. 2h–j). As shown in Fig. 2h, the Kupffer cell populations in the liver were not significantly different among the three groups, in agreement with Clementi et al. [18]. However, analysis revealed a reduction in CD80⁺ pro-inflammatory macrophages in the HF + RED group (Fig. 2i) compared to the HF + H₂O group, along with an increase in the number of CD206⁺ anti-inflammatory macrophages (Fig. 2j), suggesting that the RED extract might have the potential to modulate the liver’s contribution to the inflammatory state of obesity.

Obesity triggers the formation of crown-like structures (CLS) around individual adipocytes, typical of localized chronic inflammation [19]. We characterized the epididymal WAT histology in H&E sections. We observed a predominance of macrophages aggregated into ring patterned CLS in the HF + H₂O (6.417 ± 0.6566) group compared to the HF + RED (1.636 ± 0.2439) (Fig. 3a). No CLS were observed in the control group, where ATM were rather dispersed throughout the tissue. The decrease in HF + RED ATM was also confirmed by the reduced expression of the macrophage marker F4/80 in the epididymal WAT (Fig. 3b).

We therefore tested by flow cytometry whether ATM proliferation was reduced upon RED extract administration by staining SVF cells with the proliferation marker Ki67 and F4/80. Ki67 signal was detected in 66.94 ± 11.31% of ATM from the HF + H₂O group, against 41.50 ± 13.27% of HF + RED and only 1.240 ± 0.49% of CTR + H₂O (Fig. 3c), confirming a discrete reduction of the proliferation upon RED extract administration.

In order to characterize whether the decrease in ATM was due to a reduction in the monocyte chemoattractant protein-1 MCP-1 secretion by the tissue, we performed ELISA on epididymal WAT. As shown in Fig. 3d, the HF + RED group expresses significantly less MCP-1 (3076 ± 305.6 pg/ml) compared to the HF + H₂O group (5105 ± 404.2 pg/ml), which is consistent with a less inflammatory milieu.

We next assessed the expression of a panel of pro- and anti-inflammatory genes that represent markers of M1 and M2 macrophages respectively, to test the potential of the RED extract in promoting an ATM phenotypic switch in vivo. qRT-PCR analysis from ATM revealed that expression of all M1 inflammatory genes (TNF-α, COX-2, IL-1β, and IL-6) was dramatically downregulated in the ATM of the HF + RED group at levels comparable to those of the CTR + H₂O group (Fig. 3e–h). In addition, RED extract consumption was associated with an upregulation in the expression of all the alternative polarization markers (TGF-β, IL-10, ArgI, and FizzI) (Fig. 3i–l), confirming the results obtained in vitro of an anti-inflammatory effect of the RED extract on ATM due to its ability to induce an M2 phenotype switch.

Macrophages play a pivotal role in systemic iron recycling [20], contributing to adipogenesis and AT homeostasis. AT iron distribution was assessed by Perls’ Prussian blue method. Most of the iron depots within the AT were found coincidentally with CLS surrounding adipocytes in the HF + H₂O group, whereas lower iron staining was detected in HF + RED sections (Additional file 1: Figure S4A). This evidence was further confirmed by qRT-PCR analysis of the genes involved in iron metabolism. The HF + RED group displayed an increase in all iron metabolism genes (Additional file 1: Figure S4B–G), and a downregulation of the iron-storage gene FtL1 (Additional file 1: Figure S4H), consistent with an iron recycling phenotype.

We investigated whether our previous histological and molecular findings of the anti-inflammatory effect of RED extract on ATM also translates to a better adipocyte metabolic profile.

In adipocytes from epididymal AT, the most common adipokines (TNF-α, IL-6, IL-1β, resistin, RBP4, leptin, adiponectin, and perilipin), were all downregulated in the HF + RED condition compared to the HF + H₂O one (Fig. 4a–h). To investigate whether the inhibitory effect of the RED extract on pro-inflammatory genes and adipokine expression relates to transcription factor NF-κB, the nuclear levels of NF-κB p65 were determined in epididymal AT lysates by Western blot analysis. Despite the increased expression of α-tubulin observed in HF + RED condition, when normalized to α-tubulin, NF-κB p65 expression was significantly inhibited in the HF + RED sample compared to the HF + H₂O group after only 12 weeks of extract consumption (Fig. 4i, j), and resembled that of the untreated control group.

Given obesity is characterized by a low-grade inflammation, we investigated whether the RED extract consumption was able to exert a protective effect in vivo similar to the one observed in vitro (Additional file 1: Figure S5). SVF from the epididymal AT was exposed to 100 ng/ml LPS for 4 h. LPS-induced inflammatory cytokine expression was significantly suppressed by RED extract consumption (Fig. 5a–f). Next, we used ELISA assay to determine the production of TNF-α and IL-1β upon 24 h LPS. The data showed that cytokine production was significantly suppressed in the RED extract samples compared to both HF + H₂O and CTR + H₂O upon LPS treatment (Fig. 5g, h), confirming that the RED extract consumption indeed retains its protective effects also in vivo.