A total of 10 healthy volunteers (age, 18–40 years; 5 women and 5 men) were recruited for the study from the Department of Internal Medicine and Clinical Pharmacology (Medical University of Silesia, Silesia, Poland) all of whom were nonsmokers and were taking no medication. The ethical committee of the Medical University of Silesia accepted the study protocol. Peripheral blood mononuclear cells were extracted from the patients using Histopaque density gradient centrifugation (Sigma-Aldrich, St. Louis, MO, USA), using previously described methods (15). Subsequently, monocytes were isolated from the peripheral blood mononuclear cells via negative immunomagnetic separation, using Pan-T and Pan-B Dynabeads (Invitrogen Dynal AS, Oslo, Norway), according to the manufacturer's instructions. This procedure enabled the isolation of inactive monocytes without the use of artificial and uncontrolled stimulation. The isolated cells were labeled with a mouse anti-human monoclonal antibody (dilution, 1:100; F0844; Dako Denmark A/S, Glostrup, Denmark) against the monocyte-specific positive antigen CD14⁺. The procedure resulted in an isolated fraction containing 92% CD14⁺ cells. Monocytes were suspended in RPMI-1640 medium supplemented with 10% low endotoxin fetal calf serum, 2 mM glutamine, 100 U/ml penicillin, 100 mg/ml streptomycin and 10 mg/ml Gibco fungizone (Thermo Fisher Scientific, Inc., Grand Island, NY, USA). The cells were counted using a TC-20 automated cell counter (Bio-Rad Laboratories, Inc., Hercules, CA, USA). A constant number of cells (106 monocytes per well) were placed in a plastic 24-well plate (BD Biosciences, Franklin Lakes, NJ, USA) and left intact for 2 h for adherence. Then, the medium was changed and cultures were incubated for 72 h, with a single exchange of medium after the first 24 h. Incubations were performed in triplicate at 37°C in a humidified atmosphere containing 5% CO₂. The conversion of monocytes into macrophages was confirmed using polyclonal mouse antibodies against EMR1 (1:1,000; SAB1405756; Sigma-Aldrich). Following the 72-h incubation, the supernatant was carefully removed and replaced with medium supplemented with various combinations of metformin (0.02 or 2 mM; Sigma-Aldrich), compound C (20 µM; Sigma-Aldrich) or lipopolysaccharide (LPS; 1 µg/ml; Sigma-Aldrich) for 24 h. Certain cells were pre-incubated with metformin (0.02 or 2 mM) for 2 h to activate AMPK, then LPS (1 µg/ml) was added for 24 h. Certain cells were pre-incubated with compound C (20 µM) for 1 h to inhibit AMPK, then metformin (0.02 or 2 mM) was added. Following an additional 2 h, LPS was administered for 24 h. Compound C, at an initial concentration of 20 mM, was dissolved in dimethyl sulfoxide (DMSO; Sigma-Aldrich). A total of nine groups were thus produced, as follows: Control group (no treatment); LPS group; compound C group; 0.02 mM metformin group; 2.0 mM metformin group; compound C + 2.0 mM metformin group; LPS + 0.02 mM metformin group; LPS + 2.0 mM metformin group; and the LPS + compound C + 2.0 mM metformin group. Further dilutions were performed in the aforementioned RPMI medium and the corresponding quantities of DMSO were added to the control cultures. The final concentration of DMSO in the medium was ≤0.05% and, as previously confirmed, did not exert any effects on the cultured cells (16). Following the application of the various treatments, the cells were harvested for further analyses. Each group of experiments was performed in triplicate (17).

The cell viability was estimated using two tests. The first was based on a 0.4% trypan blue exclusion test (Sigma-Aldrich), according to manufacturers guidelines. Briefly, 10-µl aliquots of cultured cells suspended in RPMI medium with no additions were mixed with 10 µl 0.4% trypan blue. Following incubation for 3 min at 37°C, the cells were loaded onto a slide and the viability was assessed using the TC-20 automated cell counter. The second method involved MTT conversion (1). MTT was added to the medium (at a final concentration of 2.5 mg/ml) 3 h prior to the scheduled end of the experiment, then the cultures were incubated at 37°C in 5% CO₂/95% air. At the end of the experiment, after being washed twice with phosphate-buffered saline, monocytes were lysed in 100 µl DMSO at room temperature for 10 min in the dark, which enabled the release of the blue reaction product formazan. Then, 200 µl lysate was transferred to a 96-well plate (Falcon 353072; BD Biosciences). Absorbance was measured at 570 nm using a microplate reader (Dynex Technologies, Chantilly, VA, USA) using three measurements in each of ten independent experiments. The results are expressed as a percentage of the control.

The relative mRNA expression levels of GPx, CAT, SOD and p22phox were quantified using two-step RT-qPCR normalized against the expression level of glyceraldehyde 3-phosphate dehydrogenase (GAPDH) mRNA following 8 h of incubation. Total RNA was extracted from cells using TriPure Isolation Reagent (Roche Diagnostics, Basel, Switzerland) according to the manufacturer's instructions. The concentration and quality of the RNA extracts were estimated using spectrophotometry at 260 and 280 nm using a BioPhotometer (Eppendorf AG, Hamburg, Germany). Subsequently, 1 µg total RNA of each sample was reverse transcribed using an MMLV Reverse Transcriptase 1st-Strand cDNA Synthesis kit (Epicentre Technologies, Madison, WI, USA). A final RT reaction volume of 20 µl was diluted 5-fold in order to avoid possible reaction inhibition by high levels of cDNA. The qPCR was conducted using Brilliant II SYBR Green QRT-PCR 2-Step Master mix (Agilent Technologies, Inc., Santa Clara, CA, USA). Reaction mixtures consisted of 1X master mix, 300 nM of each (forward and reverse) primer complementary to one of the analyzed genes (GPx, CAT, SOD, p22phox or GAPDH), respectively, 4 µl cDNA mixture (i.e., an equivalent of 40 ng total RNA) in a total volume of 25 µl. The reaction mixtures were analyzed using a LightCycler 480 Real-Time PCR system (Roche Diagnostics) with a standard two-step thermal profile for qPCR: 95°C for 2 min, then 40 cycles of 95°C for 15 sec and 60°C for 30 sec. Following amplification, a melting curve was plotted for each sample in order to confirm the specificity of the reaction. Primers used for quantitative analysis were designed using eprimer3 software produced by European Molecular Biology Open Software Suite (EMBOSS; http://emboss.bioinformatics.nl/cgi-bin/emboss/eprimer3) according to the appropriate gene retrieved from the GenBank database (http://www.ncbi.nlm.nih.gov/genbank/) and were similar to those used in previous experiments: p22, forward 5′-TCCGGCCTGATCCTCATC-3′ and reverse 5′-AATGGAGTAGGCACCAAAGTACCA-3′; SOD, forward 5′-CTGATTTGGACAAGCAGCAA-3′ and reverse 5′-CTGGACAAACCTCAGCCCTA-3′; GPx, forward 5′-CGGGACTACACCCAGATGAA-3′ and reverse 5′-TCTCTTCGTTCTTGGCGTTC-3′; CAT, forward 5′-TCAGGCAGAAACTTTTCCATTT-3′ and reverse 5′-TGGGTCGAAGGCTATCTGTT-3′; and GAPDH, forward 5′-GAAGGTGAAGGTCGGAGTC-3′ and reverse 5′-GAAGATGGTGATGGGATTTC-3′) (18–20). Eprimer3, Primersearch and Water software were used for primer design and comparisons and are freely available on the server of the Pasteur Institute (Paris, France http://mobyle.pasteur.fr) as a part of EMBOSS (21). Additionally, randomly selected samples derived from qPCR assays were loaded onto 1.3% agarose gels (Agagel Mini; Biometra GmbH, Göttingen, Germany) in order to confirm the specificity of amplification. No other bands than expected for a particular primer pair were visible.

Total protein concentrations in samples were determined spectrophotometrically. Bovine serum albumin preparations (Fermentas, Glen Burnie, MD, USA) were used for calculation of the standard curve. Equal quantities of total protein (50 µg) mixed 1:1 with 2X sample buffer (25% glycerol, 2% sodium dodecyl sulfate and 0.02% bromophenol blue) were boiled for 6 min and loaded onto a 10% SDS-polyacrylamide gel (Sigma-Aldrich). Electrophoresis was performed at 180 V (constant voltage) until the bromophenol blue dye reached the end of the gel. Following electrophoresis, the stacking gels were removed and resolved, and sample gels were directly subjected to western blot analysis. After separation in polyacrylamide gels, the aliquots were transferred to polyvinylidene fluoride membranes (Pall Poland Ltd., Warszawa, Poland). Nonspecific antibody binding was inhibited by incubation in 20 mM Tris-buffered saline (pH 7.5) with 0.1% Tween 20 (TBST) containing 5% non-fat dried milk for 1 h at room temperature. Polyclonal antibodies against: p22phox, GPx, CAT and SOD were obtained from Sigma-Aldrich. The antibodies were diluted in TBST containing 5% skimmed milk. The membranes were incubated with the antibodies overnight at 4°C, washed with TBST, incubated at room temperature for 60 min with the appropriate goat anti-rabbit (1706518) and goat anti-mouse (1706520) alkaline phosphatase (AP)-conjugated secondary antibodies (1:1,000; Bio-Rad Laboratories Inc.) and washed twice with TBST for 5 min and once with 20 mM Tris-buffered saline (pH 7.8) for 5 min. In each assay, the colored precipitates were developed directly on the membrane using AP-chromogenic substrates (Bio-Rad Laboratories, Inc.). All membranes were photocopied and subjected to further analysis. The molecular weights of GPx, CAT, SOD and p22phox were confirmed according to their protein markers (PageRuler Unstained Protein Ladder; Fermentas). To control for the quantities of cytosolic proteins loaded in each lane, α-actin was detected in parallel using anti-α-actin antibodies (1:5,000; ab8227; Abcam, Cambridge, MA, USA). The integrated optical density (IOD) of signals was semi-quantified using Image-Pro Plus software, version 3.0 (Media Cybernetics, Inc., Rockville, MD, USA) and is expressed as the ratio of the IOD for the tested proteins to the IOD for α-actin. The experiment was repeated three times, and the relative density values were subjected to statistical analysis.

Results are expressed as the mean ± standard deviation. The normality of distribution was tested using Shapiro-Wilk's test. Statistical analysis of the data was performed using one-way analysis of variance followed by the post-hoc Tukey's significant difference test or Kruskal-Wallis test with Mann-Whitney tests according to the distribution of variables. The Bonferroni adjustment was applied for multiple comparisons. P<0.05 was considered to indicate a statistically significant difference. Statistical analysis was performed using a SPSS software, version 16.0 (SPSS, Inc., Chicago, IL, USA).

No significant differences in cell viability were observed among the cultures regardless of the reagents with which they were treated (Fig. 1).

LPS significantly elevated the expression of p22phox at the mRNA (6.34±0.56 vs. 1.0±0.21-fold; P<0.001) and protein (7.23±1.45 vs. 1.0±0.26-fold; P<0.001) levels compared with that in the control group (Fig. 2). Metformin at a low concentration (0.02 µM) did not significantly alter the mRNA expression levels of p22phox (1.12±0.23 vs. 1.0±0.21-fold; P=0.217); however, a high concentration of metformin (2.0 µM) resulted in the elevation of mRNA expression levels (1.43±0.15 vs. 1.0±0.26-fold; P=0.041) compared with those in the control. The protein expression levels of p22phox were not significantly affected by 0.02 or 2.0 mM metformin. In the LPS-treated macrophages, 0.02 mM metformin did not significantly alter the mRNA or protein expression levels of p22phox. By contrast, 2.0 mM metformin induced a significant reduction in the mRNA (4.56±0.41 vs. 6.34±0.56; P=0.007) and protein (5.33±0.94 vs. 7.23±1.45; P=0.002) expression levels of p22phox in the LPS-treated macrophages. The addition of compound C significantly inhibited the effect of metformin on the mRNA (5.95±0.39 vs. 4.56±0.51; P=0.011) and protein (7.03±1.23 vs. 5.33±0.94; P=0.009) expression levels of p22phox in the LPS-treated cells. Following treatment with compound C, these levels were comparable to those of the cells treated with LPS alone.

LPS caused a small increase in the mRNA expression level of SOD (1.32±0.19 vs. 1.0±0.13; P=0.038) compared with that in the control cells; however, it did not significantly change SOD protein expression (1.12±0.27 vs. 1.0±0.23) (Fig. 3). Metformin at 0.02 µM significantly increased the mRNA expression level of SOD (1.4±0.21 vs. 1.0±0.13; P=0.027), while 2.0 mM metformin significantly elevated the mRNA (1.8±0.18 vs. 1.0±0.13; P=0.006) and protein (1.95±0.35 vs. 1.0±0.23; P=0.007) expression levels of SOD compared with those in the control group. The addition of 0.02 mM metformin to LPS-treated cells resulted in a moderate increase in SOD mRNA expression (1.65±0.24 vs. 1.32±0.19; P=0.009) compared with that in the cells treated with LPS alone. Metformin at 2.0 µM produced a more marked difference in SOD mRNA expression (2.1±0.21 vs. 1.32±0.19; P=0.005) in the LPS-treated cells. The effect of metformin on SOD protein concentration was particularly marked in the cells treated with 0.02 µM (3.5±0.54 vs. 1.12±0.27; P<0.001) and 2.0 mM metformin (4.3±0.44 vs. 1.12±0.27; P<0.001), indicating that metformin increases the expression of SOD more effectively in an inflammatory environment. Compound C reduced the effect of 2.0 mM metformin on SOD mRNA (1.25±0.28 vs. 2.1±0.21; P=0.006) and protein (1.4±0.23 vs. 4.3±0.44; P<0.001) expression levels, which resulted in mRNA expression levels similar to those of the LPS-treated macrophages (1.25±0.28 vs. 1.32±0.19; P=0.425); however, the protein expression levels of SOD remained slightly elevated (1.4±0.23 vs. 1.12±0.27; P=0.048).

Treatment with LPS increased the expression levels of GPx mRNA (1.23±0.13 vs. 1.0±0.24; P=0.041) and protein (1.84±0.21 vs. 1.0±0.13; P=0.002) compared with those in the control group (Fig. 4). In contrast to SOD, 0.02 mM metformin did not significantly alter the mRNA or protein expression levels of GPx compared with those in the control group. However, 2.0 mM metformin, similarly to its effect on SOD expression, increased the mRNA (1.76±0.14 vs. 1.0±0.24; P=0.011) and protein (1.72±0.43 vs. 1.0±0.13; P=0.016) expression levels of GPx compared with the control. The addition of 0.02 mM metformin to the LPS-treated cells resulted in a significant increase in the mRNA (1.43±0.17 vs. 1.23±0.13; P=0.038) and protein (2.13±0.34 vs. 1.84±0.21; P=0.046) expression levels of GPx. Furthermore, 2.0 mM metformin produced a more marked increase in the mRNA (2.14±0.33 vs. 1.23±0.13; P=0.001) and protein (2.56±0.22 vs. 1.84±0.21; P=0.003) expression levels of GPx in the LPS-treated cells. Experiments involving compound C and LPS indicated that metformin expressed its effects via AMPK, as a significant inhibitory influence of compound C on the expression of GPx mRNA and protein was observed. Following the AMPK inhibition, the GPx expression was comparable to that in samples treated only with LPS (1.33±0.2 vs. 1.23±0.13; P=0.289 for mRNA) and (1.66±0.28 vs. 1.84±0.21; P=0.361 for protein).

LPS resulted in a moderate but statistically significant increase in CAT mRNA (1.3±0.24 vs. 1.0±0.24; P=0.042) and protein (1.23±0.15 vs. 1.0±0.21; P=0.029) expression levels in cultured macrophages. In 2.0 mM metformin-treated samples a mild elevation in CAT mRNA expression was observed (1.35±0.22 vs. 1.0±0.24; P=0.027); however, 0.02 mM metformin did not induce a significant effect on CAT mRNA. Notably, metformin elevated CAT protein synthesis by a similar degree in the 0.02 µM (1.21±0.15 vs. 1.0±0.21; P=0.049) and 2.0 µM (1.26±0.15 vs. 1.0±0.21; P=0.037) groups of macrophages. In LPS-treated macrophages, the CAT mRNA level was significantly increased by treatment with 0.02 µM (1.54±0.18 vs. 1.3±0.24; P=0.023) and 2.0 µM (1.76±0.25 vs. 1.3±0.24; P=0.003) metformin. However, similar changes in protein expression were not observed. Compound C inhibited the influence of 2.0 mM metformin on CAT mRNA expression in LPS-treated macrophages (1.32±0.21 vs. 1.76±0.25; P=0.005), without a significant effect on CAT protein expression (Fig. 5).