U-937 cells (purchased from American Type Culture Collection, Manassas, VA, USA and preserved in our laboratory) were cultured and maintained at a cell density between 5×10⁵ and 1×10⁶ cells/ml in RPMI-1640 media (Gibco; Thermo Fisher Scientific, Inc., Villebon-sur-Yvette, France) supplemented with 10% fetal bovine serum (Biowest, Nuaillé, France), 100 U/ml penicillin (Genom Co., Ltd., Hangzhou, China), and 100 μg/ml streptomycin (Genom Co., Ltd.). The cells were incubated at 37°C in a humidified atmosphere containing 95% air and 5% CO₂. For macrophage-like cell differentiation, the serum-starved U-937 cells were plated in 6-well plates (Corning, Inc., Corning, NY, USA) at a density of 5×10⁵ cells/ml and treated with 60 ng/ml PMA (Sigma-Aldrich, St. Louis, MO, USA) for 24 h. The non-adherent cells were removed by washing twice with phosphate-buffered saline (PBS).

The macrophages were divided into three groups: The control group, the LPS-treated group and the TSA-treated group. For the TSA-treated group, TSA (Sigma-Aldrich) was added to the medium 2 h prior to treatment with LPS (1 μg/ml; Sigma-Aldrich). The final concentrations of TSA were 50, 100, 500 and 1,000 nM [TSA was dissolved in dimethyl sulfoxide (DMSO; Sigma-Aldrich), and the final DMSO concentration was <0.4%]. For the LPS-treated group, an equal dosage of DMSO was added to the medium 2 h prior to treatment with LPS (1 μg/ml). For the control group, the same doses of DMSO and normal saline were added to the medium at the same time point. The super-natants and cells were harvested 24 h after the addition of LPS.

Cell viability was assessed using the Cell Counting kit-8 (CCK-8; Dojindo Molecular Technologies, Inc., Kumamoto, Japan). The cells (2×10⁵ cells/well) were plated in 96-well plates for the viability assay. Experimental procedures were performed according to the manufacturer's protocol.

The apoptotic rate of the macrophages was measured by flow cytometry (Cell Lab Quanta SC; Beckman Coulter, Inc., Brea, CA, USA) using an Annexin V-Fluorescein Isothiocyanate (FITC) Apoptosis Detection kit (Lianke Biotech Co., Ltd., Hangzhou, China). Briefly, non-adherent and adherent cells were collected. The cells were suspended in 500 μl binding buffer and stained with Annexin V-FITC and propidium iodide (PI) at room temperature for 5 min in the dark. After removing the unbound Annexin V-FITC and PI by centrifugation at 2,000 × g at 4°C for 5 min, the cells were resuspended in excess binding buffer. For each assay, ≥10,000 cells were analyzed by FACS. Data were expressed as the percentage of non-viable (Annexin V⁺+PI⁺) cells.

The levels of tumor necrosis factor (TNF)-α, interferon (IFN)-γ, interleukin (IL)-10 and IL-18 in the cell supernatants were measured using enzyme-linked immunosorbent assay (ELISA) kits (eBioscience, Inc., San Diego, CA, USA), according to the manufacturer's protocols.

Total RNA was extracted from the isolated macrophages using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc., Waltham, MA, USA), according to the manufacturer's protocol. RNA samples were quantified by measuring the absorbance (A) at 260 and 280 nm using a spectrophotometer (UV762; Shanghai Analysis Instrument Co., Ltd., Shanghai, China). The concentrations of RNA were calculated according to A260. Aliquots of total RNA (1 μg) from each sample were reverse transcribed into cDNA, according to the instructions of the First Strand cDNA Synthesis kit (Takara Bio, Inc., Otsu, Japan).PCR amplification was performed in a volume of 20 μl containing 2 μl cDNA, 0.25 μmol/l of each primer, 0.25 mmol/l deoxyribonucleotide triphosphates, and 1 U Takara TaqHS polymerase (Takara Bio, Inc.) using SYBR Green (Takara Bio, Inc.) as the fluorescence indicator on a Bio-Rad Cycler system (Bio-Rad Laboratories, Inc., Hercules, CA, USA). In the PCR amplification reaction, the thermocycling conditions were as follows: Initial activation at 95°C for 15 min, followed by 40 cycles of denaturation at 94°C for 15 sec, annealing for 30 sec at 55°C and extension for 30 sec at 72°C. The primers used were synthesized by Sangon Biotech Co., Ltd. (Shanghai, China), and the sequences were as follows. HDAC1, forward TAA TAA GCA GCA GAC GGA CATCG, reverse CAT AAT ACT TGC CTT TGC CAGC; HDAC2, forward TGA TGG AGA TGT ATC AAC CTA GTGC, reverse TTG AAA CAA CCC AGT CTA TCA CCAG; HDAC3, forward TGG TTA TAC TGT CCG AAA TGT TGC, reverse CTG TCA TAG GTC AGG AGG TCTGC; cyclooxygenase (COX)-2, forward AAA TCC TTG CTG TTC CCACC, reverse TTT CTC CAT AGA ATC CTG TCCG; chemokine (C-X-C motif) ligand 2 (CXCL-2), forward AAA GTG TGA AGG TGA AGT CCCCC, reverse GTG ATG CTC AAA CAC ATT AGGCG; chemokine (C-C motif) ligand 7 (CCL-7), forward ACA GAA GGA CCA CCA GTA GCCAC, reverse GCT TTG GAG TTT GGG TTT TCT TGT; and GAPDH, forward CCACATCGCTCAGACACCAT, reverse CCAGGCGCCCAATACG. The quantification cycle (Cq) value was analyzed from the amplification plots, and gene expression was normalized against the Ct of the GAPDH housekeeping gene using the 2^−ΔΔCt method (20).

The U-937 cells were washed twice with cold PBS, and lysed with lysis buffer (Beyotime Institute of Biotechnology, Beijing, China) on ice for 1 h. Following centrifugation at 12,000 × g at 4°C for 5 min, the supernatants were collected. The concentrations of protein were measured with a bicinchoninic acid protein assay kit (Beyotime Institute of Biotechnology). Briefly, 50 μg protein extracts were separated by 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred onto a polyvinylidene fluoride membrane (EMD Millipore, Billerica, MA, USA). The membranes were blocked with 5% skim milk for 1 h and then incubated with the following polyclonal primary antibodies at 4°C overnight: Anti-HDAC1 (cat. no. 2062; 1:1,000; Cell Signaling Technology, Inc., Danvers, MA, USA), anti-HDAC2 (cat. no. 2540; 1:1,000; Cell Signaling Technology, Inc.), anti-HADC3 (cat. no. 2632; 1:1,000; Cell Signaling Technology, Inc.), anti-H2A (cat. no. 2578; 1:1,000; Cell Signaling Technology, Inc.), anti-AH2A (cat. no. 2576; 1:1,000; Cell Signaling Technology, Inc.), anti-H2B (cat. no. 2934; 1:1,000; Cell Signaling Technology, Inc.), anti-AH2B (cat. no. 2571; 1:1,000; Cell Signaling Technology, Inc.), anti-H3 (cat. no. sc-8654; 1:500; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), anti-AH3 (cat. no. 8172; 1:1,000; Cell Signaling Technology, Inc.), anti-H4 (cat. no. 2935; 1:1,000; Cell Signaling Technology, Inc.), anti-AH4 (cat. no. 2591; 1:1,000; Cell Signaling Technology, Inc.), anti-p65 (cat. no. 6956; 1:1,000; Cell Signaling Technology, Inc.), anti-phosphorylated-p65 (cat. no. 3031; 1:1,000; Cell Signaling Technology, Inc.), anti-Ac-p65 (cat. no. 3045; 1:1,000; Cell signaling Technology, Inc.), anti-TLR4 (cat. no. sc-10741; 1:500; Santa Cruz Biotechnology, Inc.) and anti-β-actin (cat. no. 4967; 1:1,000; Cell Signaling Technology, Inc.). This was followed by incubation with a fluorescent secondary antibody (LI-COR Biosciences, Lincoln, NE, USA) at 37°C for 2 h. The blot was analyzed using the Odyssey Infrared Imaging system (LI-COR Biosciences). Membranes were also probed for β-actin (Cell Signaling Technology, Inc.) and histone 3 (Santa Cruz Biotechnology, Inc., Dallas, TX, USA) as additional loading controls.

Statistical analysis was performed using SPSS version 17.0 software for Windows (SPSS, Inc., Chicago, IL, USA). Data are presented as the mean ± standard deviation, and comparisons were made using Student's t-test. P<0.05 was considered to indicate a statistically significant difference.

In order to determine a suitable treatment dosage of TSA for the present study, the cytotoxic effects of TSA on the PMA-induced macrophages were detected by a CCK-8 assay. As shown in Fig. 1, the viability of macrophages decreased in a dose-dependent manner when exposed to various doses of TSA (50, 100, 500 and 1,000 nM). The cell viability was 93% when treated with 50 nM TSA, whereas the viability decreased to 42% when the cells were treated with 1,000 nM TSA. Cell viability was >80% when the dose of TSA was below 500 nM; therefore, the present study used 50, 100 and 500 nM TSA to explore the effects of TSA on macrophages.

The expression levels of TNF-α, IFN-γ, IL-10 and IL-18 were measured in the cell supernatants by ELISA. As shown in Fig. 2, TNF-α, IFN-γ, IL-10 and IL-18 expression levels were significantly increased in the LPS-treated group, as compared with the control group (P<0.05). The expression levels of TNF-α, IFN-γ, IL-10 and IL-18 were markedly decreased in the TSA-treated group, as compared with the LPS-treated group (P<0.05), in a dose-dependent manner.

To detect changes in the expression of the substrates of TSA, the mRNA and protein expression levels of class I HDACs (HDAC1, HDAC2 and HDAC3) were detected by RT-PCR and western blotting, respectively. As shown in Fig. 3, the expression levels of HDAC1, HDAC2 and HDAC3 were markedly increased in the LPS-treated group, as compared with the normal group; however, the expression levels were all decreased in the TSA-treated group. These results suggest that the expression of class I HDACs may be enhanced in the process of inflammation, and may be decreased by TSA alongside the inhibition of inflammation.

As shown in Fig. 4, as compared with the PMA-induced macrophages in normal conditions (Fig. 4A), a low dose of TSA (50 nM) had little effect on the apoptosis of macrophages (Fig. 4B). However, there was obvious apoptosis in the macrophages that were treated with higher doses of TSA (100 or 500 nM, respectively) (Fig. 4C and D). When LPS was added (Fig. 4E–H), as compared with the normal condition, the survival rate of the macrophages was significantly decreased (79.1 vs. 92.4%) (Fig. 4E). However, when treated with 50 nM TSA, the survival rate of the macrophages was increased to 88.2% (Fig. 4F). In addition, when treated with higher doses of TSA (100 or 500 nM respectively), the survival rate of the macrophages decreased to 67.3 and 52.4%, respectively (Fig. 4G and H). These results suggest that the inhibitory effects of low dose TSA (50 nM) on the release of inflammatory cytokines are not due to TSA-induced apoptosis of macrophages.

The present study detected the mRNA expression levels of COX-2, CXCL-2 and CCL-7, in order to explore whether LPS-induced gene expression could be suppressed by a low dose of TSA (50 nM). As shown in Fig. 5, the mRNA expression levels of COX-2, CXCL-2 and CCL-7 were significantly increased in the LPS-treated group, as compared with the normal group (P<0.05). In addition, COX-2 and CXCL-2 mRNA expression levels were significantly increased in the TSA-treated group, whereas CCL-7 mRNA expression levels were decreased, as compared with the LPS-treated group (P<0.05). These results suggest that TSA may differentially regulate the expression levels of specific proinflammatory genes in macrophages.

To investigate whether the inhibitory effects of TSA on inflammation were due to the acetylation of histones, the acetylation levels of histones was determined. As shown in Fig. 6, the acetylation levels of histone (H)2A, H2B, H3 and H4 were significantly increased in the LPS-treated group, as compared with the normal group (P<0.05). Furthermore, the acetylation levels of H2A, H2B, H3 and H4 were all significantly increased in the TSA-treated group (P<0.05). These results suggest that the inhibitory effects of TSA on inflammation may be caused by enhancement of histone acetylation, in order to selectively suppress the expression of proinflammatory genes.

The present study determined whether the inhibitory effects of TSA on inflammation were via regulation of the TLR4/NF-κB pathway. As shown in Fig. 7, the protein levels of TLR4 in the LPS-treated group were significantly higher, as compared with the normal group (P<0.05), whereas they were decreased in the TSA-treated group. In addition, there were alterations in the acetylation and phosphorylation of NF-κB p65. The acetylation of NF-κB p65 was significantly decreased in the LPS-treated group, as compared with the normal group, but was significantly increased in the TSA-treated group (P<0.05). Conversely, the phosphorylation of NF-κB p65 was increased in the LPS-treated group, but was decreased in the TSA-treated group (P<0.05). These results suggest that TSA may affect the TLR4/NF-κB p65 signaling pathway by regulating the balance of acetylation and phosphorylation of NF-κB p65. Therefore, the TSA-induced inhibition of inflammation may be due to regulation of the acetylation of non-histone proteins.