Flavin-containing enzymes as a source of reactive oxygen species in HEMA-induced apoptosis

Graphical abstract

Theoretical model of the formation of oxidative stress in cells exposed to dental resin monomers.

Abstract

Objective

Oxidative stress induced by compounds of dental composites like 2-hydroxyethyl methacrylate (HEMA) due to excess formation of reactive oxygen species (ROS) disturbs vital cell functions leading to apoptosis. The sources of ROS in cells exposed to resin monomers are unknown. The present study investigates functions of flavin-containing ROS and RNS (reactive nitrogen species) producing enzymes in cells exposed to HEMA.

Methods

The formation of oxidative stress in RAW264.7 mouse macrophages exposed to HEMA (0–6–8 mM) was determined by flow cytometry (FACS) after staining of cells with 2′7′-dichlorodihydrofluorescin diacetate (H 2 DCF-DA), dihydroethidium (DHE) or dihydrorhodamine 123 (DHR123). Cells in apoptosis or necrosis were identified by annexin-V-FITC/propidium iodide labeling followed by FACS analysis. Expression of ROS/RNS producing enzymes was analyzed by Western blotting.

Results

DCF fluorescence increased in cells exposed to HEMA for 1 h suggesting the production of hydroxyl radicals, H 2 O 2 , or nitric oxide and superoxide anions which form peroxynitrite (ONOO-). Increased DHR123 fluorescence after 24 h indicated the formation of mostly H 2 O 2 . The induction of apoptosis in the presence of HEMA was decreased by low concentrations of diphenylene iodonium (DPI), an inhibitor of flavin-containing enzymes. Expression of p47 phox , a regulatory subunit of the superoxide producing Nox2, was downregulated, and the expression of NOS which produces nitric oxide (NO) was possibly inhibited by feedback loop mechanisms in HEMA-exposed cultures. Inhibition of HEMA-induced apoptosis by VAS2870 or apocynin further suggested a crucial function of Nox2.

Significance

The present findings show the physiological relevance of flavin-containing enzymes in monomer-induced oxidative stress and apoptosis.

Introduction

Clinically used dental composites release resin monomers as a result of incomplete polymerization processes. Particularly hydrophilic monomers like 2-hydroxyethyl methacrylate (HEMA) are bioactive, and thus capable of interfering with tissues of the oral mucosa or the dental pulp after diffusion through dentin . Depending on the remaining dentin thickness, resin monomers like HEMA may be available at biologically relevant concentrations sufficient to influence crucial functions of pulp tissues in vivo . These compounds, which are used to direct the course of dental therapy, can cause specific stress responses in cell cultures derived from various oral target tissues. Physiologically relevant levels of resin monomers interfere with vital odontoblast cell functions in vitro such as matrix mineralizing capability or expression of gene products essential for tertiary dentin formation . Monomers may also delay essential functions of targeted cells of the innate immune system like the release of cytokines stimulated by lipopolysaccharide (LPS) released from cariogenic Gram-negative microorganisms . Moreover, it is firmly established that resin monomers trigger programmed cell death or apoptosis through the intrinsic pathway as a consequence of oxidative DNA damage as a physiological process . These biological effects observed in a variety of eukaryotic cells exposed to dental resin monomers are related to the formation of reactive oxygen species (ROS) beyond the capacities of the cellular antioxidant system causing the phenomenon of oxidative stress .

Cellular redox homeostasis is maintained as a strictly controlled balance between the formation of pro-oxidants such as ROS and the activity of non-enzymatic and enzymatic antioxidants . Recently, it has been detected that the resin monomer HEMA induced the activation of a protective network of pathways under the control of the redox-sensitive transcription factor Nrf2 (nuclear factor erythroid 2 [NF-E2]-related factor 2), a major transcriptional activator of numerous genes coding for enzymatic antioxidants. Activation of Nrf2 was identified as a fundamental mechanism that protected cells from monomer-induced apoptosis in particular, although it is certainly involved in other phenomena related to the control of excessive pro-oxidants as well . Yet, the specific nature of pro-oxidants generated in cells exposed to resin monomers like HEMA is still unknown, and the identification of their cellular origin is essential. Insight into these mechanisms will help establish effective strategies for the development of specific means to control adaptive cellular responses associated with oxidative stress in oral tissues exposed to monomers in a clinical situation.

Pro-oxidants encompass ROS as well as RNS (reactive nitrogen species) including radicals and non-radicals . ROS such as superoxide anions, hydroxyl or peroxyl radicals may function as oxidizing agents per se , or convert into molecules like hypochlorous acid (HOCl) or hydrogen peroxide (H 2 O 2 ). On the other hand, nitric oxide (NO) is a reactive nitrogen species and a radical generated by nitric oxide synthases (NOS). Finally, superoxide anions and NO combine to peroxynitrite (ONOO−) to become the basis of a cascade of reactions yielding numerous derivatives like NO 2 , NO 3, or N 2 O 3 . The formation of ONOO− is most relevant under physiological conditions since this molecule is extremely potent in the oxidation of cellular macromolecules .

Considering the vast number of bioactive derivatives with distinct physiological functions, the identification of the sources of ROS, and possibly RNS, which generate oxidative stress in monomer-exposed cells is extremely challenging and complex. So far, the analysis of resin monomer-induced oxidative stress has focused primarily on the use of DCFH 2 -DA (2′,7′-dichlorfluorescein-diacetate) as a redox-sensitive fluorescent dye . Dihydroethidium (DHE) was used to specifically indicate the formation of superoxide anions due to its oxidation into 2-hydroxyethidium, and dihydrorhodamine 123 (DHR123) allowed for the analysis of hydrogen peroxide (H 2 O 2 ) formation .

We assume that a plethora of ROS and RNS possibly generated in monomer-exposed cells might originate from the activity of flavin-containing enzymes such as NADPH oxidases (Nox), xanthine oxidoreductase (XOR), or nitric oxide synthases (NOS). NADPH oxidases as flavoenzymes transport electrons across cell membranes and reduce molecular oxygen to generate superoxide anions. Nox enzyme activities have been described as essential for cell proliferation, signaling, apoptosis or cell response to environmental stresses caused, for instance, by pathogenic microorganisms . Xanthine oxidoreductase as a molybdoflavinenzyme catalyzes purine degradation thereby reducing molecular oxygen to superoxide. The enzyme exists in two forms, xanthine reductase (XR) and xanthine oxidase (XO), depending on the cellular redox status . Finally, nitric oxide (NO) as a signaling molecule is produced by three isoforms of nitric oxide synthases identified so far. The inducible NOS (iNOS) is expressed in macrophages after physiological activation through pro-inflammatory stimuli .

We hypothesize that pharmacological inhibition of these various flavoenzymes in cells exposed to the resin monomer HEMA might be a valuable strategy for gaining insight into the multitude of ROS or RNS generated. Moreover, both the levels and quality of ROS and RNS could vary depending on exposure periods. Since the induction of apoptosis by a resin monomer has been causally related to the formation of oxidative stress , we also presumed that the inhibition of Nox, XOR, or NOS activities would indicate the origin of ROS or RNS responsible for monomer-induced apoptosis. Finally, the analyses of the expression of Nox, XOR, or NOS, and proteins directly related to the regulation of redox homeostasis under the control of Nrf2, would add even more evidence to the identification of the intracellular sources of monomer-induced oxidative stress. For this purpose, we used RAW264.7 mouse macrophages as a suitable model cell line of the innate immune system and we analyzed HEMA concentrations which induce concentration-dependent effects in these cells as repeatedly established in related recent projects . Conclusively, the findings of these analyses will encourage the creation of novel approaches for dental treatment strategies to protect human pulp tissues from oxidative stress and support a beneficial environment for tissue repair .

Materials and methods

Chemicals and reagents

2-Hydroxyethyl methacrylate (HEMA; CAS-No. 868-779) was purchased from Merck (Darmstadt, Germany). RPMI 1640 medium containing l -glutamine and 2.0 g/l NaHCO 3 came from PAN Biotech (Aidenbach, Germany). Fetal bovine serum (FBS), penicillin/streptomycin, and phosphate-buffered saline supplemented with 5 mM EDTA (PBS-EDTA) were obtained from Life Technologies, Gibco BRL (Eggenstein, Germany). Dihydroethidium (DHE; CAS-No. 104821-25-2), dihydrorhodamine 123 (DHR123; CAS-No. 109244-58-8), phorbol myristate acetate (PMA; CAS-No. 16561-29-8), oxypurinol (CAS 2465-59-0), VAS2870 (CAS 722456-31-7), and a bicinchoninic acid protein assay kit were purchased from Sigma (Taufkirchen, Germany). 2′7′-Dichlorodihydrofluorescin diacetate (DCFH 2 -DA; CAS-No. 4091-99-0) came from MoBiTec (Göttingen, Germany). An annexin V-FITC apoptosis detection kit was obtained from R&D Systems (Minneapolis, MN, USA). Diphenylene iodonium chloride (CAS 4673-26-1) was acquired from Tocris Bioscience (R&D Systems GmbH, Wiesbaden-Nordenstadt, Germany), L-NG-nitroarginine methyl ester (L-NAME, CAS 51298-62-5) and apocynin (acetovanillone, CAS 498-02-2) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA).

Anti-catalase (H-300, sc-50508), anti-heme oxygenase 1 (HO-1, sc-1797), anti-Nrf2 (H-300, sc-13032), anti-p67-phox (sc-7663) polyclonal antibodies plus anti-Cu–Zn superoxide dismutase (SOD1, sc-271014), anti-glutathione peroxidase 1/2 (GPx1/2, sc-133152), anti-p47-phox (sc-17845), and anti-xanthine oxidoreductase monoclonal antibodies (XOR, sc-398548) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-peroxiredoxin 1 (Prdx1; no. 8732), anti-nitric oxide synthase (NOS) polyclonal antibodies and anti-rabbit IgG HRP-linked antibodies (no. 7074) were purchased from Cell Signaling (NEB Frankfurt, Germany). Goat anti-mouse IgG (H + L)-HRP conjugate came from Bio-Rad Laboratories (Munich, Germany), and Amersham hyperfilm ECL was obtained from GE Healthcare (Munich, Germany). A protease inhibitor cocktail was acquired from Roche Diagnostics (Mannheim, Germany), and an anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) monoclonal antibody (6C5) came from Millipore (Schwalbach, Germany). All chemical reagents were of analytical grade.

Cell culture

RAW264.7 mouse macrophages (ATCC TIB71) were grown in RPMI 1640 medium containing glutamine, sodium-pyruvate and 2.0 g/l NaHCO 3 supplemented with 10% fetal bovine serum (FBS) and penicillin-streptomycin at 37 °C and 5% CO 2 as described in Ref. .

Formation of reactive oxygen species (ROS)

The intracellular production of ROS in RAW264.7 mouse macrophages was detected using the oxidation-sensitive probe 2′7′-dichlorodihydrofluorescin diacetate (DCFH 2 -DA), and dihydroethidium (DHE) or dihydrorhodamine 123 (DHR123) specifically indicated the formation of superoxide anions or hydrogen peroxide, respectively . Cell cultures in 6-well plates (5 × 10 4 /well) were stimulated with HEMA (0–6–8 mM) or phorbol myristate acetate (0.0–0.1–1.0 μM PMA) for 1 h or 24 h at 37 °C as described in Ref. . Cells were exposed to PMA as a positive control to show the induction of ROS .

For further analysis, cells were preincubated with diphenylene iodonium (DPI) (0–10 μM) or L-NAME (0–300 μM) for 1 h. After that, cells were exposed to HEMA (0–6–8 mM) or PMA (0.0–0.1–1.0 μM) in the presence or absence of various concentrations of DPI (0–10 μM) or L-NAME (0–300 μM) for 1 h or 24 h. The concentrations of DPI (0–10 μM) or L-NAME (0–300 μM) used here were selected after preliminary range-finding experiments. ROS formation in HEMA-exposed cell cultures was also analyzed using oxypurinol, VAS2870, apocynin, diapocynin or plumbagin but no influence of these inhibitors was detected (unpublished observations).

The formation of ROS was determined by flow cytometry (FACSCanto, Becton Dickinson) as described in Ref. . Briefly, the cells were incubated with 10 μM DCFH 2 -DA, 5 μM DHE or 5 μM DHR123 in complete medium 30 min before harvesting in PBS/5 mM EDTA. Then, cells were routinely collected by centrifugation and resuspended in 200 μl CMF–PBS. DCF fluorescence was measured at an excitation wavelength of 488 nm and an emission wavelength of 519 nm (Fl-1). DHE and DHR123 fluorescence was analyzed at an excitation wavelength of 488 and an emission wavelength of 530 nm (Fl-1, DHE) or 578 nm (Fl-2, DHR123). Mean fluorescence intensities were obtained by histogram statistics (FACSDiva™ 5.0.2 software), and individual values of fluorescence intensities in treated cell cultures were normalized to fluorescence measured in untreated controls (= 1.0) .

Detection of apoptosis and necrosis by flow cytometry

RAW264.7 mouse macrophages (1 × 10 5 cells/well) grown in 6-well plates were preincubated with diphenylene iodonium (DPI) (0–10 μM), apocynin (0–500 μM), L-NAME (0–300 μM), oxypurinol (0–100 μM), or VAS2870 (0–30 μM) for 1 h. Next, preincubation media were removed and cells were stimulated with HEMA (0–6–8 mM) in the presence or absence of DPI (0–10 μM), apocynin (0–500 μM), L-NAME (0–300 μM), oxypurinol (0–100 μM), or VAS2870 (0–30 μM) for 24 h following published procedures . Effective concentrations of the inhibitors were selected both in range finding experiments with HEMA and from their influence observed on PMA-induced apoptosis (unpublished observations).

Apoptosis or necrosis in exposed cells and untreated controls was determined as described elsewhere . Briefly, cell cultures were washed with phosphate-buffered saline (PBS/5 mM EDTA) after exposure. Then, cells were detached with PBS/5 mM EDTA and collected by centrifugation. Cells in apoptosis or necrosis were identified after staining with annexin V-FITC and propidium iodide (PI) as described in Ref. . About 1.5 × 10 5 –1 × 10 6 cells per cell culture were incubated in binding buffer with annexin V-FITC and PI, and fluorescence was measured by flow cytometry (FACSCanto). FITC fluorescence (FL-1) was analyzed by a 530/30 band pass filter, and PI fluorescence (Fl-3) by a 650 nm long pass filter. A FACSDiva™ 5.0.2 software was used for data acquisition (2 × 10 4 events for each sample) and for analysis. Numbers of viable cells (annexin V−; PI−) were counted in the lower left quadrant (unstained) of density plots, and the percentages of cells in early apoptosis (annexin V+; PI−), late apoptosis (annexin V+; PI+), or necrosis (annexin V−; PI+) were calculated as described in Ref. .

Cell lysis and protein extraction

Cellular proteins were isolated following established procedures . RAW264.7 mouse macrophages (1–2 × 10 6 cells) were first incubated in cell culture plates (150 mm in diameter) at 37 °C for 24 h. Then, cells were preincubated with diphenylene iodonium (DPI) (0–3 μM), L-NAME (0–100 μM), or VAS2870 (0–30 μM) for 1 h. Next, cells were stimulated with HEMA (0–6–8 mM) in the presence or absence of DPI, L-NAME, or VAS2870 for 24 h. After that, floating and adherent cells were collected in PBS/5 mM EDTA by centrifugation. The cell pellet was resuspended in 1 ml PBS plus 0.5 ml buffer A (10 mM Tris-Cl, 60 mM KCl, 1 mM Na 2 EDTA, 1 mM DTT, pH 7.4), and set on ice for 5 min. After centrifugation, the supernatant was discarded, and the cells were solubilized in buffer B (buffer A plus 0.4% NP-40, 5 mM NaF, 1 mM NaVO 4 , protease inhibitor cocktail) on ice for 3 min. Following centrifugation for 4 min at 16,000 × g , the supernatant was collected as a cytoplasmic fraction. The pellet was resuspended in buffer C (20 mM Tris-Cl pH 8.0, 400 mM NaCl, 1.5 mM MgCl 2 , 1.5 mM Na 2 EDTA, 25% glycerol, 1 mM DTT, 5 mM NaF, 1 mM NaVO 4 , protease inhibitor cocktail), incubated on ice, and then centrifuged at 16,000 × g for 10 min. The supernatant was collected as the nuclear fraction. Protein concentrations in the cytoplasmic and nuclear fractions were determined using a bicinchoninic acid assay .

Immunoblotting

The expression of specific proteins was detected by standard immunoblotting techniques. Briefly, proteins (15 μg/lane) were separated on a 12% sodium dodecyl sulfate-polyacrylamide gel by electrophoresis (SDS-PAGE), and transferred to a nitrocellulose membrane in 25 mM Tris-Cl, 192 mM glycine, 20% methanol (pH 8.3) at 350 mA for 60 min as described in Ref. . The membrane was washed in TBS (25 mM Tris-Cl, 150 mM NaCl, pH 7.4) and blocked with 5% nonfat milk in TBST (TBS plus 0.1% Tween 20, pH 7.4) or bovine serum albumin (detection of GPx1/2) at room temperature for 60 min. After the incubation with specific antibodies overnight at cold temperatures, the membrane was washed with TBST, and primary antibodies were detected by horseradish peroxidase-conjugated secondary antibodies in TBST. Secondary antibodies were then visualized by enhanced chemiluminescence (ECL) and exposure to X-ray films following the manufactureŕs instructions.

Statistical analyses

Individual values (n) in series of independent experiments were summarized (SPSS 23.0, SPSS, Chicago, IL, USA) as medians (25–75% quartiles). Differences between median values were statistically analysed at the 0.05 level of significance using the Mann-Whitney U -test (SPSS 23.0) for pairwise comparisons among groups.

Materials and methods

Chemicals and reagents

2-Hydroxyethyl methacrylate (HEMA; CAS-No. 868-779) was purchased from Merck (Darmstadt, Germany). RPMI 1640 medium containing l -glutamine and 2.0 g/l NaHCO 3 came from PAN Biotech (Aidenbach, Germany). Fetal bovine serum (FBS), penicillin/streptomycin, and phosphate-buffered saline supplemented with 5 mM EDTA (PBS-EDTA) were obtained from Life Technologies, Gibco BRL (Eggenstein, Germany). Dihydroethidium (DHE; CAS-No. 104821-25-2), dihydrorhodamine 123 (DHR123; CAS-No. 109244-58-8), phorbol myristate acetate (PMA; CAS-No. 16561-29-8), oxypurinol (CAS 2465-59-0), VAS2870 (CAS 722456-31-7), and a bicinchoninic acid protein assay kit were purchased from Sigma (Taufkirchen, Germany). 2′7′-Dichlorodihydrofluorescin diacetate (DCFH 2 -DA; CAS-No. 4091-99-0) came from MoBiTec (Göttingen, Germany). An annexin V-FITC apoptosis detection kit was obtained from R&D Systems (Minneapolis, MN, USA). Diphenylene iodonium chloride (CAS 4673-26-1) was acquired from Tocris Bioscience (R&D Systems GmbH, Wiesbaden-Nordenstadt, Germany), L-NG-nitroarginine methyl ester (L-NAME, CAS 51298-62-5) and apocynin (acetovanillone, CAS 498-02-2) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA).

Anti-catalase (H-300, sc-50508), anti-heme oxygenase 1 (HO-1, sc-1797), anti-Nrf2 (H-300, sc-13032), anti-p67-phox (sc-7663) polyclonal antibodies plus anti-Cu–Zn superoxide dismutase (SOD1, sc-271014), anti-glutathione peroxidase 1/2 (GPx1/2, sc-133152), anti-p47-phox (sc-17845), and anti-xanthine oxidoreductase monoclonal antibodies (XOR, sc-398548) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-peroxiredoxin 1 (Prdx1; no. 8732), anti-nitric oxide synthase (NOS) polyclonal antibodies and anti-rabbit IgG HRP-linked antibodies (no. 7074) were purchased from Cell Signaling (NEB Frankfurt, Germany). Goat anti-mouse IgG (H + L)-HRP conjugate came from Bio-Rad Laboratories (Munich, Germany), and Amersham hyperfilm ECL was obtained from GE Healthcare (Munich, Germany). A protease inhibitor cocktail was acquired from Roche Diagnostics (Mannheim, Germany), and an anti-glyceraldehyde-3-phosphate dehydrogenase (GAPDH) monoclonal antibody (6C5) came from Millipore (Schwalbach, Germany). All chemical reagents were of analytical grade.

Cell culture

RAW264.7 mouse macrophages (ATCC TIB71) were grown in RPMI 1640 medium containing glutamine, sodium-pyruvate and 2.0 g/l NaHCO 3 supplemented with 10% fetal bovine serum (FBS) and penicillin-streptomycin at 37 °C and 5% CO 2 as described in Ref. .

Formation of reactive oxygen species (ROS)

The intracellular production of ROS in RAW264.7 mouse macrophages was detected using the oxidation-sensitive probe 2′7′-dichlorodihydrofluorescin diacetate (DCFH 2 -DA), and dihydroethidium (DHE) or dihydrorhodamine 123 (DHR123) specifically indicated the formation of superoxide anions or hydrogen peroxide, respectively . Cell cultures in 6-well plates (5 × 10 4 /well) were stimulated with HEMA (0–6–8 mM) or phorbol myristate acetate (0.0–0.1–1.0 μM PMA) for 1 h or 24 h at 37 °C as described in Ref. . Cells were exposed to PMA as a positive control to show the induction of ROS .

For further analysis, cells were preincubated with diphenylene iodonium (DPI) (0–10 μM) or L-NAME (0–300 μM) for 1 h. After that, cells were exposed to HEMA (0–6–8 mM) or PMA (0.0–0.1–1.0 μM) in the presence or absence of various concentrations of DPI (0–10 μM) or L-NAME (0–300 μM) for 1 h or 24 h. The concentrations of DPI (0–10 μM) or L-NAME (0–300 μM) used here were selected after preliminary range-finding experiments. ROS formation in HEMA-exposed cell cultures was also analyzed using oxypurinol, VAS2870, apocynin, diapocynin or plumbagin but no influence of these inhibitors was detected (unpublished observations).

The formation of ROS was determined by flow cytometry (FACSCanto, Becton Dickinson) as described in Ref. . Briefly, the cells were incubated with 10 μM DCFH 2 -DA, 5 μM DHE or 5 μM DHR123 in complete medium 30 min before harvesting in PBS/5 mM EDTA. Then, cells were routinely collected by centrifugation and resuspended in 200 μl CMF–PBS. DCF fluorescence was measured at an excitation wavelength of 488 nm and an emission wavelength of 519 nm (Fl-1). DHE and DHR123 fluorescence was analyzed at an excitation wavelength of 488 and an emission wavelength of 530 nm (Fl-1, DHE) or 578 nm (Fl-2, DHR123). Mean fluorescence intensities were obtained by histogram statistics (FACSDiva™ 5.0.2 software), and individual values of fluorescence intensities in treated cell cultures were normalized to fluorescence measured in untreated controls (= 1.0) .

Detection of apoptosis and necrosis by flow cytometry

RAW264.7 mouse macrophages (1 × 10 5 cells/well) grown in 6-well plates were preincubated with diphenylene iodonium (DPI) (0–10 μM), apocynin (0–500 μM), L-NAME (0–300 μM), oxypurinol (0–100 μM), or VAS2870 (0–30 μM) for 1 h. Next, preincubation media were removed and cells were stimulated with HEMA (0–6–8 mM) in the presence or absence of DPI (0–10 μM), apocynin (0–500 μM), L-NAME (0–300 μM), oxypurinol (0–100 μM), or VAS2870 (0–30 μM) for 24 h following published procedures . Effective concentrations of the inhibitors were selected both in range finding experiments with HEMA and from their influence observed on PMA-induced apoptosis (unpublished observations).

Apoptosis or necrosis in exposed cells and untreated controls was determined as described elsewhere . Briefly, cell cultures were washed with phosphate-buffered saline (PBS/5 mM EDTA) after exposure. Then, cells were detached with PBS/5 mM EDTA and collected by centrifugation. Cells in apoptosis or necrosis were identified after staining with annexin V-FITC and propidium iodide (PI) as described in Ref. . About 1.5 × 10 5 –1 × 10 6 cells per cell culture were incubated in binding buffer with annexin V-FITC and PI, and fluorescence was measured by flow cytometry (FACSCanto). FITC fluorescence (FL-1) was analyzed by a 530/30 band pass filter, and PI fluorescence (Fl-3) by a 650 nm long pass filter. A FACSDiva™ 5.0.2 software was used for data acquisition (2 × 10 4 events for each sample) and for analysis. Numbers of viable cells (annexin V−; PI−) were counted in the lower left quadrant (unstained) of density plots, and the percentages of cells in early apoptosis (annexin V+; PI−), late apoptosis (annexin V+; PI+), or necrosis (annexin V−; PI+) were calculated as described in Ref. .

Cell lysis and protein extraction

Cellular proteins were isolated following established procedures . RAW264.7 mouse macrophages (1–2 × 10 6 cells) were first incubated in cell culture plates (150 mm in diameter) at 37 °C for 24 h. Then, cells were preincubated with diphenylene iodonium (DPI) (0–3 μM), L-NAME (0–100 μM), or VAS2870 (0–30 μM) for 1 h. Next, cells were stimulated with HEMA (0–6–8 mM) in the presence or absence of DPI, L-NAME, or VAS2870 for 24 h. After that, floating and adherent cells were collected in PBS/5 mM EDTA by centrifugation. The cell pellet was resuspended in 1 ml PBS plus 0.5 ml buffer A (10 mM Tris-Cl, 60 mM KCl, 1 mM Na 2 EDTA, 1 mM DTT, pH 7.4), and set on ice for 5 min. After centrifugation, the supernatant was discarded, and the cells were solubilized in buffer B (buffer A plus 0.4% NP-40, 5 mM NaF, 1 mM NaVO 4 , protease inhibitor cocktail) on ice for 3 min. Following centrifugation for 4 min at 16,000 × g , the supernatant was collected as a cytoplasmic fraction. The pellet was resuspended in buffer C (20 mM Tris-Cl pH 8.0, 400 mM NaCl, 1.5 mM MgCl 2 , 1.5 mM Na 2 EDTA, 25% glycerol, 1 mM DTT, 5 mM NaF, 1 mM NaVO 4 , protease inhibitor cocktail), incubated on ice, and then centrifuged at 16,000 × g for 10 min. The supernatant was collected as the nuclear fraction. Protein concentrations in the cytoplasmic and nuclear fractions were determined using a bicinchoninic acid assay .

Immunoblotting

The expression of specific proteins was detected by standard immunoblotting techniques. Briefly, proteins (15 μg/lane) were separated on a 12% sodium dodecyl sulfate-polyacrylamide gel by electrophoresis (SDS-PAGE), and transferred to a nitrocellulose membrane in 25 mM Tris-Cl, 192 mM glycine, 20% methanol (pH 8.3) at 350 mA for 60 min as described in Ref. . The membrane was washed in TBS (25 mM Tris-Cl, 150 mM NaCl, pH 7.4) and blocked with 5% nonfat milk in TBST (TBS plus 0.1% Tween 20, pH 7.4) or bovine serum albumin (detection of GPx1/2) at room temperature for 60 min. After the incubation with specific antibodies overnight at cold temperatures, the membrane was washed with TBST, and primary antibodies were detected by horseradish peroxidase-conjugated secondary antibodies in TBST. Secondary antibodies were then visualized by enhanced chemiluminescence (ECL) and exposure to X-ray films following the manufactureŕs instructions.

Statistical analyses

Individual values (n) in series of independent experiments were summarized (SPSS 23.0, SPSS, Chicago, IL, USA) as medians (25–75% quartiles). Differences between median values were statistically analysed at the 0.05 level of significance using the Mann-Whitney U -test (SPSS 23.0) for pairwise comparisons among groups.

Results

Formation of reactive oxygen species (ROS) in HEMA-exposed cells

First, the formation and the levels of various kinds of ROS in RAW264.7 mouse macrophages exposed to HEMA were analyzed using different oxidation-sensitive fluorescent dyes. ROS production on cell cultures exposed to phorbol myristate acetate (PMA), which stimulates the formation of superoxide by NADPH oxidases, was used as a reference .

Increasing levels of ROS, as indicated by increasing fluorescence intensities, were detected in HEMA-exposed cells depending on the length of the exposure period. The oxidation of DCFH 2 -DA to 2,7-dichlorofluorescein (DCF) specifically increased in cells exposed to HEMA for 1 h ( Fig. 1 A). DCF fluorescence was significantly enhanced in cells exposed to 6 or 8 mM HEMA about 1.5-fold compared to untreated controls (p = 0.002). While DCF formation only slightly but significantly increased (p = 0.002), PMA caused a clear and concentration-dependent increase in DHE and DHR123 fluorescence. DHE fluorescence was enhanced about 2.9-fold (p = 0.002) and DHR123 fluorescence increased by a factor of 5.5 (p = 0.002) in cells exposed to 1.0 μM PMA ( Fig. 1 A). In contrast, a considerable increase in DHE and DHR123 fluorescence was absent in cultures exposed to HEMA for 1 h ( Fig. 1 A). This pattern of the differential induction of ROS formation by HEMA or PMA was inversed after a long exposure period. No increase in DCF fluorescence was observed in cells exposed to HEMA for 24 h compared to untreated cultures. Noteworthy was a clear increase in DHR123 fluorescence by 1.7-fold (p = 0.002) in cells exposed to 6 mM, or 2.0-fold (p = 0.002) in cultures treated with 8 mM HEMA ( Fig. 1 B). In contrast to HEMA, DCF fluorescence increased about 5-fold in cells exposed to PMA for 24 h, however, DHE and DHR123 fluorescence intensities were lower than after 1 h exposure ( Fig. 1 B).

Fig. 1
Generation of reactive oxygen species (ROS) in RAW264.7 mouse macrophages.
Cells were exposed to HEMA (2-hydroxyethyl methacrylate) (0–6–8 mM) or PMA (phorbol myristate acetate) (0–0.1–1.0 μM) for 1 h (A) or 24 h (B). The formation of ROS was measured using the oxidation-sensitive fluorescent probe 2′7′-dichlorodihydrofluorescin diacetate (DCF fluorescence), dihydroethidium (DHE) for the detection of superoxide anions or dihydrorhodamine 123 (DHR123) for the detection of hydrogen peroxide. Bars represent median values (and 25%/75% percentiles) calculated from individual histograms (n = 6).
UC = untreated cell cultures (0 mM HEMA).
a = significant differences between median values of normalized DCF fluorescence detected in untreated cell cultures (UC; fluorescence factor = 1.0) and cell cultures exposed to HEMA or PMA;
b = significant differences between median values of normalized DHE fluorescence detected in untreated cell cultures and cell cultures exposed to HEMA or PMA;
c = significant differences between median values of normalized DHR123 fluorescence detected in untreated cell cultures and cell cultures exposed to HEMA or PMA.

The influence of diphenylene iodonium on ROS formation through flavoenzymes

Various inhibitors of enzyme activities were used as tools to gain insight into the sources of enhanced ROS formation in HEMA-exposed cells. As a commonly used unspecific inhibitor, DPI (diphenylene iodonium) is known to modify the activities of NADPH oxidases (Nox) and other flavin-containing enzymes like nitric oxide synthase (NOS), xanthine oxidoreductase (XOR), NAD(P)H:quinone oxidoreductase (complex I), or cytochrome P-450 reductase . The influence of DPI on the effect of HEMA was studied in the present investigation at a very broad concentration range from 0.001 to 10 μM, and representative results for HEMA-induced DCF and DHR123 fluorescence are shown ( Fig. 2 ). First, in these series of numerous experiments the increase in DCF and DHR123 fluorescence observed in RAW264.7 mouse macrophages exposed to HEMA for short (1 h) or long (24 h) time periods, as presented in Fig. 1 , remained highly reproducible. A clear concentration-dependent effect was detected with DPI. Low concentrations (0.001–0.1 μM) of DPI alone slightly reduced DCF fluorescence after a short exposure period. The identical pattern of a small, reproducible but statistically insignificant decrease in DCF fluorescence was detectable in HEMA-exposed cultures in the presence of DPI as well ( Fig. 2 A). Most remarkable is that DCF fluorescence increased about 1.5–2-fold in a concentration-dependent manner by high DPI concentrations alone (1–10 μM). It appeared as if this DPI-induced pattern was added to the increase observed in the presence of HEMA, but no further influence of high DPI concentrations on the effect of HEMA was detected ( Fig. 2 B). After a 24 h exposure period, DCF fluorescence was greatly increased about 5-fold even by 0.1 μM DPI ( Fig. 2 C). The same increase in DCF fluorescence was present in culture exposed to 1 or 3 μM DPI ( Fig. 2 D). A decline in DCF fluorescence observed with 10 μM DPI is discussed below. Oxidative stress induced by DPI after a 24 h exposure period was inhibited in the presence of HEMA ( Fig. 2 C and D).

Nov 22, 2017 | Posted by in Dental Materials | Comments Off on Flavin-containing enzymes as a source of reactive oxygen species in HEMA-induced apoptosis
Premium Wordpress Themes by UFO Themes