Ó 2015 Eur J Oral Sci
Eur J Oral Sci 2015; 123: 282–287 DOI: 10.1111/eos.12189 Printed in Singapore. All rights reserved
European Journal of Oral Sciences
Cell toxicity of 2-hydroxyethyl methacrylate (HEMA): the role of oxidative stress
Else Morisbak1, Vibeke Ansteinsson1,2, Jan T. Samuelsen1 1
Nordic Institute of Dental Materials, Oslo; Department of Clinical Dentistry, Faculty of Medicine and Dentistry, University of Bergen, Bergen, Norway
2
Morisbak E, Ansteinsson V, Samuelsen JT. Cell toxicity of 2-hydroxyethyl methacrylate (HEMA): the role of oxidative stress. Eur J Oral Sci 2015; 123: 282–287. © 2015 Eur J Oral Sci 2-Hydroxyethyl methacrylate (HEMA) is a methacrylate monomer used in polymerbased dental-restorative materials. In this study, the viability of human lung epithelial cells, BEAS-2B, was investigated after exposure to this monomer. Exposure to HEMA reduced the viability of the BEAS-2B cells as a result of increased apoptosis, interruption of the cell cycle, and decreased cell proliferation. Depletion of cellular glutathione and increased levels of reactive oxygen species (ROS) were seen after exposure of BEAS-2B cells to HEMA. The glutathione synthase inhibitor, L-buthioninesulfoximine (BSO), was used to study whether the reduced viability was caused by glutathione depletion and increased levels of ROS. Similarly to incubation with HEMA, incubation with BSO resulted in glutathione depletion and increased ROS levels, without increasing cell death or inhibiting cell growth. The results indicate that HEMA-induced cell damage is not caused exclusively by these mechanisms. Mechanisms other than glutathione depletion and ROS formation seem to be of importance for the toxic effect of HEMA on lung epithelial cells.
Polymer-based composite materials are a frequent first choice in restorative dentistry. The composite is combined with an adhesive system that provides bond to the dentin and the enamel. The adhesive can be considered as an integral part of the composite system (1). Composites consist of two main phases: the fillers, usually silica particles of various sizes; and the matrix, which in general is a mixture of methacrylate monomers. The composites are cured in situ by polymerization of the methacrylates (2). Dental personnel handle the uncured materials and are potentially exposed to the monomers on a daily basis. Both direct skin contact and airway exposure to uncured methacrylate occurs (3–6). In addition, methacrylate monomers have been demonstrated in saliva from patients undergoing dental treatment (7), and it has been shown that methacrylate monomers can pass through dentine channels and enter the circulation (1). Detailed knowledge of the mechanism by which chemicals may exert their effect on cells is of great value for risk evaluation of new and existing dental materials. In vitro, methacrylate monomers have been shown to have cytotoxic potential (1, 8–12). Glutathione depletion by adduct formation between glutathione and the methacrylate monomers, followed by formation of reactive oxygen species (ROS), has been suggested as a major mechanism (13, 14). In line with this hypothesis, the antioxidant N-acetyl cysteine (NAC) is reported to counteract the cytotoxic effects of methacrylates (15).
Else Morisbak, Nordic Institute of Dental Materials, Sognsveien 70 A, N-0855 Oslo, Norway E-mail: [email protected] Key words: adhesives; apoptosis; cell proliferation; glutathione; toxicology Accepted for publication March 2015
However, NAC can induce additional effects on the exposed system by spontaneously adducting with several methacrylate monomers, thus merely resulting in a methacrylate-scavenging mechanism and not an antioxidant mechanism (16). It has been shown that other antioxidants do not fully counteract the DNA damage response induced by 2-hydroxyethyl methacrylate (HEMA) (17). Thus, the role of glutathione depletion and ROS formation as a major contributor in the cytotoxic effect of methacrylate monomers has not been fully elucidated. L-buthioninesulfoximine (BSO) is often used to inhibit the synthesis of glutathione in vitro. L-buthioninesulfoximine depletes cellular glutathione content by inhibition of the glutathione synthase enzyme. Glutathione is an important antioxidant in the human body and is found in high concentrations in cells in the cytosol, mitochondria, and the cell nucleus. Glutathione contains the amino acid cysteine, and the antioxidative activity is connected to the thiol group in this amino acid (18). Thus, decreased levels of glutathione may result in increased levels of ROS (19). In the present study we hypothesized that HEMA induces cell-cycle and cell-proliferation disturbances through glutathione depletion and formation of ROS. To test the hypothesis, glutathione depletion and ROS formation were studied in BEAS-2B cells exposed to either HEMA or BSO. Similar cellular effects of the two exposure conditions would support that
Cell toxicity of HEMA: the role of oxidative stress
glutathione depletion followed by increased ROS formation is a major mechanism of HEMA toxicity, whilst different results would suggest that other mechanisms may be involved in HEMA-induced toxicity.
Material and methods Chemicals Lechner and La Veck (LHC9) medium was purchased from GIBCO (Life Technologies, Foster City, CA, USA). Collagen (PureCol) was purchased from Inamed Biomaterials (Fremont, CA, USA). 2-Hydroxyethyl methacrylate (CAS no. 868-77-9), purity ≥97%, was from Fluka Chemie (Buchs, Switzerland). L-buthioninesulfoximine, monobromobimane (mBrB), 20 ,70 -dichlorofluorescein diacetate (DCFH-DA), thiazolyl blue tetrazolium bromide (MTT), dimethyl sulphoxide (DMSO), Hoechst 33342, and propidium iodide (PI), were purchased from Sigma–Aldrich (St Louis, MO, USA). Nuclear isolation and staining solution was purchased from NPE systems (Pembroke Pines, FL, USA). All other chemicals were purchased from commercial sources and were of the highest purity available. Cell cultures and treatment BEAS-2B, an SV40 hybrid (Ad12-SV40)-transformed human bronchial epithelial cell line, was purchased from the European Tissue Type Culture Collection (ECACC). The cells were cultured in serum-free Lechner and LaVeck (LHC9) medium that was replaced every second day. The cells were passaged when they had reached >85% confluence. Culture flasks and wells were precoated in HEPESbuffered saline (HBS) with collagen (30 lg/ml). Twenty four hours before exposure, cells were seeded on flat bottom plates (6 and 24 wells) or in single dishes (9.5 cm2) (Costar, Corning, NY, USA). One hour before the start of the experiments, the cell culture medium was replaced with fresh medium. A stock solution of HEMA was prepared by dissolving HEMA in cell culture medium to a concentration of 500 mM using a Vortex mixer. 2-Hydroxyethyl methacrylate was added to cell culture wells to final concentrations of 2.5, 5, and 10 mM. A stock solution of BSO was prepared by dissolving solid BSO in cell culture medium to a concentration of 1 mM using a Vortex mixer. L-buthioninesulfoximine was added to cell culture wells to a final concentration of 10 lM. Control cells were cultured in medium without addition of HEMA or BSO.
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Intracellular ROS The level of intracellular ROS was measured using the fluorescent probe DCFH-DA, as previously described (21). The principle of the test is based on the diffusion of DCFH-DA into cells, where it is hydrolyzed to non-fluorescent 2,7-dichlorofluorescin (DCFH). Intracellular ROS then causes oxidation of DCFH to a measurable fluorescent product, DCF. After exposure to HEMA and BSO, DCFH-DA (20 lM) was added to the cells. The cells were harvested after 15 min and washed in PBS. Then, DCF fluorescence was measured using flow cytometry (Cell Lab Quanta SC; Beckman Coulter). Cell-viability
PI/Hoechst staining: To determine plasma-membrane damage and changes in nuclear morphology, the cells were stained with PI (5 lg/ml) and Hoechst 33342 (10 lg/ml), then analysed using fluorescence microscopy (Olympus BX51; Olympus Europe, Hamburg, Germany). Cells with clearly condensed and/or fragmented nuclei (both PI-negative and PI-positive), as well as cells with partially condensed chromatin (PI-negative), were counted as apoptotic and included in the fraction of the total number of cells. Propidium iodide-stained cells that showed a rounded morphology and a homogeneously stained nucleus, or partially condensed chromatin with less fluorescence intensity, were termed necrotic. Nonapoptotic cells, excluding PI, were considered as viable cells. Approximately 300–400 cells from each sample were counted. MTT: The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide) assay was used as a toxicity assay. Cleavage of the tetrazolium salt MTT into a bluecoloured product (formazan) is dependent on the activity of the mitochondrial enzyme, succinate dehydrogenase (SDH) (22). In short, 300 ll MTT solution (0.5 mg/ml of MTT in PBS) was added to each well containing unexposed, HEMA exposed (24 h) and BSO exposed (24 h) cells. After incubation for 1 h at 37°C, the MTT solution was removed and DMSO was added to solubilize the formazan product. The plate was shaken and absorption at 570 nm was measured using a spectrophotometer (Synergy H1; BioTek Instruments, Winooski, VT, USA). The value recorded for each sample was used as a measure of the density of viable cells. Cell proliferation
Cellular glutathione The relative concentrations of intracellular glutathione were determined according to a previously described assay using mBrB (20). Monobromobimane binds to the SH group of glutathione (reduced form), thereby forming a fluorescent adduct. A stock solution of 40 mM mBrB in methanol was diluted with PBS to a final concentration of 40 lM. After exposure to HEMA and BSO, the cells were trypsinized and incubated with mBrB (40 lM) for 10 min. The total amount of fluorescence per cell was measured using flow cytometry (Cell Lab Quanta SC; Beckman Coulter, Brea, CA, USA; excitation filter of 350 nm, emission filter of 465 nm). At least 5,000 cells were recorded in each sample, and the average value was used to compare relative GSH levels between samples.
BEAS-2B cells were seeded on dishes containing a 2 mm grid (Corning 430196; Corning, Corning, NY, USA). The following day, the cells were exposed to HEMA and BSO. Specific areas of each dish were marked, and cell proliferation in these areas was monitored by phase contrast microscopy (Olympus CKX41 with Olympus C7070; Olympus Europe) and photographed at the start of the exposure and at various time points up to 48 h. Cell cycle analysis After treatment, cells were harvested, and cellular DNA was obtained and stained using a nuclear staining solution. Fluorescence was measured using a flow cytometer (Cell Lab Quanta SC; Beckman Coulter). Different phases of
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the cell cycle were distinguished on the basis of DNA content. DNA histograms were analysed using MULTICYCLE software (Phoenics Flow Systems, San Diego, CA, USA).
A
Statistics Statistical significance of treatment was evaluated using one-way ANOVA followed by Bonferroni’s test for multiple comparisons. All analyses were performed using the statistical software GRAPHPAD PRISM (GraphPad Software, San Diego, CA, USA). Results were calculated as mean SD, and P < 0.05 was considered statistically significant.
B
Results Effect of HEMA and BSO on glutathione and ROS levels
Twenty-four hours of exposure to HEMA and 24 h of exposure to BSO resulted in significantly reduced glutathione levels in BEAS-2B cells. There was no significant difference between the glutathione depletion obtained in the two conditions; relative to unexposed cells, the reduction in glutathione concentration was 47 21% after exposure to HEMA and 34 17% after exposure to BSO (Fig. 1A). The levels of ROS were measured 6 and 24 h after exposure to either HEMA or BSO. Treatment with either HEMA or BSO resulted in significantly increased levels of ROS in BEAS-2B cells (Fig. 1B). There was no significant difference in the levels of ROS between cells exposed to HEMA and cells exposed to BSO. After 6 h, the increase relative to unexposed cells was 145 16% and 123 4%, for cells exposed to HEMA and BSO, respectively, and the increase after 24 h of exposure was 126 24% and 148 37%, for cells exposed to HEMA and BSO, respectively.
Fig. 2. Density of viable cells relative to control cells, assessed using the thiazolyl blue tetrazolium bromide (MTT) assay, measuring succinate dehydrogenase activity after 24 h of exposure to 2.5, 5, and 10 mM 2-hydroxyethyl methacrylate (HEMA) and to 10 lM L-buthioninesulfoximine (BSO) (A). The percentage of apoptotic cells after 24 h of exposure to 5 mM HEMA and 10 lM BSO is shown in (B). The data are mean SD of at least three separate experiments. *Statistical difference compared with unexposed cells (control); #statistical difference between cells exposed to HEMA (5 and 10 mM) and cells exposed to BSO (P < 0.05).
Cell viability and cell death induced by HEMA or BSO
A significant decrease in cell viability was observed after 24 h of exposure to 5 and 10 mM HEMA (13 4%, and 31 8%, respectively, relative to unexposed cells; P < 0.05) but not after exposure to BSO (4 3%) (Fig. 2A). 2-Hydroxyethyl methacrylate induced a significant increase in apoptotic cell death
A
B
Fig. 1. Relative amount of glutathione (GSH) (A) in cells exposed to 5 mM 2-hydroxyethyl methacrylate (HEMA) or to 10 lM L-buthioninesulfoximine (BSO) for 24 h, and the relative amount of reactive oxygen species (ROS) (B) in cells exposed to 5 mM 2-hydroxyethyl methacrylate (HEMA) (solid bars) or to 10 lM L-buthioninesulfoximine (BSO) (open bars) for 6 and 24 h. The data are presented as mean SD of at least three separate experiments. *Statistical difference from unexposed cells (control) (P < 0.05).
Fig. 3. Photographs of phase-contrast microscopy images of BEAS-2B cell cultures, unexposed (control) and exposed to 5 mM 2-hydroxyethyl methacrylate (HEMA) or to 10 lM L-buthioninesulfoximine (BSO). To document cell proliferation, photographs were taken of the same areas at the start of exposure (0 h) and after 24 and 48 h of exposure.
Cell toxicity of HEMA: the role of oxidative stress
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Discussion
Fig. 4. Cell density, measured as a percentage of control cell density, was assessed using the thiazolyl blue tetrazolium bromide (MTT) assay after 24 and 48 h of exposure to 5 mM 2-hydroxyethyl methacrylate (HEMA) (solid bars) or to 10 lM L-buthioninesulfoximine (BSO) (open bars). The data are presented as mean SD of at least three separate experiments. *Statistical difference (P < 0.05) compared with unexposed (control) cells at the same time point.
(8 3%, (3 1%, apoptotic (4 2%)
P < 0.05) compared with control cells P < 0.05); however, no significant increase in cell death was observed in BSO-treated cells (Fig. 2B).
Reduction in cell growth
BEAS-2B cells exposed to HEMA (5 mM) for up to 48 h showed a marked reduction in cell growth compared with the control cells (Fig. 3). In contrast, BSO did not induce a similar reduction in cell growth in cell culture. The microscopy observations were confirmed by the results of the MTT assay, which showed reduced cell density in the HEMA-exposed cells compared with unexposed cells. No change was seen in the BSOexposed cells (Fig. 4). Cell cycle arrest
Representative cell cycle histograms of unexposed cells and cells exposed to HEMA and BSO are shown in Fig. 5. Cells exposed to HEMA showed marked accumulation of cells in early S-phase (Fig. 5B) compared with unexposed cells (Fig. 5A). The relative amount of unexposed BEAS-2B cells in S-phase was 31 2%, whereas HEMA induced a significant accumulation of cells in S-phase (52 8%, P < 0.05). In contrast to HEMA, BSO did not induce a significant increase of the number of cells in S-phase compared with unexposed cells (34 2%) (Fig. 6). A
B
We have compared the effects of the methacrylate monomer, HEMA, with the effects of the glutathione synthase inhibitor, BSO, on a human bronchial epithelial cell line (BEAS-2B). It has previously been found that methacrylate monomers induce glutathione depletion by adduct formation (14, 23). The glutathione depletion is assumed to be the cause of the reported ROS increase in cells exposed to methacrylate (14, 24). Increased intracellular ROS is potentially toxic to cells, and we wanted to study whether glutathione depletion by BSO had similar effects on cells as exposure to HEMA. We found a similar degree of glutathione depletion in cells exposed to either HEMA or BSO. In addition, increased levels of ROS were observed under the two exposure conditions. Based on these findings, we evaluated the use of BSO as an appropriate model to study the cytotoxic effects of glutathione depletion. BEAS-2B cells were employed after reports on HEMA detected in the air at dental surgeries. This implies exposure of bronchial cells, of both staff and patients, to HEMA (3, 5). Significantly decreased viability and increased numbers of apoptotic cells were observed for BEAS-2B cells exposed to HEMA for 24 h, whereas no change was observed after exposure to BSO. Exposure both to HEMA and to BSO resulted in glutathione depletion and increased ROS. These results do not support the hypothesis that HEMA exerts its toxicity by lowering the cellular glutathione level, at least not exclusively. A slight delay in cellular glutathione depletion and ROS formation in BSO-treated cells could result in a delayed onset of apoptosis and explain the difference measured after 24 h of exposure. Arguing this, we did not observe increased cell death when extending the BSO exposure time to 48 h (data not shown). The previously reported lack of correlation between glutathione depletion and cell viability after exposure to different methacrylate monomers (23) also suggests additional toxic mechanisms of at least some methacrylates. The absence of a direct link between glutathione level and cell death is further supported by a study in which no cell death was observed in a mouse macrophage cell
C
Fig. 5. Cell-cycle histograms of one representative experiment. Unexposed cells (A) and cells exposed to 2-hydroxyethyl methacrylate (HEMA) (5 mM) (B) or to L-buthioninesulfoximine (BSO) (10 lM) (C) for 24 h. Data shown are one representative of at least four experiments.
Fig. 6. Percentage of cells in S-phase, without exposure or after 24 h of exposure to 2-hydroxyethyl methacrylate (HEMA) (5 mM) or to L-buthioninesulfoximine (BSO) (10 lM). The data are mean SD of at least four separate experiments. *Statistical difference between unexposed and HEMA-exposed cells; #statistical difference between HEMAexposed and BSO-exposed cells (P < 0.05). The data are calculated from the histograms shown in Fig 5.
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line treated with BSO for 24 h (24). This study, however, showed increased cell death after exposure to BSO and HEMA in combination compared with HEMA alone. Taken together, glutathione depletion alone does not explain the toxic response to HEMA, but low cellular levels of glutathione could make the cells more susceptible to HEMA exposure. Glutathione–HEMA adduct formation may be an advantageous event through lowering the concentration of free intracellular HEMA. This is in agreement with the current understanding of the phase II metabolism of xenobiotics, where, for example, glutathione conjugation of xenobiotics is considered a beneficial detoxification reaction (18). Similarly to the different effects of HEMA and BSO on cell death, only HEMA reduced proliferation of BEAS2B cells. Both microscopic observations and flow cytometric measurements indicated cell-growth disturbance after exposure to HEMA, but failed to detect a difference in cell growth between BSO-treated cells and the control. The visual appearance of the DNA histograms also differed markedly. 2-Hydroxyethyl methacrylate (HEMA) induced a ‘tail’ in the transition phase between the G1 and S-phases which was not observed in BSO-treated cells or the control cells. Furthermore, the proportion of cells in S-phase was significantly increased in HEMAexposed cells, but not in BSO-treated cells. The role of glutathione in cell proliferation is reported to be of importance to the control of tumour growth, and glutathione depletion is a desired effect of several anticancer drugs (25). However, the different pattern of cell growth and cell death after exposure to HEMA and BSO argues that glutathione depletion alone is the cause of these HEMA-induced changes. An alternative mechanism for the observed cell cycle changes in HEMA-exposed cells could be the binding of HEMA to nucleophilic groups in DNA, thereby forming a HEMA–DNA adduct (15, 26). The finding, of KRIFKA et al. (24), of increased cytotoxity in cells incubated with both HEMA and BSO, could be in accordance with a theory of glutathione serving as a detoxifying agent, binding free HEMA in the cell and thereby preventing HEMA–DNA adduct formation. In the present study we hypothesized that HEMA induces cell-proliferation disturbances and cell toxicity through glutathione depletion and subsequent ROS formation. In contrast to HEMA, exposure to BSO, which leads to a similar depletion of glutathione and elevation of ROS, did not induce cell toxicity and cell cycle disturbances. This indicates that HEMA-induced cell toxicity is not caused by glutathione depletion and ROS formation alone.
2. 3. 4.
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17. Acknowledgements – The authors wish to thank Professor Jon E. Dahl for critical revision of the manuscript. We are also thankful to Dr John E. Tibballs for useful discussion and comments on the manuscript. Conflicts of interest – The authors have no conflicts to declare.
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HOET P, VAN MEERBEEK B. How much do resin-based dental materials release? A meta-analytical approach Dent Mater 2011; 27: 723–747. FERRACANE JL. Resin composite–state of the art. Dent Mater 2011; 27: 29–38. HAGBERG S, LJUNGKVIST G, ANDREASSON H, KARLSSON S, BARREGARD L. Exposure to volatile methacrylates in dental personnel. J Occup Environ Hyg 2005; 2: 302–306. HELMIS CG, TZOUTZAS J, FLOCAS HA, HALIOS CH, STATHOPOULOU OI, ASSIMAKOPOULOS VD, PANIS V, APOSTOLATOU M, SGOUROS G, ADAM E. Indoor air quality in a dentistry clinic. Sci Total Environ 2007; 377: 349–365. HENRIKS-ECKERMAN ML, ALANKO K, JOLANKI R, KEROSUO H, KANERVA L. Exposure to airborne methacrylates and natural rubber latex allergens in dental clinics. J Environ Monit 2001; 3: 302–305. MARQUARDT W, SEISS M, HICKEL R, REICHL FX. Volatile methacrylates in dental practices. J Adhes Dent 2009; 11: 101–107. MICHELSEN VB, KOPPERUD HB, LYGRE GB, BJORKMAN L, JENSEN E, KLEVEN IS, SVAHN J, LYGRE H. Detection and quantification of monomers in unstimulated whole saliva after treatment with resin-based composite fillings in vivo. Eur J Oral Sci 2012; 120: 89–95. BECHER R, KOPPERUD HM, AL RH, SAMUELSEN JT, MORISBAK E, DAHLMAN HJ, LILLEAAS EM, DAHL JE. Pattern of cell death after in vitro exposure to GDMA, TEGDMA, HEMA and two compomer extracts. Dent Mater 2006; 22: 630–640. GEURTSEN W, LEHMANN F, SPAHL W, LEYHAUSEN G. Cytotoxicity of 35 dental resin composite monomers/additives in permanent 3T3 and three human primary fibroblast cultures. J Biomed Mater Res 1998; 41: 474–480. SAMUELSEN JT, DAHL JE, KARLSSON S, MORISBAK E, BECHER R. Apoptosis induced by the monomers HEMA and TEGDMA involves formation of ROS and differential activation of the MAP-kinases p38, JNK and ERK. Dent Mater 2007; 23: 34–39. SCHWEIKL H, SPAGNUOLO G, SCHMALZ G. Genetic and cellular toxicology of dental resin monomers. J Dent Res 2006; 85: 870–877. THONEMANN B, SCHMALZ G, HILLER KA, SCHWEIKL H. Responses of L929 mouse fibroblasts, primary and immortalized bovine dental papilla-derived cell lines to dental resin components. Dent Mater 2002; 18: 318–323. KRIFKA S, PETZEL C, HILLER KA, FRANK EM, BOSL C, SPAGNUOLO G, REICHL FX, SCHMALZ G, SCHWEIKL H. Resin monomer-induced differential activation of MAP kinases and apoptosis in mouse macrophages and human pulp cells. Biomaterials 2010; 31: 2964–2975. SAMUELSEN JT, KOPPERUD HM, HOLME JA, DRAGLAND IS, CHRISTENSEN T, DAHL JE. Role of thiol-complex formation in 2-hydroxyethyl- methacrylate-induced toxicity in vitro. J Biomed Mater Res A 2010; 96: 395–401. SCHWEIKL H, HARTMANN A, HILLER KA, SPAGNUOLO G, BOLAY C, BROCKHOFF G, SCHMALZ G. Inhibition of TEGDMA and HEMA-induced genotoxicity and cell cycle arrest by N-acetylcysteine. Dent Mater 2007; 23: 688–695. NOCCA G, D’ANTO V, DESIDERIO C, ROSSETTI DV, VALLETTA R, BAQUALA AM, SCHWEIKL H, LUPI A, RENGO S, SPAGNUOLO G. N-acetyl cysteine directed detoxification of 2-hydroxyethyl methacrylate by adduct formation. Biomaterials 2010; 31: 2508–2516. ANSTEINSSON V, SOLHAUG A, SAMUELSEN JT, HOLME JA, DAHL JE. DNA-damage, cell-cycle arrest and apoptosis induced in BEAS-2B cells by 2-hydroxyethyl methacrylate (HEMA). Mutat Res 2011; 723: 158–164. BOELSTERLI UA. Mechanistic toxicology, 2nd ed. New York: Informa healthcare, 2009; 117–175. GRIFFITH OW, MEISTER A. Potent and specific inhibition of glutathione synthesis by buthionine sulfoximine (S-n-butyl homocysteine sulfoximine). J Biol Chem 1979; 254: 7558–7560. COTGREAVE IA, MOLDEUS P. Methodologies for the application of monobromobimane to the simultaneous analysis of soluble and protein thiol components of biological systems. J Biochem Biophys Methods 1986; 13: 231–249.
Cell toxicity of HEMA: the role of oxidative stress 21. WANG H, JOSEPH JA. Quantifying cellular oxidative stress by dichlorofluorescein assay using microplate reader. Free Radical Biol Med 1999; 27: 612–616. 22. MOSMANN T. Rapid colorimetric assay for cellular growth and survival: application to proliferation and cytotoxicity assays. J Immunol Methods 1983; 65: 55–63. 23. ANSTEINSSON V, KOPPERUD HB, MORISBAK E, SAMUELSEN JT. Cell toxicity of methacrylate monomers-The role of glutathione adduct formation. J Biomed Mater Res A 2013; 101: 3504–3510. 24. KRIFKA S, HILLER KA, SPAGNUOLO G, JEWETT A, SCHMALZ G, SCHWEIKL H. The influence of glutathione on redox regulation
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Eur J Oral Sci 2015; 123: 282–287 DOI: 10.1111/eos.12189 Printed in Singapore. All rights reserved
European Journal of Oral Sciences
Cell toxicity of 2-hydroxyethyl methacrylate (HEMA): the role of oxidative stress
Else Morisbak1, Vibeke Ansteinsson1,2, Jan T. Samuelsen1 1
Nordic Institute of Dental Materials, Oslo; Department of Clinical Dentistry, Faculty of Medicine and Dentistry, University of Bergen, Bergen, Norway
2
Morisbak E, Ansteinsson V, Samuelsen JT. Cell toxicity of 2-hydroxyethyl methacrylate (HEMA): the role of oxidative stress. Eur J Oral Sci 2015; 123: 282–287. © 2015 Eur J Oral Sci 2-Hydroxyethyl methacrylate (HEMA) is a methacrylate monomer used in polymerbased dental-restorative materials. In this study, the viability of human lung epithelial cells, BEAS-2B, was investigated after exposure to this monomer. Exposure to HEMA reduced the viability of the BEAS-2B cells as a result of increased apoptosis, interruption of the cell cycle, and decreased cell proliferation. Depletion of cellular glutathione and increased levels of reactive oxygen species (ROS) were seen after exposure of BEAS-2B cells to HEMA. The glutathione synthase inhibitor, L-buthioninesulfoximine (BSO), was used to study whether the reduced viability was caused by glutathione depletion and increased levels of ROS. Similarly to incubation with HEMA, incubation with BSO resulted in glutathione depletion and increased ROS levels, without increasing cell death or inhibiting cell growth. The results indicate that HEMA-induced cell damage is not caused exclusively by these mechanisms. Mechanisms other than glutathione depletion and ROS formation seem to be of importance for the toxic effect of HEMA on lung epithelial cells.
Polymer-based composite materials are a frequent first choice in restorative dentistry. The composite is combined with an adhesive system that provides bond to the dentin and the enamel. The adhesive can be considered as an integral part of the composite system (1). Composites consist of two main phases: the fillers, usually silica particles of various sizes; and the matrix, which in general is a mixture of methacrylate monomers. The composites are cured in situ by polymerization of the methacrylates (2). Dental personnel handle the uncured materials and are potentially exposed to the monomers on a daily basis. Both direct skin contact and airway exposure to uncured methacrylate occurs (3–6). In addition, methacrylate monomers have been demonstrated in saliva from patients undergoing dental treatment (7), and it has been shown that methacrylate monomers can pass through dentine channels and enter the circulation (1). Detailed knowledge of the mechanism by which chemicals may exert their effect on cells is of great value for risk evaluation of new and existing dental materials. In vitro, methacrylate monomers have been shown to have cytotoxic potential (1, 8–12). Glutathione depletion by adduct formation between glutathione and the methacrylate monomers, followed by formation of reactive oxygen species (ROS), has been suggested as a major mechanism (13, 14). In line with this hypothesis, the antioxidant N-acetyl cysteine (NAC) is reported to counteract the cytotoxic effects of methacrylates (15).
Else Morisbak, Nordic Institute of Dental Materials, Sognsveien 70 A, N-0855 Oslo, Norway E-mail: [email protected] Key words: adhesives; apoptosis; cell proliferation; glutathione; toxicology Accepted for publication March 2015
However, NAC can induce additional effects on the exposed system by spontaneously adducting with several methacrylate monomers, thus merely resulting in a methacrylate-scavenging mechanism and not an antioxidant mechanism (16). It has been shown that other antioxidants do not fully counteract the DNA damage response induced by 2-hydroxyethyl methacrylate (HEMA) (17). Thus, the role of glutathione depletion and ROS formation as a major contributor in the cytotoxic effect of methacrylate monomers has not been fully elucidated. L-buthioninesulfoximine (BSO) is often used to inhibit the synthesis of glutathione in vitro. L-buthioninesulfoximine depletes cellular glutathione content by inhibition of the glutathione synthase enzyme. Glutathione is an important antioxidant in the human body and is found in high concentrations in cells in the cytosol, mitochondria, and the cell nucleus. Glutathione contains the amino acid cysteine, and the antioxidative activity is connected to the thiol group in this amino acid (18). Thus, decreased levels of glutathione may result in increased levels of ROS (19). In the present study we hypothesized that HEMA induces cell-cycle and cell-proliferation disturbances through glutathione depletion and formation of ROS. To test the hypothesis, glutathione depletion and ROS formation were studied in BEAS-2B cells exposed to either HEMA or BSO. Similar cellular effects of the two exposure conditions would support that
Cell toxicity of HEMA: the role of oxidative stress
glutathione depletion followed by increased ROS formation is a major mechanism of HEMA toxicity, whilst different results would suggest that other mechanisms may be involved in HEMA-induced toxicity.
Material and methods Chemicals Lechner and La Veck (LHC9) medium was purchased from GIBCO (Life Technologies, Foster City, CA, USA). Collagen (PureCol) was purchased from Inamed Biomaterials (Fremont, CA, USA). 2-Hydroxyethyl methacrylate (CAS no. 868-77-9), purity ≥97%, was from Fluka Chemie (Buchs, Switzerland). L-buthioninesulfoximine, monobromobimane (mBrB), 20 ,70 -dichlorofluorescein diacetate (DCFH-DA), thiazolyl blue tetrazolium bromide (MTT), dimethyl sulphoxide (DMSO), Hoechst 33342, and propidium iodide (PI), were purchased from Sigma–Aldrich (St Louis, MO, USA). Nuclear isolation and staining solution was purchased from NPE systems (Pembroke Pines, FL, USA). All other chemicals were purchased from commercial sources and were of the highest purity available. Cell cultures and treatment BEAS-2B, an SV40 hybrid (Ad12-SV40)-transformed human bronchial epithelial cell line, was purchased from the European Tissue Type Culture Collection (ECACC). The cells were cultured in serum-free Lechner and LaVeck (LHC9) medium that was replaced every second day. The cells were passaged when they had reached >85% confluence. Culture flasks and wells were precoated in HEPESbuffered saline (HBS) with collagen (30 lg/ml). Twenty four hours before exposure, cells were seeded on flat bottom plates (6 and 24 wells) or in single dishes (9.5 cm2) (Costar, Corning, NY, USA). One hour before the start of the experiments, the cell culture medium was replaced with fresh medium. A stock solution of HEMA was prepared by dissolving HEMA in cell culture medium to a concentration of 500 mM using a Vortex mixer. 2-Hydroxyethyl methacrylate was added to cell culture wells to final concentrations of 2.5, 5, and 10 mM. A stock solution of BSO was prepared by dissolving solid BSO in cell culture medium to a concentration of 1 mM using a Vortex mixer. L-buthioninesulfoximine was added to cell culture wells to a final concentration of 10 lM. Control cells were cultured in medium without addition of HEMA or BSO.
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Intracellular ROS The level of intracellular ROS was measured using the fluorescent probe DCFH-DA, as previously described (21). The principle of the test is based on the diffusion of DCFH-DA into cells, where it is hydrolyzed to non-fluorescent 2,7-dichlorofluorescin (DCFH). Intracellular ROS then causes oxidation of DCFH to a measurable fluorescent product, DCF. After exposure to HEMA and BSO, DCFH-DA (20 lM) was added to the cells. The cells were harvested after 15 min and washed in PBS. Then, DCF fluorescence was measured using flow cytometry (Cell Lab Quanta SC; Beckman Coulter). Cell-viability
PI/Hoechst staining: To determine plasma-membrane damage and changes in nuclear morphology, the cells were stained with PI (5 lg/ml) and Hoechst 33342 (10 lg/ml), then analysed using fluorescence microscopy (Olympus BX51; Olympus Europe, Hamburg, Germany). Cells with clearly condensed and/or fragmented nuclei (both PI-negative and PI-positive), as well as cells with partially condensed chromatin (PI-negative), were counted as apoptotic and included in the fraction of the total number of cells. Propidium iodide-stained cells that showed a rounded morphology and a homogeneously stained nucleus, or partially condensed chromatin with less fluorescence intensity, were termed necrotic. Nonapoptotic cells, excluding PI, were considered as viable cells. Approximately 300–400 cells from each sample were counted. MTT: The MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide) assay was used as a toxicity assay. Cleavage of the tetrazolium salt MTT into a bluecoloured product (formazan) is dependent on the activity of the mitochondrial enzyme, succinate dehydrogenase (SDH) (22). In short, 300 ll MTT solution (0.5 mg/ml of MTT in PBS) was added to each well containing unexposed, HEMA exposed (24 h) and BSO exposed (24 h) cells. After incubation for 1 h at 37°C, the MTT solution was removed and DMSO was added to solubilize the formazan product. The plate was shaken and absorption at 570 nm was measured using a spectrophotometer (Synergy H1; BioTek Instruments, Winooski, VT, USA). The value recorded for each sample was used as a measure of the density of viable cells. Cell proliferation
Cellular glutathione The relative concentrations of intracellular glutathione were determined according to a previously described assay using mBrB (20). Monobromobimane binds to the SH group of glutathione (reduced form), thereby forming a fluorescent adduct. A stock solution of 40 mM mBrB in methanol was diluted with PBS to a final concentration of 40 lM. After exposure to HEMA and BSO, the cells were trypsinized and incubated with mBrB (40 lM) for 10 min. The total amount of fluorescence per cell was measured using flow cytometry (Cell Lab Quanta SC; Beckman Coulter, Brea, CA, USA; excitation filter of 350 nm, emission filter of 465 nm). At least 5,000 cells were recorded in each sample, and the average value was used to compare relative GSH levels between samples.
BEAS-2B cells were seeded on dishes containing a 2 mm grid (Corning 430196; Corning, Corning, NY, USA). The following day, the cells were exposed to HEMA and BSO. Specific areas of each dish were marked, and cell proliferation in these areas was monitored by phase contrast microscopy (Olympus CKX41 with Olympus C7070; Olympus Europe) and photographed at the start of the exposure and at various time points up to 48 h. Cell cycle analysis After treatment, cells were harvested, and cellular DNA was obtained and stained using a nuclear staining solution. Fluorescence was measured using a flow cytometer (Cell Lab Quanta SC; Beckman Coulter). Different phases of
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the cell cycle were distinguished on the basis of DNA content. DNA histograms were analysed using MULTICYCLE software (Phoenics Flow Systems, San Diego, CA, USA).
A
Statistics Statistical significance of treatment was evaluated using one-way ANOVA followed by Bonferroni’s test for multiple comparisons. All analyses were performed using the statistical software GRAPHPAD PRISM (GraphPad Software, San Diego, CA, USA). Results were calculated as mean SD, and P < 0.05 was considered statistically significant.
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Results Effect of HEMA and BSO on glutathione and ROS levels
Twenty-four hours of exposure to HEMA and 24 h of exposure to BSO resulted in significantly reduced glutathione levels in BEAS-2B cells. There was no significant difference between the glutathione depletion obtained in the two conditions; relative to unexposed cells, the reduction in glutathione concentration was 47 21% after exposure to HEMA and 34 17% after exposure to BSO (Fig. 1A). The levels of ROS were measured 6 and 24 h after exposure to either HEMA or BSO. Treatment with either HEMA or BSO resulted in significantly increased levels of ROS in BEAS-2B cells (Fig. 1B). There was no significant difference in the levels of ROS between cells exposed to HEMA and cells exposed to BSO. After 6 h, the increase relative to unexposed cells was 145 16% and 123 4%, for cells exposed to HEMA and BSO, respectively, and the increase after 24 h of exposure was 126 24% and 148 37%, for cells exposed to HEMA and BSO, respectively.
Fig. 2. Density of viable cells relative to control cells, assessed using the thiazolyl blue tetrazolium bromide (MTT) assay, measuring succinate dehydrogenase activity after 24 h of exposure to 2.5, 5, and 10 mM 2-hydroxyethyl methacrylate (HEMA) and to 10 lM L-buthioninesulfoximine (BSO) (A). The percentage of apoptotic cells after 24 h of exposure to 5 mM HEMA and 10 lM BSO is shown in (B). The data are mean SD of at least three separate experiments. *Statistical difference compared with unexposed cells (control); #statistical difference between cells exposed to HEMA (5 and 10 mM) and cells exposed to BSO (P < 0.05).
Cell viability and cell death induced by HEMA or BSO
A significant decrease in cell viability was observed after 24 h of exposure to 5 and 10 mM HEMA (13 4%, and 31 8%, respectively, relative to unexposed cells; P < 0.05) but not after exposure to BSO (4 3%) (Fig. 2A). 2-Hydroxyethyl methacrylate induced a significant increase in apoptotic cell death
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B
Fig. 1. Relative amount of glutathione (GSH) (A) in cells exposed to 5 mM 2-hydroxyethyl methacrylate (HEMA) or to 10 lM L-buthioninesulfoximine (BSO) for 24 h, and the relative amount of reactive oxygen species (ROS) (B) in cells exposed to 5 mM 2-hydroxyethyl methacrylate (HEMA) (solid bars) or to 10 lM L-buthioninesulfoximine (BSO) (open bars) for 6 and 24 h. The data are presented as mean SD of at least three separate experiments. *Statistical difference from unexposed cells (control) (P < 0.05).
Fig. 3. Photographs of phase-contrast microscopy images of BEAS-2B cell cultures, unexposed (control) and exposed to 5 mM 2-hydroxyethyl methacrylate (HEMA) or to 10 lM L-buthioninesulfoximine (BSO). To document cell proliferation, photographs were taken of the same areas at the start of exposure (0 h) and after 24 and 48 h of exposure.
Cell toxicity of HEMA: the role of oxidative stress
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Discussion
Fig. 4. Cell density, measured as a percentage of control cell density, was assessed using the thiazolyl blue tetrazolium bromide (MTT) assay after 24 and 48 h of exposure to 5 mM 2-hydroxyethyl methacrylate (HEMA) (solid bars) or to 10 lM L-buthioninesulfoximine (BSO) (open bars). The data are presented as mean SD of at least three separate experiments. *Statistical difference (P < 0.05) compared with unexposed (control) cells at the same time point.
(8 3%, (3 1%, apoptotic (4 2%)
P < 0.05) compared with control cells P < 0.05); however, no significant increase in cell death was observed in BSO-treated cells (Fig. 2B).
Reduction in cell growth
BEAS-2B cells exposed to HEMA (5 mM) for up to 48 h showed a marked reduction in cell growth compared with the control cells (Fig. 3). In contrast, BSO did not induce a similar reduction in cell growth in cell culture. The microscopy observations were confirmed by the results of the MTT assay, which showed reduced cell density in the HEMA-exposed cells compared with unexposed cells. No change was seen in the BSOexposed cells (Fig. 4). Cell cycle arrest
Representative cell cycle histograms of unexposed cells and cells exposed to HEMA and BSO are shown in Fig. 5. Cells exposed to HEMA showed marked accumulation of cells in early S-phase (Fig. 5B) compared with unexposed cells (Fig. 5A). The relative amount of unexposed BEAS-2B cells in S-phase was 31 2%, whereas HEMA induced a significant accumulation of cells in S-phase (52 8%, P < 0.05). In contrast to HEMA, BSO did not induce a significant increase of the number of cells in S-phase compared with unexposed cells (34 2%) (Fig. 6). A
B
We have compared the effects of the methacrylate monomer, HEMA, with the effects of the glutathione synthase inhibitor, BSO, on a human bronchial epithelial cell line (BEAS-2B). It has previously been found that methacrylate monomers induce glutathione depletion by adduct formation (14, 23). The glutathione depletion is assumed to be the cause of the reported ROS increase in cells exposed to methacrylate (14, 24). Increased intracellular ROS is potentially toxic to cells, and we wanted to study whether glutathione depletion by BSO had similar effects on cells as exposure to HEMA. We found a similar degree of glutathione depletion in cells exposed to either HEMA or BSO. In addition, increased levels of ROS were observed under the two exposure conditions. Based on these findings, we evaluated the use of BSO as an appropriate model to study the cytotoxic effects of glutathione depletion. BEAS-2B cells were employed after reports on HEMA detected in the air at dental surgeries. This implies exposure of bronchial cells, of both staff and patients, to HEMA (3, 5). Significantly decreased viability and increased numbers of apoptotic cells were observed for BEAS-2B cells exposed to HEMA for 24 h, whereas no change was observed after exposure to BSO. Exposure both to HEMA and to BSO resulted in glutathione depletion and increased ROS. These results do not support the hypothesis that HEMA exerts its toxicity by lowering the cellular glutathione level, at least not exclusively. A slight delay in cellular glutathione depletion and ROS formation in BSO-treated cells could result in a delayed onset of apoptosis and explain the difference measured after 24 h of exposure. Arguing this, we did not observe increased cell death when extending the BSO exposure time to 48 h (data not shown). The previously reported lack of correlation between glutathione depletion and cell viability after exposure to different methacrylate monomers (23) also suggests additional toxic mechanisms of at least some methacrylates. The absence of a direct link between glutathione level and cell death is further supported by a study in which no cell death was observed in a mouse macrophage cell
C
Fig. 5. Cell-cycle histograms of one representative experiment. Unexposed cells (A) and cells exposed to 2-hydroxyethyl methacrylate (HEMA) (5 mM) (B) or to L-buthioninesulfoximine (BSO) (10 lM) (C) for 24 h. Data shown are one representative of at least four experiments.
Fig. 6. Percentage of cells in S-phase, without exposure or after 24 h of exposure to 2-hydroxyethyl methacrylate (HEMA) (5 mM) or to L-buthioninesulfoximine (BSO) (10 lM). The data are mean SD of at least four separate experiments. *Statistical difference between unexposed and HEMA-exposed cells; #statistical difference between HEMAexposed and BSO-exposed cells (P < 0.05). The data are calculated from the histograms shown in Fig 5.
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line treated with BSO for 24 h (24). This study, however, showed increased cell death after exposure to BSO and HEMA in combination compared with HEMA alone. Taken together, glutathione depletion alone does not explain the toxic response to HEMA, but low cellular levels of glutathione could make the cells more susceptible to HEMA exposure. Glutathione–HEMA adduct formation may be an advantageous event through lowering the concentration of free intracellular HEMA. This is in agreement with the current understanding of the phase II metabolism of xenobiotics, where, for example, glutathione conjugation of xenobiotics is considered a beneficial detoxification reaction (18). Similarly to the different effects of HEMA and BSO on cell death, only HEMA reduced proliferation of BEAS2B cells. Both microscopic observations and flow cytometric measurements indicated cell-growth disturbance after exposure to HEMA, but failed to detect a difference in cell growth between BSO-treated cells and the control. The visual appearance of the DNA histograms also differed markedly. 2-Hydroxyethyl methacrylate (HEMA) induced a ‘tail’ in the transition phase between the G1 and S-phases which was not observed in BSO-treated cells or the control cells. Furthermore, the proportion of cells in S-phase was significantly increased in HEMAexposed cells, but not in BSO-treated cells. The role of glutathione in cell proliferation is reported to be of importance to the control of tumour growth, and glutathione depletion is a desired effect of several anticancer drugs (25). However, the different pattern of cell growth and cell death after exposure to HEMA and BSO argues that glutathione depletion alone is the cause of these HEMA-induced changes. An alternative mechanism for the observed cell cycle changes in HEMA-exposed cells could be the binding of HEMA to nucleophilic groups in DNA, thereby forming a HEMA–DNA adduct (15, 26). The finding, of KRIFKA et al. (24), of increased cytotoxity in cells incubated with both HEMA and BSO, could be in accordance with a theory of glutathione serving as a detoxifying agent, binding free HEMA in the cell and thereby preventing HEMA–DNA adduct formation. In the present study we hypothesized that HEMA induces cell-proliferation disturbances and cell toxicity through glutathione depletion and subsequent ROS formation. In contrast to HEMA, exposure to BSO, which leads to a similar depletion of glutathione and elevation of ROS, did not induce cell toxicity and cell cycle disturbances. This indicates that HEMA-induced cell toxicity is not caused by glutathione depletion and ROS formation alone.
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17. Acknowledgements – The authors wish to thank Professor Jon E. Dahl for critical revision of the manuscript. We are also thankful to Dr John E. Tibballs for useful discussion and comments on the manuscript. Conflicts of interest – The authors have no conflicts to declare.
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