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Azerbaycan Saytlari

 »  Home  »  Endodontic Articles 1  »  In vitro cytotoxicity of a composite resin and compomer
In vitro cytotoxicity of a composite resin and compomer
Introduction - Materials and methods.

Many new dental restorative materials have entered the market in recent years. It is claimed these products have many advantages, such as fluoride release and improved adhesion to enamel and dentine. However, in many cases, the product is released onto the market without prior independent evaluation. In this context, cytotoxicity testing is an assessment of the probability that a material will be hazardous to the body, particularly with respect to the material’s potential to cause pulpal problems (Wataha et al . 1994).
In the past, many varieties of restorative materials have been tested and found to be hazardous. Kawahara et al . (1968) found that silicate and zinc phosphate cements were toxic to cultured cells, particularly early in the setting process. Meryon & Riches (1982) subsequently reported that composite resin materials were cytotoxic to fibroblasts and macrophages and Muller et al . (1990) found that some glass ionomer cements were toxic to primary rabbit pulpal fibroblasts.
In addition to problems arising from leakage and bacterial contamination that particularly affected silicate cements (Powis et al . 1988), the direct toxicity of restorative materials largely results from the elution of compounds from the material. For example, Rathbun et al . (1991) studied the elution of the components of composite-resin restorative materials into various organic solvents. They found that the monomer Bis-GMA (bisglycidyldimethacrylate) was the main material eluted and they also identified a benzophenone UV stabiliser that did not participate in the photo-activated polymerization reaction. Furthermore, Geurtsen et al . (1998) showed that the elution of TEGDMA (triethylene glycol dimethacrylate) was one of the main causes for the cytotoxic reactions evoked by the light-cured glassionomer cements and compomers that they investigated. In fact, the National Institute of Occupational Safety and Health has classified TEGDMA as being irritating to various tissues (RTECS 1995).
It has been found that the greater the extent of the photo-initiated polymerization reaction, the fewer the monomers available to be leached (Ferracane 1994). This relationship was demonstrated by Rueggeberg & Craig (1988) for a light-cured composite consisting of the monomers Bis-GMA and TEGDMA, who showed that the degree of light-curing of this monomer mixture and the degree of conversion of carbon-carbon double bonds, to form the hardened material, was reduced by curing the composite through varying thickness of previously cured composite which reduced the light intensity.
Another significant factor to consider is whether or not the dentine provides protection for the pulp. Several features of dentine may influence the potential toxicity of a restorative material. The limited wetability of dentine limits the dissolution of the applied material, and the buffering capacity of the dentine hydroxyapaptite allows for the use of acid-base reaction materials and liquid acids (Pashley 1996). Furthermore, the presence of a smear layer may further reduce the permeability of dentine by acting as a diffusion barrier (Pashley et al . 1981). However, it can also be argued that the smear layer could harbour bacteria, which would then multiply beneath the applied restoration (Brannstrom 1996). Indeed Hanks et al . (1994), using a ‘split chamber device’, found that the diffusion of biological and synthetic materials through dentine was indirectly proportional to the dentine thickness. Similarly, Gerzina & Hume (1994) compared the diffusion of TEGDMA through dentine to the pulp space with its release directly into aqueous solution in vitro and reported that dentine provided an effective method of retarding the release of TEGDMA.
However, it is important to note that a restorative material which is hazardous in vitro may not necessarily be toxic in vivo , possibly due to biological barriers or insufficient time of contact between the offending restorations and susceptible tissues (Wataha et al . 1994). Therefore, in order to obtain an accurate risk assessment, the in vitro testing model must reflect the clinical situation as closely as possible. Two main strategies have been employed to date. The first is testing of the components of materials to cells in monolayer culture, constructing dose–response curves and then using this to estimate the cytotoxic potential of these components in vivo ; secondly, the use of barriers between the material and the cells to mimic barriers which might exist in vivo (Wataha et al . 1994). In the present study the second strategy was used, in which a dentine barrier was placed between cultured cells and the materials to be tested in vitro in order to simulate possible pulpal effects in the clinical situation.
All cells have the ability to die through activation of an internally encoded suicide programme. Once activated, this programme initiates a characteristic form of cell death called apoptosis (Thompson 1995). One of the essential features of apoptosis is that, although the membrane is thrown into massive convolutions (blebbing), it remains intact. The dead cells are rapidly removed through engulfment by macrophages and any leakage of their noxious contents and consequent inflammatory response is avoided (Cotter & Martin 1995). In contrast, necrosis is a pathological form of cell death that results from an overwhelming cellular injury. It is associated with an early loss of cell membrane integrity, resulting in leakage of cytoplasmic contents and the induction of an inflammatory response. Unlike apoptosis, where there is controlled auto-digestion of the cell, necrotic cells spill their contents out over the surrounding cells, thus spreading any toxic or infectious agent and resulting in a spreading wave of necrosis (Thompson 1995).
There have been few studies of the mode of cell death induced by dental restorative materials. In view of this, morphological and enzymatic assessments of the mode of cell death were performed after light curing samples of restorative materials for different time periods.

Materials and methods.

Maintenance and storage of cells.
Two different mammalian cell lines were obtained  from the European Collection of Animal Cell Cultures  (E.C.A.C.C.). The human endothelial cell line (ECV-304),  which has also been used in previous apoptosis studies  (Escargueil-Blanc  et al  . 1998), was chosen to simulate  the  in vivo  situation. The human promyelocytic leukaemia  cell line (HL-60) was the model used for apoptosis studies  ( Jarvis  et al  . 1994). 
The ECV-304 adherent cells were grown in medium  199 supplemented with foetal calf serum (FCS, 10%  v  /  v  ),  l  -glutamine (2 m mol L  1  ), penicillin (100 U mL  –1  ) and  streptomycin (100  g mL  1  ). The nonadherent HL-60  cells were grown in RPMI medium that had been supplemented  with FCS (20%  v  /  v  ),  l  -glutamine (2 m mol L  1  ),  penicillin (100 U mL  –1  ) streptomycin (100  g mL  1  ),  and amphotericin B solution (2%). The stock cultures were maintained at 37  C in a humidified atmosphere  (Forma CO  2  -water-jacketed incubator, Forma Scientific,  OH, USA) with 5% CO  2  in 95% air, in 75 cm  2  tissue  culture flasks (Grenier® Laboratories, Frickenhausen,  Germany). The ECV-304 cells were passaged twice weekly,  upon reaching confluence and re-seeded at a cell density  of 1.0  10  5  cells mL  –1  . The HL-60 cells were passaged  three times weekly and re-seeded at a cell density of  1.5  10  5  cells mL  –1  .

Moulds and dentine preparation.
Human maxillary canine teeth (10), which had been  extracted for orthodontic reasons, were obtained with  patient consent (from the Dublin Dental Hospital) and  stored in physiological saline until required. Both the  crown and root portions of the teeth were embedded  whole in clear methyl methacrylate resin under vacuum.  The crown portion was later sectioned into 70-micron  sections, using a hard tissue sectioner (Leica AG, Bensheim,  Germany) under water spray. The dentine of the root  portion was not used experimentally. 
To ensure reproducible dimensions of sample materials  employed – Spectrum® composite resin (Dentsply, Surrey,  UK) and Dyract® AP compomer (Dentsply) – a custom  mould was created so that material discs of 3 mm diameter  and 2 mm thickness were produced. The mould was  constructed in additional vinyl polysiloxane impression  material (3M Express®, 3M Dental Products, MN, USA). 
Each dentine section was cut into a diameter similar to  that of a well from a 96 well plate and then etched for 30 s  using 37% phosphoric acid gel, washed for 30 s in water,  dried and then placed on a sterile glass block. Primer  (Prime & Bond®, OSB Bond®, Dentsply) was dispensed  into a plastic watch-glass and applied with a brush to the  dentine section to form a thin layer and then light-cured  for 10 s using a Visilux 2 visible-light curing unit (3M  Dental Products). 
A second layer of primer was applied and light-cured,  as before, for 10 s. The mould was then placed centrally on  the dentine section and the sample material (Spectrum®,  Dyract® AP) was dispensed into the mould using a  preloaded compule tip gun. The material was then fully  cured for 40 s. 
The mould was removed and a layer of soft red wax  was placed around three sides of the restorative material  attached to the centre of the dentine section, the fourth  remained free of wax to allow for the passage of air and  CO  2  to the underlying cells. The samples were inserted  into the plate well so that the dentine, in contact with the  culture medium, formed a physical barrier between the  restorative material and the cells. This was achieved via  the wax seal, which was mouldable at room temperature,  allowing the sample to be lowered into position. 
The above system was used for studies of cell viability  by the MTT (3,-[4,5-dimethylthiazol-2-yl]-2,5-diphenyl  tetrazolium bromide) reduction and the LDH (lactate  dehydrogenase) release procedures with ECV-304 cells,  as described below. 
For studies with HL-60 cells no dentine sections were  used. A custom mould was created, as described above, so  that material samples of 3 mm diameter and 1 mm thickness  were produced, and the mould was placed directly  onto the sterile glass block. The restorative material  (Spectrum® or Dyract® AP) was dispensed, using the  preloaded compule gun, into the well of the mould and  light-cured for 1 s (partially cured), 4 s (until the restoration  was solid to probing) or 40 s (fully cured). The mould  was then removed and the materials were stored in sterile  containers in the dark until required.

Cell viability assays.
For both the LDH and MTT assays, ECV-304 cells were  seeded, at a density of 1.0  10  5  cells mL  1  , into three 96-  well microplates and incubated at 37  C in 5% CO  2  , 95%  air and left to adhere overnight. A fully cured sample of  Dyract® AP or Spectrum® was inserted into each well so  that the dentine disc with the attached material was in  direct contact with the culture medium, leaving a space  of approximately 5 mm between the sample and cell  population. In addition, the samples were held in place by  a wax seal. Controls of wax and dentine alone (70  m  thick, surrounded by wax) were added in triplicate to  each plate. As before, controls were in contact with the  culture medium. Each plate was further incubated at  37  C in 5% CO  2  , 95% air over a period of 1–3 days. 
Cell viability was determined using the MTT assay  (Mosmann 1983). Upon incubation with viable cells, the  tetrazolium ring of MTT (pale yellow) is cleaved by cellular  dehydrogenases and at sites in the mitochondrial electrontransport  system (Vistica  et al  . 1991) to yield a purple  formazan product. MTT solution (0.5 mg mL  –1  per well)  was added to each plate and they were incubated for  2.5 h at 37  C in the dark. The formazan crystals were  solubilized with DMSO (200  L, dimethylsulfoxide) and  the absorbance determined at  = 595 nm using an  ELISA plate reader (Thermo  max  Microplate Reader, Molecular  Devices, CA, USA). 
Cell lysis was determined using an LDH assay kit under  conditions described by the manufacturer (Promega  Corporation, Madison, WI, USA). The essence of this assay is that LDH is a cytosolic enzyme that is released  upon cell lysis. The released LDH is measured in a coupled  assay that results in the conversion of a tetrazolium  salt into a red formazan product. The amount of colour  formed, expressed as a percentage of the total LDH activity,  is proportional to the number of lysed cells. Total LDH  was determined after lysis of untreated cells using the  detergent, Triton® X-100 (9%  v  /  v  , Korzeniewski &  Callewaert 1983). Following sample incubation, 50  L  aliquots were transferred from all wells to a fresh 96-well  plate. To each well 50  L of the reconstituted substrate  mix was added and incubated for 30 min at room  temperature. Stop solution (50  L) was added to each  well and the absorbance was measured within 1 h at  = 490 nm using an ELISA plate reader. Both the LDH  and MTT assays were carried out in triplicate and  repeated in three separate experiments.

Morphological staining of cells.
HL-60 cells were seeded, at a density of 5.0  10  5  cells mL,  into a sterile 96-well microplate and incubated at 37  C in  a humidified atmosphere (Forma CO  2  -water-jacketed  incubator) with 5% CO  2  in 95% air overnight. The following  day 20 fully cured or partially cured samples (one  sample per well) of Dyract® AP or Spectrum® were added  to the plates, whilst the control remained untreated and  the cells were further incubated over a period of 2–12 h. 
Every 2 h one of each sample type was removed and  the cells (150  L; 5.0  10  5  cells mL  1  ) were loaded into  the prepared cytospin cup and slide set-up and cytocentrifuged.  The slides (Shandon Corporation, PA, USA)  were then air-dried at room temperature before being  differentially stained using the RAPI-DIFF II stain pack  (Diachem International Ltd, Lancashire, UK). The slides  were allowed to air-dry and then viewed by light microscopy  under  40 magnification. Apoptotic cells were  characterized by cell shrinkage, membrane blebbing,  nuclear condensation and fragmentation, whilst necrotic  cells were identified by cell swelling and loss of cell  membrane. The percentage of apoptotic and necrotic  cells was determined by taking counts of 100 cells in  three separate fields of view and correlating this to the  total number of cells present. This procedure was carried  out in triplicate and repeated three times for the fully and  partially cured materials.

Determination of caspase-3 activity.
Caspases, a family of cysteine proteases, are thought  to play a vital role in the disassembly of the cell during  apoptosis. These enzymes ensure the ultimate demise of  the cell by preventing DNA repair, condensing the cytoplasm  and inhibiting DNA synthesis and splicing (Villa  et al  . 1997). Caspase-3 activity was determined using  the fluorometric peptide substrate Asp-Glu-Val-Asp-  Aminomethyl Coumarin (DEVD-AMC, Alexis Corporation,  Nottingham, UK) which contains the specific aspartic  acid cleavage site for this peptidase. The caspase-3 activity  was quantified as the amount of the fluorescent leaving  group (AMC) released per min per mg of protein, according  to the method of Kaushal  et al  . (1997). 
The HL-60 cells were seeded at a density of 5.0  10  5  cells mL  1  into 96-well microplates and incubated at  37  C in 5% CO  2  , 95% air for 24 h. Thirty samples each of  Dyract® AP and Spectrum® that had been cured for  either one, four or 40 s were added to one of the 96-well  plates and incubated in a humidified atmosphere at 37  C  in 5% CO  2  , 95% air for a further 10 h. The control consisted  of untreated cells. For the etoposide treatment, 4 h  prior to harvesting the cells, following completion of  sample incubation, etoposide (50  mol L  1  ) was added  to 5 mL of HL-60 cells (5.0  10  5  cells mL  1  ). This was  used as a positive control, because etoposide is a DNAdamaging  agent that induces the structural changes  associated with apoptosis (Ritke  et al  . 1994). 
Following sample incubation, the cells were pelleted by  centrifugation, washed in PBS, and recentrifuged. The  pellets were resuspended in 20 m mol  1  HEPES (N-  [2-hydroxyethyl] piperazine-N  -[2-ethanesulfonic acid),  pH 7.5, containing 10% sucrose, 0.1% CHAPS (3-[(3-  cholamidopropyl) dimethylammonio]-1-propanesulfonate),  0.1% NP40 (Nonidet P40), 1 m mol L  1  EDTA  (ethylenediaminetetraacetic acid), 1 m mol L  1  PMSF  (phenylmethylsulfonyl fluoride), 2 m mol L  1  DTT (DLdithiothreitol),  1  g mL  1  leupeptin and 1  g mL  1  pepstatin. 
The supernatants obtained after centrifugation were  used to determine the enzyme activity. A 50-  g sample of  protein from each set of samples was incubated with  100 m mol L  1  HEPES, pH 7.5 (containing 10% sucrose,  0.1% CHAPS, 10 m mol L  1  DTT) and 20  m DEVD-AMC  substrate in a total reaction volume of 3 mL. After  incubation for 60 min at 25  C, the fluorescence was  determined using a spectrophotofluorometer (Perkin-  Elmer Corporation, Ueberlingen, Germany) at an excitation  wavelength of 380 nm and an emission wavelength  of 460 nm. The amount of fluorescence from the cells  was correlated to the amount of liberated AMC using  an AMC calibration curve. Protein concentrations of  the samples were determined by the Bradford (1976)  method.

Statistical significance was determined by Student’s  t  -  test with a  P  -value of <0.05 being considered significant.