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ABSTRACT
Salivary conditioning films (SCF) play an important role in protecting the enamel surface against abrasion and erosion by providing better lubrication and the formation of a barrier against diffusion of acidic components of beverages toward and of Ca2+ and PO43‐ ions out of the enamel surface. However, the physical mechanism behind this dual protection offered by SCFs is not well understood.
Here, we use a SnF2 containing mouthrinse to demonstrate the importance of structural and glycosylation changes in SCFs, as induced by Sn2+ ions in the protection of enamel surfaces against erosion and abrasion. Quartz crystal microbalance with dissipation monitoring (QCM‐D) showed that SCFs became rigid after exposure to a SnF2 containing mouthrinse, which we attributed to cross‐linking of adsorbed proteins by Sn2+ ions. During renewed exposure to saliva, the SnF2 treated SCF recruited more salivary proteins, thereby increasing the adsorbed mass and degree of glycosylation in the SCF, as determined from QCM‐D and X‐ray photoelectron spectroscopy, respectively. The renewed adsorbed film on a SnF2 treated SCF provided a lower friction than when formed on an untreated SCF. Moreover, such rigid, more heavily glycosylated and lubricious SCFs yielded a lower calcium loss during exposure to a citric acid solution than untreated SCFs. Therewith, this is the first study to demonstrate physical changes in SCFs due to Sn2+ adsorption that can be related to the control of erosion and abrasion of enamel surfaces in vitro.
INTRODUCTION
Salivary conditioning films (SCF) cover all oral surfaces and have been shown to reduce enamel abrasion [1] through lubrication as well as to prevent enamel erosion by acting as a diffusion barrier for acids from beverages toward and of Ca2+ and PO43‐ ions out of enamel surface [2, 3]. Glycosylated mucins in adsorbed SCFs play an important role in preventing erosion [4] and are known to improve lubrication [5, 6] This is supported by observations that xerostomic patients suffer from a lower secretion of mucins and have an increased probability of enamel demineralisation [7] .
In preventive dentistry, scientific efforts to prevent enamel erosion focus on identifying optimal fluoride formulations. Common formulations used in caries prevention are based on sodium fluoride, but these have only limited effects on erosion [8‐10]. In contrast, solutions of stannous fluoride (SnF2), titaniumtetrafluoride (TiF4) and hydrofluoric acid (HF) have been shown to be effective in preventing erosion as determined in vitro and in situ [11, 12].
Moreover, SnF2 has been described as the “most promising erosion protective agent” on the market [13].
The physical changes in SCFs after treatment with SnF2 solutions responsible for the protective nature of SCF against erosion and abrasion are largely unknown, although glycosylation and structure of SCFs can be chemically (e.g. by toothpaste and mouthrinse components) [14] and mechanically (e.g. by toothbrushing) [15]
perturbed. Therefore, the aim of this study was to demonstrate the importance of structural and glycosylation changes in SCFs, as induced by Sn2+ ions in the protection of enamel surfaces against erosion and abrasion.
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MATERIALS AND METHODS Saliva collection
Human whole saliva from 20 volunteers (10 male and 10 female) of both genders was collected into ice‐cooled beakers after stimulation by chewing Parafilm® and then pooled, centrifuged, dialyzed, and lyophilized for storage. Prior to lyophilization, phenylmethylsulfonylfluoride was added to a final concentration of 1 mM as a protease inhibitor in order to reduce protein breakdown and preserve high‐molecular weight mucins. Note that recently it has been shown that freeze‐
thawing does not alter saliva which has been stored at ‐20oC and ‐80oC for a period of 6 months [16]. For experiments, lyophilized saliva was reconstituted at 1.5 mg ml‐1 in buffer (50 mM potassium chloride, 2 mM potassium phosphate, 1 mM calcium chloride, pH 6.8). This reconstituted human whole saliva will be referred to as “saliva”. Volunteers gave their informed consent to saliva donation, in agreement with the guidelines set out by the Medical Ethical Committee at the University Medical Center Groningen, Groningen, The Netherlands.
Quartz crystal microbalance with dissipation monitoring
Kinetics of adsorption of salivary proteins and structural softness of adsorbed SCFs were determined using a QCM‐D device, model Q‐sense E4 (Q‐sense, Gothenburg, Sweden). Gold (Au) plated AT‐cut quartz crystals with a sensitivity constant of 17.7 ng cm‐2 for a 5 MHz sensor crystal, were used as substrata. The Au‐plated crystals were cleaned 2 h prior to use by 10 min UV/ozone treatment, followed by immersion into a 3:1:1 mixture of ultrapure water, NH3 and H2O2 at 70°C for 10 min, drying with N2 and another UV/ozone treatment. The QCM‐D chamber is disc‐shaped with a volume of approximately 40 l and a diameter of 11.1 mm with the inlet and outlet facing the crystal surface. The chamber was first perfused with buffer by a peristaltic pump (Ismatec SA, Glattbrugg, Switzerland).
When stable base lines for both frequency ∆f and dissipation ∆D were achieved, saliva was introduced. Experiments were conducted at 20°C, at a flow rate of 50
l min‐1, corresponding with a shear rate of 3 s‐1. Saliva was flown through the QCM‐D chamber for 2 h to form a SCF, followed by 2 min exposure to buffer or 0.63% w/v of SnF2 containing mouthrinse (3M ESPE, Dental Products, Minnesota, USA) and renewed flow of saliva was initiated for 2 h. After each step, a buffer rinse was applied for 10 min. The structural softness of the SCF was determined by their ∆D3/∆f3 ratio recorded in QCM‐D. Higher changes in ∆f3 toward more negative values were interpreted to indicate an increase in hydrated mass of the SCF, while a higher ∆D3/∆f3 ratio indicated an increase in structural softness.
Atomic force microscopy
Friction force, surface topography and repulsive force toward a colloidal probe AFM [17] were measured in presence of buffer with an AFM (Nanoscope IV Dimensiontm 3100) equipped with a Dimension Hybrid XYZ SPM scanner head (Veeco, New York, USA) on the different adsorbed SCFs. To this end, rectangular, tipless cantilevers (length (l), width (w) and thickness (t) of 300, 35 and 1 μm, respectively) with a stiffness of 0.05 N m‐1 were calibrated for their exact torsional and normal stiffness using AFM Tune IT v2.5 software [18]. The normal stiffness (Kn) was in the range of 0.01 to 0.04 N m‐1, while the torsional stiffness (Kt) was in the range of 2 to 4 x 10‐9 N‐m rad‐1.
Subsequently, a silica particle of 4.74 μm diameter (d) (Bangs laboratories, Fishers, IN, USA) was glued to a cantilever with an epoxy glue (Pattex, Brussels, Belgium) using a micromanipulator (Narishige group, Tokyo, Japan) to prepare a colloidal probe. The deflection sensitivity (α) of the colloidal probe was recorded
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at a constant compliance with bare Au‐plated crystal in buffer to calculate the applied normal force (Fn) using
where ∆Vn is the voltage output from the AFM photodiode due to normal deflection of the colloidal probe. The torsional stiffness and geometrical parameters of the colloidal probe were used to calculate the friction force (Ff) [18, 19] according to
the AFM and ΔVL corresponds to the voltage output from the AFM photodiode due to lateral deflection of the colloidal probe. Lateral deflection was observed at a scanning angle of 90 degrees over a scan area of 5 x 5 µm2 and at a scanning frequency of 1 Hz. The scanning angle, distance and frequency were kept constant throughout all friction force measurements.
The colloidal probe was incrementally loaded and unloaded up to a maximal normal force of 32 nN in buffer. At each normal force, 10 friction loops were recorded to yield the average friction force.
Repulsive force‐distance curves measured in the contact mode between a colloidal probe and the films were obtained at a threshold force of 5 nN and at an approach and retraction velocity of 10 µm s‐1. All surface topography imaging by
colloidal probe was performed at 5 nN normal force, and the surface height was calculated from these surface topography images.
X‐ray photoelectron spectroscopy
The degree of glycosylation of the adsorbed salivary films was determined by using XPS (S‐probe, Surface Science Instruments, Mountain View, CA, USA). Films adsorbed on Au‐plated quartz crystals as removed from the QCM‐D chamber, were dried in the pre‐vacuum chamber of the XPS, and then subjected to a vacuum of 10‐7 Pa. X‐rays (10 kV, 22 mA), at a spot size of 250 1000 m, were produced using an aluminum anode. Scans of the overall spectrum in the binding energy range of 1‐1100 eV were made at low resolution (pass energy 150 eV). The area under each peak was used to yield elemental surface concentrations for C, O, N, Sn, F and Au after correction with sensitivity factors provided by the manufacturer. The O1S peak was split into three components, i.e. for oxygen involved in amide groups (C=O‐N; 531.3 eV), carboxyl groups (C‐O‐H; 532.7 eV) and oxygen arising from the Au‐plated quartz crystal. Accordingly, the fraction of the O1s peak at 532.7 eV (%O532.7) was used to calculate the amount of oxygen involved in glycosylated moieties (%Oglyco)
%Oglyco = %O532.7 * %Ototal (4)
where %Ototal is the total percentage of oxygen.
Ex vivo erosion study
In this study, a 0.63% w/v SnF2 containing mouthrinse (3M PerioMed Oral Rinse, 3M ESPE, St Paul, MN, USA) was used and results compared with a control (water). A sintered hydroxyapatite (HAP) disc (Himed, Old Bethesda, NY, USA;
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batch #100406) was placed in the buccal sulcus of the lower jaw of a human volunteer in close proximity to the first molar at 9.00 am. Eating, drinking, brushing and smoking were not allowed from 1 h before insertion until removal of the samples from the oral cavity. After 1 h of wearing the disc, volunteers were asked to rinse with the SnF2 containing mouthrinse or water for 2 min and the HAP discs were left in the oral cavity for another 1 h. Subsequently, the discs were removed and ex vivo exposed for 2 min to 3 ml of an erosive solution (50 mM citric acid, pH 3.0) under agitation (100 rpm) and afterwards rinsed with 2 ml of demineralised water. Calcium loss into the acid solution was measured using atomic absorption spectroscopy, as described previously [20]. Three volunteers, two males and one female, participated in this part of the study, after giving their informed consent under the approval by the University Medical Center Groningen Investigators Research Board (UMCG IRB #2008109).
RESULTS
Salivary proteins adsorbed to Au‐plated crystal surfaces within seconds after exposure to saliva and the Δf3 decreased within minutes to ‐90 Hz. Simultaneous ΔD3 increased to 12 x 10‐6. The frequency and dissipation remained constant during 120 min of saliva flow (see Fig. 1). A 15 min rinse with buffer yielded a slight desorption of loosely adsorbed salivary proteins, indicated by an increase in Δf3 and decrease in ΔD3. Subsequent buffer exposure caused a further small change in the Δf3 and the ΔD3 (Fig. 1A), whereas 2 min of exposure to a SnF2 containing mouthrinse decreased Δf3 to ‐185 Hz and increased ΔD3 up to 45 x 10‐6 (Fig. 1B), indicating adsorption and of SnF2. Removal of the mouthrinse by a 10 min rinse with buffer rinsing increased Δf3 to ‐110 Hz and decreased ΔD3 to 9 x 10‐6. After renewed exposure of buffer treated SCFs to saliva for 120 min, only a small decrease in Δf3 and increase in ΔD3 was observed, whereas SnF2 treated
SCFs showed a much larger decrease in the Δf3 down to ‐145 Hz and much larger increase in ΔD3 up to 18 x 10‐6 upon renewed salivary protein adsorption.
Figure 1 Influence of SnF2 on adsorbed SCFs and adsorption of salivary proteins during renewed flow of saliva.
Example of the QCM‐D response to salivary protein adsorption on Au‐plated crystal surfaces, chemical perturbation and continued adsorption of salivary proteins as a function of time, expressed as changes in third harmonic frequency (∆f3, thick line) and dissipation (∆D3, thin line). Perturbation is due to exposure of buffer (A) or a SnF2 containing mouthrinse (B). The QCM‐D chamber was first perfused with saliva, after which a buffer rinse was applied in all cases, followed
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by 2 min exposure by buffer or SnF2, intermediate buffer rinsing for 15 min and continued perfusion of the chamber with saliva.
The structural softness of the SCF after exposure to a SnF2 containing mouthrinse decreased as compared with the softness after buffer treatment (Fig. 2A). After renewed adsorption of salivary proteins, differences in structural softness between SCFs treated with buffer or a SnF2 containing mouthrinse disappeared (Fig. 2A). Small amounts of Sn and F were measured in SCFs treated with the SnF2 containing mouthrinse, that were absent in SCFs exposed to buffer only (Table 1).
Moreover, the degree of glycosylation in SCFs formed on the SCF treated with the SnF2 containing mouthrinse was higher than in SCFs formed on SCFs treated with buffer (see Fig. 2B).
Au‐plated crystal surfaces were extremely smooth (Fig. 3A), whereas SCFs were characterized by adsorbed, globular structures (Figs. 3B, C). Protein globules on SCFs formed on buffer treated films were approximately 20 nm in height, with a width of 500 nm, whereas the SCF formed after treatment with a SnF2 containing mouthrinse possessed larger protein globules with a height and width of 38 nm and 900 nm, respectively (see Fig. 3D). The SCF formed after treatment with a SnF2 containing mouthrinse furthermore exerted repulsive forces to the colloidal probe over longer separation distances than SCFs formed on a buffer treated SCF (Fig. 3E).
Figure 2 Influence of SnF2 on structure and glycosylation of SCFs.
(A) structural softness of SCFs immediately after exposure to buffer or a SnF2 containing mouthrinse and after renewed SCFs formation..
(B) degree of glycosylation of SCFs formed on SCFs treated with buffer or a SnF2 containing mouthrinse.
Error bars represent the standard deviations over three independent experiments. *Statistically significant (p < 0.05, two tailed Student t‐test) differences between buffer and SnF2 treatment.
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Table 1 Elemental composition on SCFs formed on SCF exposed to buffer or a SnF2
containing mouthrinse. ± indicates standard deviation over three separate experiments.
C 62.9 ± 2.9 55.3 ± 0.3
O 22.6 ± 2.0 23.1 ± 1.3
N 14.3 ± 1.2 12.0 ± 0.4
force, indicating that friction was independent of the normal force applied (Fig.
4A). In contrast, friction forces on SCFs formed on films treated with a SnF2 containing mouthrinse were lower corresponding with a coefficient of friction of 0.07.
Ex vivo calcium loss after citric acid exposure in HAP discs worn in human volunteers was two‐fold lower when in vivo SCFs were treated with a SnF2 containing mouthrinse than with water (see Fig. 4B).
Figure 3 Surface topography (5 µm x 5 µm) and repulsive forces of SCFs exposed to buffer or SnF2 using colloidal probe AFM.
(A) bare Au‐plated QCM crystal
(B) SCF formed on SCF treated with buffer
(C) SCF formed on SCF treated with a SnF2 containing mouthrinse
(D) height as a function of the globular structures found on the different SCFs.
Black line, SCF treated with buffer, brown line, SCF treated with SnF2 and orange line, Au‐plated quartz crystal.
(E) example of the repulsive force as a function of separation distance between the colloidal probe and the different SCFs.
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Figure 4 Lubrication and erosion protection provided SCFs exposed to buffer or a SnF2.
(A) friction force as a function of increasing (closed symbols) and decreasing (open symbols) normal forces on SCFs formed on SCFs treated with buffer or a SnF2 containing mouthrinse. Error bars represent standard deviation over 9 experiments.
(B) ex vivo calcium loss due to citric acid exposure of HAP discs, worn by human volunteers after use of a SnF2 containing mouthrinse or control (water). Error bars represent standard deviations over three experiments in each volunteer.
Experiments in each volunteer were done in triplicate.
*Statistically significant (p < 0.05, two tailed Student t‐test) differences in properties of the buffer with respect to SnF2.
DISCUSSION
Abrasion and erosion are mechanical and chemical wear phenomena that eventually result in loss of enamel and dentin. This study shows that treatment of SCFs with a SnF2 containing mouthrinse caused the SCF to become more rigid, while furthermore SnF2 treated films recruited more glycosylated salivary proteins during renewed exposure to saliva, giving rise to long range repulsion of a colloidal probe and reduced friction. Reduced friction will be responsible for protection of the enamel surface against abrasion, while it was experimentally determined that SCFs formed on SnF2 treated films were better protected against erosion than in absence of SnF2 treatment.
Glycosylated, high‐molecular weight mucins adsorb in loops and trains to a surface, while smaller proteins, like proline‐rich proteins, histatins, lysozymes, amylases may be found underneath these loops and between the trains, as illustrated in Fig. 5A. Here we suggest a model for the interaction of Sn2+ ions with SCFs and the continued interaction of such films with salivary proteins. Sn2+
ions cause cross‐linking of negatively charged proteins, including adsorbed glycosylated mucins, yielding a rigid base layer (see in Fig. 5B). Not all Sn2+ will be involved in cross‐linking, yielding an initially less negatively charged outer surface of the film, that interacts with glycosylated mucins during renewed exposure to saliva and recruits them to form a soft outer surface of the final film (see Fig. 5C).
This results in an increased hydrated mass, as determined by QCM‐D with a higher degree of glycosylation in the SCF, as determined by XPS. A higher degree of glycosylation in the SCF reduces friction [5, 6], while increased rigidity adds stability in the SCF to withstand higher normal and shear forces. Also, SCFs with
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more hydration mass and higher glycosylation may decrease the interaction between citric acid and the HAP surface through increased diffusion time [2, 3].
Although it is known that glycosylated mucins can provide protection against demineralisation after exposure to citric acid [4], our results are the first to demonstrate that this protection is due to a combination of structural and compositional features of the SCF.
Figure 5 Architecture of SCFs after exposure to SnF2 and renewed adhesion of salivary proteins.
(A) adsorbed SCF, showing glycosylated mucins adsorbed in loops and trains over a layer of adsorbed, densely packed low‐molecular weight proteins, including proline‐rich proteins, histatins and lysozymes.
(B) SCFs after adsorption of SnF2. The Sn2+ ions diffuse into the SCFs and cross‐link negatively charged, glycosylated mucins, causing collapse of the glycosylated structure.
(C) SCFs with adsorbed Sn2+ ions and after renewed adsorption of salivary proteins. Reduced electrostatic repulsion due to the presence of Sn2+ ions residing on the SCF allow recruitment of additional glycosylated mucins to form a soft mucinous layer over a rigid, cross‐linked SCF.
In summary, we show that Sn2+ triggers the structure in SCF formed after renewed flow of saliva over SnF2 treated SCF into a rigid base layer and enhanced hydrated mass due to recruitment of glycosylated mucins over this rigid base layer. The synergy between the rigidity and hydrated overlayer with glycosylated mucins provides lubrication stability to improve abrasion resistance as well as a stronger diffusion barrier to citric acids to improve protection against erosion in the SCF.
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