Introduction
Bacteria in the oral cavity consist of more than 700 bacterial species. In this niche, commensal and pathogenic bacteria co-exist and form biofilms [1]. The Infectious diseases in the oral cavity include dental caries, endodontitis and periodontitis. Dental caries is induced by Streptococcus mutans [2]. Periodontopathogens are classified as a red complex consisting of Porphyromonas gingivalis, Tannerella forsythia and Treponema denticola [3]. These pathogenic bacteria form biofilms according to an ordered case sequence [4]. Commensal bacteria bind on teeth and form the initial biofilm. The initial biofilm was developed to mature biofilm by exopolysaccharides as glucans which are secreted from S. mutans. S. mutans is crucial in the formation of cariogenic biofilms by abundant production of acid and glucan. P. gingivalis and T. forsythia also form biofilms with the commensal bacteria through Fusobacterium nucleatum in the gingival pocket [5,6]. The bacteria in biofilm are more complex compared to planktonic bacteria because they are in organized communication with each other and produce specific matrix molecules [4]. Therefore, biofilms are not easily controlled by chemical treatments. The effective bactericidal concentration of antibiotics for biofilm bacteria may be up to 100-1000 fold higher than that of planktonic or floating bacteria [7,8]. Nevertheless, the studies of virulences, antimicrobial agents and characterization about pathogens have conventionally been performed using planktonic bacteria. Control of pathogenic biofilms is essential for maintenance of human health and for prevention of disease.
Electrolyzed water is generated by an electric current that is passed through water using metal electrodes. Although the detailed mechanisms of electrolyzed water remain elusive, the electrolyzed water has varied characteristics depending on the metal electrodes, which include copper, silver, palladium and gold [9-11]. Electrolyzed water can be acid, basic and neutral. Electrolyzed water has been implicated in anti-cancer effects, oxidative stress and antibacterial activity [12-14]. Acidic electrolyzed water has bactericidal effect on Staphylococcus aureus and Escherichia coli [9,12]. Basic electrolyzed water affects oral hygiene including bactericidal activity, biofilm removal and bacterial growth inhibition [15]. Slightly acidic electrolyzed water removes bacterial biofilms on dental unit water systems [16]. However, the antimicrobial effect of electrolyzed water on biofilms of oral microbes has not been studied. This study analyzed and compared the antimicrobial activity of electrolyzed water using various electrodes on biofilms of oral microbes.
Materials and Methods
Oral microbes and cultivation
Streptococcus mutans ATCC 25175 was cultured using brain heart infusion (BHI) broth (BD Bioscience, Franklin Lakes, NJ, USA). Fusobacterium nucleatum ATCC 25586, and Porphyromonas gingivalis ATCC 33277 were cultivated using BHI broth supplemented with hemin (1 μ g/ml) and vitamin K (0.2 μg/ml) in anaerobic condition (5% H2, 10% CO2 and 85% N2) at 37℃. Tannerella forsythia ATCC 43037 was cultured modified in new oral spirochete (NOS) broth [17] at 37℃ anaerobically.
Production of various electrolyzed waters
Tap water (1 L) was subjected to electrolysis for 5 min with 24V of DC 400 mA using copper, silver or platinum electrode (cylindrical, 3 mm x 10 cm) in an whole tank undivided anode chamber and cathode chamber. The DC power supply was PowerPac basic (Bio-Rad, Hercules, CA, USA). pH level of the electrolyzed water was measured by an Orion 520A pH meter (Thermo Scientific, Beverly, MA, USA) using a pH electrode
Antimicrobial activity of the electrolyzed waters against oral microbes
S. mutans, P. gingivalis, and T. forsythia were cultivated in each specific medium for 36 h and counted by a Petroff- Hausser bacterial counting chamber (Hausser Scientific, Horsham, PA, USA). The bacterial density was adjusted to 1 x 107 cells/ml with BHI broth. One milliliter of each bacterial suspensions was harvested by centrifugation at 6,000 x g for 5 min 4℃ and washed twice with phosphate buffered saline (PBS, pH7.2). The bacterial pellet was treated with 1 ml of electrolyzed water for 30 sec or 1 min, and then immediately added to 1 ml of BHI broth. After diluting serially 10-fold to 106, 50 μl of each diluted suspension was spread on agar plate and incubated at 37℃ anaerobically until colonies could be counted. All procedure of antibacterial assay was performed in anaerobic chamber.
Biofilm formation of oral microbes
For biofilm formation of S. mutans, pooled saliva of 10 healthy donors was centrifuged at 9,000 x g for 10 min at 4℃, and then the supernatant was transferred to a new tube. The same volume of PBS was added. After filtration through 0.22 μm polyvinylidene fluoride (PVDF) membrane, the saliva sample was dispensed into 12-well polystyrene plate. The plate was dried at 40℃ and sterilized in an UV sterilizer. S. mutans was inoculated into BHI broth including 1% sucrose and incubated at 37℃ for 48 h anaerobically [2]. To form biofilm of P. gingivalis and T. forsythia, the protein of F. nucleatum was extracted by methanol-chloroform methods after disruption by sonicator (VCS-1300; SONICS, Newtown, CT. USA). The protein was coated on 12-well polystyrene plate. The plate was dried at 40℃ and sterilized using UV. P. gingivalis or T. forsythia were inoculated into BHI broth supplemented with hemin and vitamin K or modified NOS broth, respectively and incubated at 37℃ for 48 h in an anaerobic condition.
Antimicrobial activity of the electrolyzed water against biofilm of oral microbes
Biofilm was reacted with 2 ml of tap water and electrolyzed water for 30 sec or 1 min and the solution was immediately aspirated by vacuum pump. Biofilm of S. mutans, P. gingivalis and T. forsythia received 1 ml of unsupplemented BHI broth or BHI broth supplemented with hemin and vitamin K and modified NOS broth, respectively. Biofilms were mechanically disrupted with a scraper and transferred into a new 1.5 ml tube. The tubes were vortexed for 30 s (in case of S. mutans, glass beads were added in the tube). The biofilm suspensions were serially diluted from 10 to 108 with each specific medium. Fifty microliters of diluted biofilm suspension was spread on BHI agar, BHI agar supplemented with hemin and vitamin K, and modified NOS agar using triangle spreader, respectively. The bacteria were incubated at 37℃ for 72 h in an anaerobic condition. Colony forming units of each bacterium were counted.
Comparison of biofilm biomass and viable bacteria
Bacterial biofilms were treated with electrolyzed water and washed twice with PBS to remove planktonic bacteria. Each biofilm was stained with crystal violet and washed three times with PBS. Acetone-alcohol (10:90) solution was added and, the optical density of the supernatant was measured at a wavelength 590 nm by spectrophotometer to determine the biofilm biomass. To detect viable bacteria, electrolyzed water-treated S. mutans biofilms were observed by LSM 5 Pascal confocal laser scanning microscope (Carl-Zeiss, Oberkochen, Germany) using Z-stack scan from 0 to 30 after bacterial live/dead staining using LIVE/DEAD® BacLightTM kit (Invitrogen, Eugene, OR, USA). Briefly, live bacteria are stained by binding SYTO 9 on the bacterial membrane, and dead bacteria are stained by binding of propidium iodide (PI) to nucleic acid. SYTO 9 and PI fluoresce green and red, respectively. Image was visualized using Zeiss image program (Carl-Zeiss).
Statistical analysis
Differences between control and samples were analyzed by Krusical-wallis and Mann-Whitney test using SPSS 10 (SPSS Inc., Chicago, IL, USA). p-values less than 0.05 were considered statistically significant.
Results
Antimicrobial activity of the electrolyzed water against oral microbes
The typical pathogens of 3 species related with oral diseases such as caries, gingivitis and periodontitis were selected to investigate antimicrobial activity of the electrolyzed water. When planktonic pathogens were treated with various electrolyzed water for 30 sec or 1 min, the electrolyzed water using platinum electrode (EWP) exhibited antimicrobial activity against S. mutans, P. gingivalis and T. forsythia (Fig. 1). The electrolyzed water using copper electrode (EWC) and silver electrode (EWS) did not affect oral microbes.
Antimicrobial activity of the electrolyzed water against biofilms of oral microbes
Oral pathogens form biofilm in oral cavity and express specific genes related with resistance against antimicrobial agents via bacterial communication [8]. Therefore, after the bacteria were formed biofilm, the electrolyzed water was evaluated antimicrobial activity against the biofilm of oral microbes. Among the electrolyzed water, EWP showed strong antimicrobial activity against the biofilm of oral microbes (Fig. 2). The viability of S. mutans biofilm was decreased in treating with EWP for 1 min (Fig. 2A). Also, the viable count of P. gingivalis biofilm was slightly reduced by 1 min treatment with the EWS, and 30 sec or 1 min treatment with EWP. In case of T. forsythia biofilm, 30 sec or 1 min treatment with the EWP decreased the bacterial viability, as did not treatment with EWC and EWS (Fig. 2C).
Disruptive effect of the electrolyzed water on biofilm of oral microbes
The antimicrobial efficiency of the electrolyzed waters differed between biofilm and planktonic bacteria. Especially, EWP showed greater antimicrobial efficiency against biofilms than planktonic microbes for P. gingivalis and T. forsythia (Figs. 1B, 1C, 2B, and 2C). After treatment with the electrolyzed water, biofilm biomass was investigated. The EWC and the EWS did not disrupt T. forsythia biofilm (Fig. 3C), while EWP significantly reduced the biomass of P. gingivalis and T. forsythia biofilms (Figs. 3B and C). However, none of the electrolyzed waters disrupted S. mutans biofilm (Fig. 3A).
Antimicrobial activity of the electrolyzed water using platinum electrode against S. mutans biofilm.
EWP showed antibacterial activity against planktonic S. mutans but did not disrupt S. mutans biofilms. Bacteria in biofilm are phenotypically heterogeneous, unlike fungi. Therefore, the antibacterial activity of EWP was investigated by confocal laser scanning microscopy after bacterial live/dead staining. S. mutans biofilms were treated with tap water as control fluoresced green (Fig. 4A). When S. mutans biofilms were treated with EWP for various times, they tended to fluoresce mostly green with some yellow in treating EWP for 30 sec (Fig. 4B), and green, yellow and red color in 1 min treatment with EWP (Fig. 4C). Especially, the red fluorescence tended to be in the interior of biofilms, indicating that the antibacterial activity of EWP affected the interior of S. mutans biofilm (Fig. 4C).
Discussion
Oral microbes exist in planktonic and biofilm forms in oral cavity. Biofilms as dental plaque are associated with oral disease [2,4,8]. Etiologically, P. gingivalis, T. forsythia and T. denticola are closely associated with acute and chronic periodontitis [3]. Since S. mutans is an acidogenic and aciduric bacteria, they are considered as cariogenic bacteria [2]. For these reasons, the management of oral biofilm is important for prevention of oral disease and has been studied by various groups [2,18]. Electrolyzed water is water through which an electric current is passed using a metal electrode. This process alters physicochemical properties of the water [19]. Electrolyzed water has antibacterial activity against food [10]. Most electrolyzed water is usually generated by electrolysis of solution including a little sodium chloride in reparative chamber between the cathode and anode [10]. Since electrolyzed water is acidic or basic, its bactericidal effect has been tried to apply for food and equipment [19,20]. In this study, neutral pH electrolyzed water was generated using copper, silver, and platinum electrodes with relatively high conductivity in an unseparated chamber.
When the antimicrobial activity of the three types of electrolyzed water against planktonic oral microbes was examined, the EWP exhibited to have relatively strong antimicrobial activity. Some studies were suggested that the bactericidal effect of electrolyzed water has been correlated with free chlorine [21,22]. However, the EWP used in this study did not contain detectable free chlorine (data not shown). Therefore, the antimicrobial effect of electrolyzed water might be also involved other molecules, such as dissolved hydrogen and oxygen radicals, as well as free chlorine. The chemical reaction is H2O →1/2 O2 +2H+ + 2e-, 2Cl- → Cl2 +2e- and Cl2+H2O ↔ HClO + HCl at the anode, and H2O + 2e- → 1/2H2 + OH- at the cathode. However the electrolyzed water in this study had neutral pH. High conductivity of a palladium electrode compared to copper and silver electrodes may produce oxygen radicals (O2-), and reactive hydrogen (H) on the basis of high concentration of dissolved hydrogen. These ions may destroy the bacterial wall by binding surface components, and inhibit cytoplasmic signaling by irregular aggregation of cytoplasmic molecules. Transmission electron microscopy of bacterial morphology after treatment with electrolyzed water has revealed cell wall damage and irregular aggregation of cytoplasmic small granules compared to non-treated bacteria [23].
The antimicrobial effect of electrolyzed waters on biofilms of the same oral pathogens was investigated, since biofilms are classically more resistant to antibiotics [8]. The EWP had an antimicrobial effect on biofilm of oral microbes such as S. mutans, P. gingivalis and T. forsythia biofilms. Unexpectedly, the antimicrobial activity of electrolyzed water for biofilm of oral microbes was only slightly higher level or similar to the planktonic counterparts. As noted above, the biofilm is more resistant to antibiotics molecules than planktonic microbes. Therefore, the disruptive effect of electrolyzed water on biofilms was evaluated. Biomass of biofilms can be measured by staining with crystal violet. Of the electrolyzed waters, only the EWP changed biofilm biomass. The EWP decreased the biomass of P. gingivalis and T. forsythia biofilm. These results indicate that the EWP has antimicrobial activity against S. mutans and both antimicrobial and disruptive effect on P. gingivalis and T. forsythia. S. mutans, unlike P. gingivalis and T. forsythia, produces exopolysaccharide as glucan [24]. S. mutans can synthesize both water-soluble and -insoluble glucan using glucosyltransferase [2]. Glucans assist in the attachment and colonization of bacteria, and are important in the formation and development of biofilms [25]. Also, glucans are a protective barrier of biofilms and protect oral bacteria by inhibiting diffusion of antibiotics into biofilms [8]. The lack of S. mutans biofilm disruption by the EWP may reflect the presence of the exopolysaccharide. Thus, the antimicrobial activity of the EWP against S. mutans biofilm was analyzed by confocal laser scanning microscopy after fluorescence staining for live and dead bacteria. The EWP had bactericidal effect on interior of S. mutans biofilm. Bacteria in biofilms are physiologically heterogeneous, with bacteria in the interior and on the surface being metabolically inert and active, respectively [26]. The EWP may affect S. mutans in stationary state.
The present study demonstrates the antimicrobial activity of electrolyzed water against biofilm and planktonic oral pathogens. The EWP showed the highest level of antimicrobial activity against oral pathogens in the both conditions. The neutral electrolyzed water has the merit of easy and safe application for humans than chemical solutions; safety for human in drinking up to at least 1.2 L/day has been reported [27].
Electrolyzed water generated using a palladium electrode may have potential value as a gargle solution for prevention of oral diseases induce by pathogens and denture-related stomatitis.
Conflict of interest
The authors declare that they have no competing interest.
