
Dental units, which are essential devices in modern dentistry, contain dental unit waterlines (DUWLs) that allow water transfer and movement. Previous studies have demonstrated that water discharged from DUWLs is contaminated with high levels of microorganisms due to biofilm formation on the inner surfaces of DUWLs1-4). Various microorganisms, including bacteria, fungi, and protozoa, have been observed in the water discharged from DUWLs5-7). Among these microorganisms, bacteria are the most common, and diverse bacterial species have been detected in DUWLs7,8). Most were waterborne bacteria with low pathogenicity; however, several opportunistic pathogens, such as Pseudomonas aeruginosa and Legionella pneumophila were also observed. Thus, bacteria from DUWLs may cross-infect dentists and patients undergoing dental treatment, particularly those with low immunity9-11). In addition, mature biofilms in DUWLs can provide a favorable environment for the growth of other bacteria, including opportunistic pathogens3,12).
Biofilms are formed at several stages, including adhesion, microcolony formation, maturation, and dispersion. During the early stages of biofilm formation, adhesion between bacteria and other microorganisms and specific recognition between bacteria leads to coaggregation13). Therefore, coaggregation contributes to the formation and maturation of complex multispecies13-15). Coaggregation is a highly specific cell-to-cell mechanism by which genetically distinct bacteria recognize and adhere to each other and is mediated by specific cell surface polymers16,17). These polymers are composed of an adhesin (a protein) on one partner and a receptor (a saccharide-containing polymer) on the other16). Coaggregation is also mediated by protein-protein interactions and protein–saccharide interactions18). Coaggregation also enables interactions such as protection from adverse environmental conditions, cell-cell communication, and exchange of genetic information within the biofilm, providing metabolic benefits to coaggregating bacteria13,14,19,20). Therefore, a better understanding of the coaggregation of biofilm bacteria and their mechanisms can be utilized to formulate effective ways to inhibit the formation and removal of biofilms. Biofilms are present in a variety of environments, including DUWLs, oral cavities, water distribution systems, contact lens cases, and more13,15,21). Although coaggre-gation in dental plaque, an oral biofilm, has been extensively studied, little information is available regarding coaggregation in other environments14,15,17,19,20). Coaggregation between strains isolated from water distribution systems and contact lens cases has been shown, and in the case of water distribution systems, research is being conducted to identify key bacteria in biofilm formation that act as a bridge between early and late colonizers, such as Fusobacterium nucleatum in dental plaque15,22-24). Although studies have been conducted to elucidate the specific formation mechanisms of biofilms in different environments, there is a lack of research on the mechanisms of DUWLs biofilm. The water in DUWLs is discharged from the water tank installed in dentist’s clinic through the pressure generated when the operation button is pressed. The water remains in the waterline and tank except during treatment hours25). Water in DUWLs is known to contain bacteria from the oral cavity and water, providing it a different environment from the oral cavity and typical water distribution systems7,8,26). Therefore, to understand the biofilms that form in unique environments such as DUWLs, it is necessary to identify the formation mechanisms of the bacteria that compose the DUWLs biofilm, which can then be utilized to inhibit and eliminate its formation. Additionally, to understand the mechanism of biofilm formation on DUWLs, coaggregation reactions among the bacteria that comprise DUWLs biofilm should be investigated.
Currently, disinfectants containing a variety of chemicals are used to remove DUWLs biofilm; however, side effects such as blocking waterlines and corrosion of components have been reported, prompting the development of new methods to remove DUWLs biofilms25,27). To develop new methods for removing DUWL biofilms, information about specific agents on the surface of coaggregating bacteria can be utilized. Previous studies have reported that the addition of sugars to protein-saccharide-mediated coaggregation can inhibit coaggregation by blocking lectin protein sites through competitive inhibition15,17,23). Through studies on the inhibition of coaggregation with added sugars, lactose was found to inhibit coaggregation between many oral bacteria but less often between freshwater bacteria28,29). Galactosamine, fucose, and mannose were found to be among the different sugars known to inhibit coaggregation of freshwater bacteria18,30,31). Arginine has also been shown to inhibit the coaggregation of some oral and freshwater bacterial pairs18,32).
However, to date, the mechanisms of biofilm formation and bacterial coaggregation in DUWLs, as well as the inhibition of coaggregation using monosaccharides, have not been studied.
This study investigated the coaggregation of bacterial isolates in the DUWLs and assess the possible mechanisms of coaggregation and inhibition.
A previous study isolated a total of 28 bacterial species (Table 1) from DUWLs, which were stored at –70°C and used in the present study33). Each bacterial species was cultured at 26°C for 7 days in R2A solid medium (Becton, Dickinson and Company, Sparks, MD, USA). A solution of the bacterial culture in R2A liquid medium was used for the experiments.
Bacterial Isolates Used in This Study
Isolate | GenBank | Species match [GenBank accession number] | Homology (%) |
---|---|---|---|
HY1 | MG763899 | Acidovorax delafieldii strain PCWCS4 [GQ284437] | 100 |
HY2 | MW188653 | Acidovorax soli strain DT17-1 [KC920935] | 99 |
HY3 | MW188654 | Afipia broomeae strain 30-RHI-2 [HF558416] | 99 |
HY7 | MW188655 | followed by Beijerinckia derxii subsp. venezuelae strain DSM2329 [AJ563934] | 99 |
HY8 | MW188656 | Bradyrhizobium subterraneum strain 55 1-1 [KP308153] | 100 |
HY10 | MG763900 | Brevundimonas subvibrioides strain ATCC15264 [CP002102] | 99 |
HY11 | MW188657 | Caulobacter segnis strain ATCC21756 [CP002008] | 100 |
HY12 | MG763901 | Cupriavidus pauculus strain KPS201 [AM418462] | 100 |
HY16 | MW188658 | Methylobacterium fujisawaense strain DSM5686 [NR_112232] | 100 |
HY21 | MG763903 | Microbacterium testaceum strain 38A [KC329834] | 100 |
HY23 | MW188661 | Novosphingobium capsulatum strain GIFU11526 [NR_025838] | 100 |
HY31 | MW188662 | Novosphingobium aromaticivorans strain DSM12444 [NR_074261] | 99 |
HY29 | MW188663 | Novosphingobium stygium strain IFO16085 [NR_040826] | 99 |
HY32 | MW188664 | Novosphingobium resinovorum strain NCIMB8767 [EF029110] | 100 |
HY33 | MW188665 | Novosphingobium humi strain R1-4 [NR_157799] | 100 |
HY35 | MW188666 | Novosphingobium nitrogenifigens strain DSM19370 [NR_043857] | 100 |
HY39 | MW188669 | Phenylobacterium muchangponense strain A8 [NR_117783] | 100 |
HY40 | MG763906 | Polaromonas aquatica strain CCUG39797 [AM039831] | 100 |
HY47 | MW188670 | Hydrotalea flava strain CCUG51397 [NR_117026] | 100 |
HY48 | MW188671 | Sphingobium limneticum strain 301 [NR_109484] | 100 |
HY49 | MG763908 | Sphingobium xenophagum strain D5AP82 [JF459960] | 100 |
HY53 | MW188672 | Sphingobium yanoikuyae Q1 [SYU37525] | 100 |
HY54 | MG763909 | Sphingomonas echinoides strain S32312 [AB649019] | 99 |
HY61 | MW188673 | Sphingomonas ginsenosidimutans strain T7AP25-Sg1 [HF930756] | 100 |
HY66 | MW188674 | Sphingomonas oligophenolica strain 0A644 [MH929906] | 100 |
HY67 | MW188675 | Sphingomonas paucimobilis strain NBRC13934 [AB680525] | 100 |
HY69 | MW188676 | Sphingomonas wittichii strain RW1 [NR_074268] | 99 |
HY70 | MG763910 | Sphingopyxis panaciterrae strain Gsoil164 [AB245354] | 100 |
To test the coaggregation of bacterial species, the visual analysis method suggested by Cisar et al.34) was modified and used18). The bacteria were cultured in R2A liquid medium and centrifuged at 3,000×g for 20 minutes. After centrifugation, the bacteria were resuspended in sterile distilled water, measured using a spectrophotometer, and adjusted to 1.5 (optical density [OD]600). A previous study showed that bacterial species isolated from water exhibited reduced coaggregation in buffers with high ionic strength35). Sterile, distilled water was used in this study. A total of 0.2 ml of each concentration-adjusted bacterial solution was put in a glass tube (13 mm) and mixed on a stirring plate for at least 10 seconds. After incubation at room temperature for 1 hour, the aggregation was scored using the following criteria:
0, no coaggregates observed macroscopically; 1, small coaggregates observed macroscopically in turbid suspension; 2, coaggregates easily identified macroscopically; however, the suspension was cloudy; 3, large coaggregates settled immediately, and the supernatant was slightly cloudy; and 4, large coaggregates settled immediately, and the supernatant was clear.
The coaggregation reaction assay was performed in duplicate, and if there was a difference between the two results, it was repeated.
The concentration-adjusted bacterial solution was stained with SYTOⓇ 9 (Invitrogen, Carlsbad, CA, USA) and propidium iodide (Invitrogen)18,36). The stained bacteria were washed twice and resuspended in sterile distilled water. Each bacterial solution was vortexed for 10 seconds and incubated at room temperature for 1 hour to induce coaggregation. Solutions showing coaggregation were transferred onto glass slides, covered with cover glass, and observed under a fluorescence microscope (Leica Microsystems, Heidelberg, Germany).
To assess whether simple sugars and amino acids can inhibit coaggregation, simple sugars and amino acids were added to the coaggregation pairs with scores of three points or higher, and inhibition of coaggregation was observed. The following simple sugars and amino acids were used: galactose (Sigma‐Aldrich Ltd., Gillingham, UK), lactose (Sigma‐Aldrich Ltd.), mannose (Sigma‐Aldrich Ltd.), glucose (Sigma‐Aldrich Ltd.), galactosamine (Sigma‐Aldrich Ltd.), fucose (Sigma‐Aldrich Ltd.), and L-arginine (Sigma-Aldrich Ltd.). A filter-sterilized solution (500 mmol/L) of each simple sugar and amino acid was added to the coaggregated bacterial solution at a final concentration of 50 mmol/L22). This was mixed for 10 seconds after the addition of simple sugars or amino acids and incubated at room temperature for 1 hour for coaggregation. Coaggregation was scored according to the criteria described above.
Coaggregation was observed in 360 out of the 406 coaggregation pairs (Table 2), with coaggregation scores ranging from 1 to 3 points. Five pairs (1.4%) scored 3 points while forty-three pairs (11.9%) scored 2 points. Among the 28 bacterial species, Cupriavidus pauculus HY12 had the highest number of coaggregation pairs, with 10 pairs having scores of 2 or 3 points, followed by Beijerinckia derxii subsp. venezuelae HY7, which had eight coaggregation pairs with scores of 2 or 3 points. Under fluorescence microscopy, additional coaggregation pairs with coaggregation scores of 3 points (Sphingobium limneticum HY48 and C. pauculus HY12) were observed (Fig. 1). In contrast, no coaggregates were observed in pairs that did not show coaggregation responses.
Coaggregation Scores between Bacteria Isolated from Dental Unit Waterlines
Acidovorax delafieldii HY1 | Acidovorax soli HY2 | Afipia broomeae HY3 | Beijerinckia derxii subsp· Venezuelae HY7 | Bradyrhizobium subterraneum HY8 | Brevundimonas subvibrioides HY10 | Caulobacter segnis HY11 | Cupriavidus pauculus HY12 | Methylobacterium fujisawaense HY16 | Microbacterium testaceum HY21 | Novosphingobium capsulatum HY23 | Novosphingobium aromaticivorans HY31 | Novosphingobium stygium HY29 | Novosphingobium resinovorum HY32 | Novosphingobium humi HY33 | Novosphingobium nitrogenifigens HY35 | Phenylobacterium muchangponense HY39 | Polaromonas aquatica HY40 | Hydrotalea flava HY47 | Sphingobium limneticum HY48 | Sphingobium xenophagum HY49 | Sphingobium yanoikuyae HY53 | Sphingomonas echinoides HY54 | Sphingomonas ginsenosidimutans HY61 | Sphingomonas oligophenolica HY66 | Sphingomonas paucimobilis HY67 | Sphingomonas wittichii HY69 | Sphingopyxis panaciterrae HY70 | |
---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
Acidovorax delafieldii HY1 | 1 | 0 | 1 | 0 | 0 | 0 | 0 | 0 | 1 | 0 | 0 | 0 | 1 | 0 | 2 | 1 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | 1 | 0 | 0 | 0 | 1 |
Acidovorax soli HY2 | 0 | 1 | 0 | 0 | 1 | 0 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | 2 | 1 | 1 | 1 | 1 | 1 | 3 | 2 | 1 | 1 | 1 | 0 | 1 | 1 | |
Afipia broomeae HY3 | 1 | 2 | 1 | 1 | 1 | 1 | 1 | 0 | 0 | 1 | 0 | 1 | 1 | 1 | 2 | 0 | 1 | 2 | 1 | 0 | 1 | 1 | 1 | 1 | 1 | 1 | ||
Beijerinckia derxii subsp. Venezuelae HY7 | 1 | 1 | 2 | 1 | 1 | 1 | 2 | 0 | 1 | 2 | 0 | 0 | 1 | 1 | 2 | 1 | 1 | 1 | 2 | 1 | 2 | 1 | 2 | 2 | 1 | |||
Bradyrhizobium subterraneum HY8 | 0 | 1 | 0 | 1 | 1 | 0 | 1 | 1 | 0 | 1 | 0 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | 0 | 1 | 0 | 0 | 1 | 1 | ||||
Brevundimonas subvibrioides HY10 | 1 | 1 | 2 | 2 | 1 | 0 | 1 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | |||||
Caulobacter segnis HY11 | 0 | 1 | 1 | 0 | 1 | 1 | 1 | 2 | 1 | 0 | 1 | 0 | 1 | 1 | 1 | 1 | 0 | 1 | 1 | 0 | 1 | 1 | ||||||
Cupriavidus pauculus HY12 | 2 | 3 | 1 | 0 | 1 | 0 | 1 | 2 | 1 | 1 | 2 | 1 | 3 | 2 | 2 | 1 | 1 | 1 | 2 | 2 | 2 | |||||||
Methylobacterium fujisawaense HY16 | 2 | 1 | 2 | 1 | 0 | 1 | 1 | 2 | 1 | 0 | 1 | 2 | 2 | 0 | 2 | 0 | 0 | 1 | 0 | 1 | ||||||||
Microbacterium testaceum HY21 | 0 | 0 | 0 | 0 | 0 | 1 | 0 | 1 | 0 | 0 | 1 | 0 | 1 | 1 | 1 | 0 | 1 | 0 | 0 | |||||||||
Novosphingobium capsulatum HY23 | 1 | 1 | 0 | 1 | 1 | 0 | 0 | 1 | 0 | 0 | 1 | 1 | 0 | 1 | 0 | 0 | 0 | 1 | ||||||||||
Novosphingobium aromaticivorans HY31 | 1 | 0 | 1 | 2 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | |||||||||||
Novosphingobium stygium HY29 | 0 | 1 | 1 | 1 | 0 | 0 | 0 | 1 | 1 | 1 | 0 | 1 | 0 | 1 | 1 | 1 | ||||||||||||
Novosphingobium resinovorum HY32 | 0 | 0 | 1 | 0 | 2 | 1 | 1 | 1 | 2 | 1 | 1 | 0 | 1 | 2 | 2 | |||||||||||||
Novosphingobium humi HY33 | 2 | 2 | 1 | 1 | 1 | 1 | 3 | 0 | 1 | 1 | 2 | 2 | 3 | 1 | ||||||||||||||
Novosphingobium nitrogenifigens HY35 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | 1 | 1 | 0 | 1 | 1 | 1 | |||||||||||||||
Phenylobacterium muchangponense HY39 | 1 | 0 | 1 | 1 | 1 | 1 | 0 | 1 | 1 | 0 | 0 | 0 | ||||||||||||||||
Polaromonas aquatica HY40 | 0 | 0 | 1 | 1 | 0 | 0 | 1 | 0 | 1 | 1 | 2 | |||||||||||||||||
Hydrotalea flava HY47 | 1 | 2 | 1 | 1 | 1 | 0 | 1 | 1 | 1 | 1 | ||||||||||||||||||
Sphingobium limneticum HY48 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | 1 | |||||||||||||||||||
Sphingobium xenophagum HY49 | 1 | 1 | 1 | 1 | 1 | 1 | 2 | 1 | ||||||||||||||||||||
Sphingobium yanoikuyae HY53 | 1 | 1 | 1 | 0 | 0 | 1 | 1 | |||||||||||||||||||||
Sphingomonas echinoides HY54 | 1 | 1 | 1 | 1 | 1 | 1 | ||||||||||||||||||||||
Sphingomonas ginsenosidimutans HY61 | 1 | 1 | 1 | 1 | 0 | |||||||||||||||||||||||
Sphingomonas oligophenolica HY66 | 1 | 0 | 0 | 0 | ||||||||||||||||||||||||
Sphingomonas paucimobilis HY67 | 1 | 1 | 1 | |||||||||||||||||||||||||
Sphingomonas wittichii HY69 | 0 | 0 | ||||||||||||||||||||||||||
Sphingopyxis panaciterrae HY70 | 0 |
Inhibition of coaggregation was tested in pairs that had coaggregation scores of 3 points or higher. Acidovorax soli HY2 and S. xenophagum HY49 coaggregates were completely inhibited by l-arginine. C. pauculus HY12 and Methylobacterium fujisawaense HY16 coaggregates were completely inhibited by mannose, lactose, and galactose, whereas Novosphingobium humi HY33 and Sphingomonas wittichii HY69 coaggregates were completely inhibited by mannose and lactose (Table 3). Mannose most frequently inhibited coaggregation.
Inhibitory Effect of Sugars or Amino Acids on Coaggregation between Selected Pairs of Isolates
Coaggregation mixture | No. inhibitor | Sugar | Amino acid | ||||||
---|---|---|---|---|---|---|---|---|---|
Galactose | Mannose | Glucose | Lactose | Galactosamine | Fucose | L-arginine | |||
Acidovorax soli HY2+Sphingobium xenophagum HY49 | 3 | 1 | 1 | 1 | 1 | 1 | 1 | 0 | |
Cupriavidus pauculus HY12+Methylobacterium fujisawaense HY16 | 3 | 1 | 0 | 1 | 0 | 0 | 1 | 1 | |
Cupriavidus pauculus HY12+Sphingobium limneticum HY48 | 3 | 1 | 1 | 1 | 2 | 2 | 1 | 2 | |
Novosphingobium humi HY33+Sphingobium xenophagum HY49 | 3 | 3 | 1 | 2 | 1 | 1 | 2 | 1 | |
Novosphingobium humi HY33+Sphingomonas wittichii HY69 | 3 | 1 | 0 | 1 | 0 | 2 | 1 | 1 |
Biofilms mature by adhesion of bacteria to the conditioning film formed on surfaces, coaggregation of early and late adherent bacteria, and binding of extracellular polysaccharides produced by the bacteria37). Bacterial coaggregation is inherently involved in biofilm formation and is affected by the diversity of bacteria in the biofilm17). Therefore, recent studies have focused on investigating the coaggregation responses between isolated bacterial species to understand the mechanisms of biofilm formation in various environments. Examples include studies on the coaggregation of different bacterial species that constitute oral biofilms28,34,38,39). A better understanding of the mechanisms of biofilm formation is essential to develop efficient methods for removing biofilms from DUWLs. However, no studies have specifically investigated the coaggregation of DUWL biofilm-forming bacteria. Therefore, this is the first study to assess the coaggregation between bacterial isolates from DUWL biofilms and investigate the mechanisms of bacterial coaggregation and inhibition.
In our study, a visual assay was used to analyze the coaggregation reaction between the isolates in the DUWLs biofilm. Visual and spectrophotometric assays are the most commonly used methods for coaggregation assays. Buswell et al.22) analyzed the coaggregation response of 19 aquatic biofilm-forming isolates using both visual and spectrophotometric assays. As a result, the sensitivity of the spectrophotometric assay to measure the percentage of coaggregation was high, but owing to the high inter-experiment variability, only a partial result of the spectrophotometric assay was presented22). Based on these results, a visual assay was performed to analyze the coaggregation reaction.
In our study, a microscopic analysis of some coaggregation pairs was performed, in addition to a visual assay. Microscopic analysis was performed to confirm the degree of coaggregation by visual analysis, as visual assays provide high inter-experiment reproducibility but low sensitivity24). In addition, microscopic analyses were performed to determine the morphology of particular interactions and the structure of coaggregates between coaggregation pairs18,36). Coaggregation depends on the relative size and morphology of coaggregating bacteria, and may depend on the density of interacting ligands on the bacterial surface22). The coaggregation that occurs between the oral bacteria F. nucleatum and Streptococcus sanguinis has the structure of ‘corn cobs’40). The formation of these structures is believed to reduce the oxygen tension around F. nucleatum and mask it from other oral species, allowing improved integration within dental plaque18,41,42). In our study, the coaggregation scores revealed by the visual assay and the degree of coaggregation observed by microscopic analysis were similar, and in coaggregates with high coaggregation scores, each bacterium was found to be mixed and integrated with the other. However, not all coaggregation pairs were identified, and the limitations of the maximum magnification did not confirm the detailed structure and density of the ligands. Therefore, it is necessary to observe the coaggregate morphology at high magnification using a microscope, such as an electron microscope24,36).
As a result of the coaggregation analysis of a total of 28 strains by visual analysis, among the isolates tested in this study, B. derxii subsp. venezuelae HY7 and C. pauculus HY12 coaggregated with most strains and formed the highest number of coaggregation pairs with a score of 2 or 3 points. Thus, these isolates may mediate the early and late adhesion of bacteria. According to the results of a previous study on coaggregation reactions between strains isolated from aquatic environments, Micrococcus luteus coaggregated with all 19 isolates tested, except self-aggregation, suggesting the possibility of a bridge role22). Another study suggested Acinetobacter calcoaceticus as a bridge bacterium because it coaggregated with all six isolates tested, including self-aggregation, with a high coaggregation score24). Bridge bacteria are important because they temporarily retain other bacteria on a nascent surface and eventually facilitate colonization for formation of the biofilm23). F. nucleatum has been identified as a major bridge bacterium in dental plaque, and adhesins on F. nucleatum can bind to receptors on the surface of early- and late-colonizing bacteria23). Therefore, inhibiting F. nucleatum adhesion or coaggregation may prevent the accumulation of pathogenic late-colonizing bacteria and this mechanism has been studied43,44). To identify the bridge bacteria among the strains isolated from DUWLs, coaggregation assays with more isolates and further studies to characterize adhesins are needed. Coaggregation between pairs of bacteria is highly specific and is generally mediated by a protein ‘adhesin’ on one cell type and a complementary saccharide ‘receptor’ on the other17). Through this mechanism, the addition of simple sugars can inhibit coaggregation17). In other words, information on the sugars that inhibit coaggregation can be utilized as an anti-adhesion/anti-biofilm method45). Moreover, as anti-adhesion/anti-biofilm methods utilizing sugars do not kill bacteria or inhibit their growth, biofilm formation can be inhibited without the emergence of resistant strains45). Disinfectants containing various chemicals have been used to manage DUWL biofilms, and their disruptive and removal effects have been reported25). However, reports on the adverse effects of disinfectants indicate the need to develop new methods for DUWL biofilm removal. Therefore, the sugars identified in this study that can inhibit coaggregation between strains isolated from DUWLs can be utilized for the development of new DUWLs biofilm removal methods.
Coaggregation was observed between different species of bacteria isolated from water discharged from DUWLs. The bacterial coaggregation score was 1∼3 points. In most cases, coaggregation with scores of 0 or 1 point, indicating weak reactions, was observed. This is consistent with the findings of a previous study in which the level of coaggregation between bacterial species isolated from water was low21,22).
In addition, the inhibition of coaggregation by simple sugars and amino acids was assessed. Among the inhibitors tested, mannose had the strongest inhibitory effect, suggesting that bacterial coaggregation was mediated by carbohydrates. Consistent with our findings, previous studies have shown that the coaggregation of bacterial species isolated from water is inhibited by sugars21,22). Therefore, the coaggregation of these bacterial water isolates, including those isolated from DUWLs, may be mediated by carbohydrates.
In conclusion, this study demonstrated coaggregation between bacterial species isolated from DUWLs and suggested that their coaggregation may be mediated by carbohydrates. This study provides information on the mechanism of DUWL biofilm formation and a basis for the development of more efficient methods for removing DUWL biofilms.
Because oral bacteria belonging to the Streptococcus genus are also found in DUWLs, coaggregation reactions between aquatic and oral bacteria isolated from DUWLs should also be investigated26).
Disinfectant-containing chemicals are commonly used46). Although chemical disinfectants have been reported to successfully inhibit DUWL biofilms, their use is limited by adverse effects such as damage to equipment from chemical reactions, irritation to oral tissues from residues, or reactions with restorations25). Continuous use of chemical disinfectants can lead to the emergence of resistant bacteria. Therefore, the use of coaggregation-inhibiting sugars identified in our study, particularly mannose, as disinfectants, either alone or in combination, may reduce concerns regarding side effects and resistant bacteria45).
The presence or absence of B. derxii subsp. venezuelae HY7 and C. pauculus HY12, which are hypothesized to be the main bacteria playing a bridging role in the formation of DUWL biofilms, can either reduce or promote biofilm formation31). Therefore, the adhesive properties of B. derxii subsp. venezuelae HY7 and C. pauculus HY12 need to be further investigated, and inhibition of their activity and coaggregation may be more effective in inhibiting DUWLs biofilm formation. Based on our results, the utilization of mannose, which showed the greatest inhibitory effect on the coaggregation of C. pauculus HY12, is recommended.
In a previous study that conducted high-throughput DNA sequencing of bacteria in DUWLs, 233 and 394 bacterial genera were identified, respectively. As potentially greater numbers of bacterial species are present in DUWLs than those isolated and tested in this study, future studies must assess coaggregation between more isolates and determine the effects of such coaggregation reactions on the formation of DUWL biofilms7,8). Despite issues with reproducibility, spectrophotometric assays are often used to confirm coaggregation20,22). Because only the degree of coaggregation was visualized in this study, it is necessary to confirm the degree of coaggregation using spectrophotometric assays with higher sensitivity. In addition, considering the duration of DUWL use in dentistry, it is necessary to determine the change in coaggregation over time when incubated for longer than 1 hour. Coaggregation is influenced by the relative size and morphology of the bacteria; however, the relative size and morphology of the DUWLs isolates were not determined in this study22). Future research should address these limitations.
None.
Conflict of interest
No potential conflict of interest relevant to this article was reported.
Ethical approval
This article is not necessary for IRB screening because it is an experimental paper using strains isolated from DUWLs.
Author contributions
Conceptualization: Hye Young Yoon and Si Young Lee. Data acquisition: Hye Young Yoon and Si Young Lee. Formal analysis: Hye Young Yoon and Si Young Lee. Supervision: Hye Young Yoon. Writing-original draft: Si Young Lee. Writing-review & editing: Hye Young Yoon and Si Young Lee.
Funding
None.
Data availability
Please contact the corresponding author for data availability.
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