Antimicrobial activity of a quaternary ammonium methacryloxy silicate-containing acrylic resin: a randomised clinical trial

Si-ying Liu, Lige Tonggu, Li-na Niu, Shi-qiang Gong, Bing Fan, Liguo Wang, Ji-hong Zhao, Cui Huang, David H Pashley, Franklin R Tay, Si-ying Liu, Lige Tonggu, Li-na Niu, Shi-qiang Gong, Bing Fan, Liguo Wang, Ji-hong Zhao, Cui Huang, David H Pashley, Franklin R Tay

Abstract

Quaternary ammonium methacryloxy silicate (QAMS)-containing acrylic resin demonstrated contact-killing antimicrobial ability in vitro after three months of water storage. The objective of the present double-blind randomised clinical trial was to determine the in vivo antimicrobial efficacy of QAMS-containing orthodontic acrylic by using custom-made removable retainers that were worn intraorally by 32 human subjects to create 48-hour multi-species plaque biofilms, using a split-mouth study design. Two control QAMS-free acrylic disks were inserted into the wells on one side of an orthodontic retainer, and two experimental QAMS-containing acrylic disks were inserted into the wells on the other side of the same retainer. After 48 hours, the disks were retrieved and examined for microbial vitality using confocal laser scanning microscopy. No harm to the oral mucosa or systemic health occurred. In the absence of carry-across effect and allocation bias (disks inserted in the left or right side of retainer), significant difference was identified between the percentage kill in the biovolume of QAMS-free control disks (3.73 ± 2.11%) and QAMS-containing experimental disks (33.94 ± 23.88%) retrieved from the subjects (P ≤ 0.001). The results validated that the QAMS-containing acrylic exhibits favourable antimicrobial activity against plaque biofilms in vivo. The QAMS-containing acrylic may also be used for fabricating removable acrylic dentures.

Figures

Figure 1. CONSORT flow diagram of subject…
Figure 1. CONSORT flow diagram of subject randomisation and selection criteria.
Figure 2. Representative BacLight-stained 48-hour biofilm grown…
Figure 2. Representative BacLight-stained 48-hour biofilm grown on the surface of QAMS-containing acrylic resin (experimental disk) with >25% kill within the biovolume (Subject-3).
(a) 3-D projection for the green, red and merged channels of the biofilm (the depth axis (Z-axis) was scaled-in to enhance presentation of the thickness of the biofilm). Green channel: live microorganisms; red channel: dead microorganisms. (b) Live/dead microorganism distribution from the bottom to the top of the biofilm, indicating the percentage of kill in different layers of the biofilms. (c) Perspectives of the biofilm from the X-Z plane of the 3-D projection (rectangular blocks beneath the projection) taken at one-quarter (0.25), one-half (0.50) and three-quarter (0.75) width of the Y-axis of the biofilm. (d) Perspectives of the biofilm from the Y-Z plane of the 3-D projection taken at 0.25, 0.50 and 0.75 width of the X-axis of the biofilm.
Figure 3. The corresponding BacLight-stained 48-hour biofilm…
Figure 3. The corresponding BacLight-stained 48-hour biofilm grown on the surface of QAMS-free acrylic resin (control disk) of the subject presented in Figure 3 (Subject-3).
(a) 3-D projection for the green, red and merged channels of the biofilm (Z-axis scaled-in to the same extent as Fig. 3). Green channel: live microorganisms; red channel: dead microorganisms. (b) Live/dead microorganism distribution from the bottom to the top of the biofilm, indicating the percentage of kill in different layers of the biofilms. (c) Perspectives of the biofilm viewed from the X-Z plane at 0.25, 0.50 and 0.75 width of the Y-axis. (d) Perspectives of the biofilm viewed from the Y-Z plane at 0.25, 0.50 and 0.75 width of the X-axis.
Figure 4. Representative BacLight-stained 48-hour biofilm grown…
Figure 4. Representative BacLight-stained 48-hour biofilm grown on the surface of QAMS-containing acrylic resin (experimental disk) with
(a) 3-D projection for the green, red and merged channels of the biofilm (Z-axis scaled-in to the same extent as Fig. 3). Green channel: live microbes; red channel: dead microbes. (b) Live/dead microbe distribution from the bottom to the top of the biofilm. (c) Perspectives of the biofilm viewed from the X-Z plane at 0.25, 0.50 and 0.75 width of the Y-axis. (d) Perspectives of the biofilm viewed from the Y-Z plane at 0.25, 0.50 and 0.75 width of the X-axis.
Figure 5. Example of plaque biofilms containing…
Figure 5. Example of plaque biofilms containing mixed bacterial-fungal components.
(a) BacLight-stained CLSM merged image showing the presence of live (green) fungal hyphae (open arrowhead) within the biovolume of a biofilm taken from a control disk of one subject (subject 21). (b) The corresponding BacLight-stained CLSM merged image showing the presence of dead (red) fungal hyphae killed by the QAMS-containing acrylic (open arrowhead) from an experimental disk of the same subject (subject 21).
Figure 6. Appliance design.
Figure 6. Appliance design.
(a) Schematic illustrating the relationship between a test disk and its fitting well within the retainer. The disk surface to be examined was turned toward the palate but not in direct contact with the palate. Each well was protected from the oral cavity by orthodontic acrylic to prevent disturbance of the biofilm by the tongue. (b) Each appliance contained 4 wells to house two 6-mm diameter retrievable control disks and 2 similar diameter retrievable experimental PMMA disks. Allocation of the control disks to the left or right side of the retainer was decided by a randomised number sheet; the experimental disks were inserted subsequently into the wells on the contralateral side.

References

    1. Okunseri C., Pajewski N. M., McGinley E. L. & Hoffmann R. G. Racial/ethnic disparities in self-reported pediatric visits in the United States. J. Public Health Dent. 67, 217–223 (2007).
    1. Whitesides J., Pajewski N. M., Bradley T. G., Iacopino A. M. & Okunseri C. Socio-demographics of adult orthodontic visits in the United States. Am. J. Orthod. Dentofacial Orthop. 133, 489. e9–14 (2008).
    1. Manski R. J. & Brown E. Dental Procedures, United States, 1999 and 2009. Statistical Brief #368. Agency for Healthcare Research and Quality, Rockville, MD. (2012) Available at: . (Accessed: 26th January, 2016).
    1. Littlewood S. J., Millett D. T., Doubleday B., Bearn D. R. & Worthington H. V. Orthodontic retention: a systematic review. J. Orthod. 33, 205–212 (2006).
    1. Westerlund A. et al. Stability and side effects of orthodontic retainers – a systematic review. Dentistry 4, 258 (2014).
    1. Pathak A. K. & Sharma D. S. Biofilm associated microorganisms on removable oral orthodontic appliances in children in the mixed dentition. J. Clin. Pediatr. Dent. 37, 335–339 (2013).
    1. Batoni G. et al. Effect of removable orthodontic appliances on oral colonisation by mutans streptococci in children. Eur. J. Oral Sci. 109, 388–392 (2001).
    1. Addy M., Shaw W. C., Hansford P. & Hopkins M. The effect of orthodontic appliances on the distribution of Candida and plaque in adolescents. Br. J. Orthod. 9, 158–163 (1982).
    1. Bjerklin K., Gärskog B. & Rönnerman A. Proximal caries increment in connection with orthodontic treatment with removable appliances. Br. J. Orthod. 10, 21–24 (1983).
    1. Hibino K., Wong R. W., Hägg U. & Samaranayake L. P. The effects of orthodontic appliances on Candida in the human mouth. Int. J. Paediatr. Dent. 19, 301–308 (2009).
    1. Chang C. S., Al-Awadi S., Ready D. & Noar J. An assessment of the effectiveness of mechanical and chemical cleaning of Essix orthodontic retainer. J. Orthod. 41, 110–117 (2014).
    1. Patel A., Burden D. J. & Sandler J. Medical disorders and orthodontics. J. Orthod. 36, 1–21 (2009).
    1. Coulthwaite L. & Verran J. Potential pathogenic aspects of denture plaque. Br. J. Biomed. Sci. 64, 180–189 (2007).
    1. Verran J. Malodour in denture wearers: an ill-defined problem. Oral Dis. 11 (Suppl 1), 24–28 (2005).
    1. Vento-Zahra E., De Wever B., Decelis S., Mallia K. & Camilleri S. Randomized, double-blind, placebo-controlled trial to test the efficacy of nitradine tablets in maxillary removable orthodontic appliance patients. Quintessence Int. 42, 37–43 (2011).
    1. Fathi H., Martiny H. & Jost-Brinkmann P. G. Efficacy of cleaning tablets for removable orthodontic appliances: an in vivo pilot study. J. Orofac. Orthop. 76, 143–151 (2015).
    1. Xu X. M. & Costin S. Antimicrobial polymeric dental material. In Cerrada M. (ed). Polymeric materials with antimicrobial activity: from synthesis to applications, pp. 279–304 (The Royal Society of Chemistry. 2014).
    1. Kenawy el-R., Worley S. D. & Broughton R. The chemistry and applications of antimicrobial polymers: a state-of-the-art review. Biomacromolecules 8, 1359–1384 (2007).
    1. Siedenbiedel F. & Tiller J. C. Antimicrobial polymers in solution and on surfaces: Overview and functional principles. Polymers 4, 46–71 (2012).
    1. Raj P. A. & Dentino A. R. Denture polymers with antimicrobial properties: a review of the development and current status of anionic poly(methyl methacrylate) polymers. Future. Med. Chem. 5, 1635–1645 (2013).
    1. Carmona-Ribeiro A. M. & de Melo Carrasco L. D. Cationic antimicrobial polymers and their assemblies. Int. J. Mol. Sci. 14, 9906–9946 (2013).
    1. Gong S. Q. et al. Quaternary ammonium silane-functionalized, methacrylate resin composition with antimicrobial activities and self-repair potential. Acta Biomater. 8, 3270–3282 (2012).
    1. Gong S. Q. et al. An ORMOSIL-containing orthodontic acrylic resin with concomitant improvements in antimicrobial and fracture toughness properties. PLoS One 7, e42355 (2012).
    1. Gong S. Q. et al. Effect of water-aging on the antimicrobial activities of an ORMOSIL-containing orthodontic acrylic resin. Acta Biomater. 9, 6964–6973 (2013).
    1. U. S. Food and Drug Administration. 510(k) premarket notification K141439. (2014) Available at: (Accessed: 21th August 2015).
    1. Shu M., Wong L., Miller J. H. & Sissons C. H. Development of multi-species consortia biofilms of oral bacteria as an enamel and root caries model system. Arch. Oral Biol. 45, 27–40 (2000).
    1. Guggenheim B., Guggenheim M., Gmür R., Giertsen E. & Thurnheer T. Application of the Zürich biofilm model to problems of cariology. Caries Res. 38, 212–222 (2004).
    1. Chávez de Paz L. E. Development of a multispecies biofilm community by four root canal bacteria. J. Endod. 38, 318–323 (2012).
    1. Al-Ahmad A. et al. The in vivo dynamics of Streptococcus spp., Actinomyces naeslundii, Fusobacterium nucleatum and Veillonella spp. in dental plaque biofilm as analysed by five-colour multiplex fluorescence in situ hybridization. J. Med. Microbiol. 56 (Pt 5), 681–687 (2007).
    1. Zijnge V. et al. Oral biofilm architecture on natural teeth. PLoS One 5, e9321 (2010).
    1. van’t Hof W., Veerman E. C., Nieuw Amerongen A. V. & Ligtenberg A. J. Antimicrobial defense systems in saliva. Monogr. Oral Sci. 24, 40–51 (2014).
    1. Bjarnsholt T. et al. The in vivo biofilm. Trends Microbiol. 21, 466–474 (2013).
    1. Dewhirst F. E. et al. The human oral microbiome. J. Bacteriol. 192, 5002–5017 (2010).
    1. Toyofuku M. et al. Environmental factors that shape biofilm formation. Biosci. Biotechnol. Biochem. Jun 23:1-6 [Epub ahead of print] (2015).
    1. Goo E., An J. H., Kang Y. & Hwang I. Control of bacterial metabolism by quorum sensing. Trends Microbiol. 23, 567–576. (2015).
    1. Dixon E. F. & Hall R. A. Noisy neighbourhoods: quorum sensing in fungal polymicrobial infections. Cell Microbiol. 10.1111/cmi.12490.[Epubahead of print] (2015).
    1. Lesaffre E., Philstrom B., Needleman I. & Worthington H. The design and analysis of split-mouth studies: what statisticians and clinicians should know. Stat. Med. 28, 3470–3482 (2009).
    1. Pandis N., Walsh T., Polychronopoulou A., Katsaros C. & Eliades T. Split-mouth designs in orthodontics: an overview with applications to orthodontic clinical trials. Euro. J. Ortho. 35, 783–789 (2013).
    1. Suresh K. P. An overview of randomization techniques: An unbiased assessment of outcome in clinical research. J. Hum. Reprod. Sci. 4, 8–11 (2011).
    1. Jüni P., Altman D. G. & Egger M. Systematic reviews in health care: Assessing the quality of controlled clinical trials. Br. Med. J. 323, 42–46 (2001).
    1. Boulos L., Prévost M., Barbeau B., Coallier J. & Desjardins R. LIVE/DEAD BacLight: application of a new rapid staining method for direct enumeration of viable and total bacteria in drinking water. J. Microbiol. Methods. 37, 77–86 (1999).
    1. Berney M., Hammes F., Bosshard F., Weilenmann H. U. & Egli T. Assessment and interpretation of bacterial viability by using the LIVE/DEAD BacLight Kit in combination with flow cytometry. Appl. Environ. Microbiol. 73, 3283–3290 (2007).
    1. Stiefel P., Schmidt-Emrich S., Maniura-Weber K. & Ren Q. Critical aspects of using bacterial cell viability assays with the fluorophores SYTO9 and propidium iodide. BMC Microbiol. 15, 36 (2015).
    1. Schwank S., Rajacic Z., Zimmerli W. & Blaser J. Impact of bacterial biofilm formation on in vitro and in vivo activities of antibiotics. Antimicrob. Agents Chemother. 42, 895–898 (1998).
    1. Jentsch H. F., Eckert F. R., Eschrich K., Stratul S. I. & Kneist S. Antibacterial action of Chlorhexidine/thymol containing varnishes in vitro and in vivo. Int. J. Dent. Hyg. 12, 168–173 (2014).
    1. Gottenbos B., Grijpma D. W., van der Mai H. C., Feijen J. & Busscher H. J. Antimicrobial effects of positively charged surfaces on Gram-positive and Gram-negative bacteria. J. Antimicro. Chemo. 48, 7–13 (2001).
    1. von Canstein H., Kelly S., Li Y. & Wagner-Dobler I. Species diversity improves the efficiency of mercury-reducing biofilms under changing environmental conditions. Appl. Environ. Microbiol. 68, 2829–2837 (2002).
    1. Leriche V., Briandet R. & Carpentier B. Ecology of mixed biofilms subjected daily to a chlorinated alkaline solution: spatial distribution of bacterial species suggests a protective effect of one species to another. Environ. Microbiol. 5, 64–71 (2003).
    1. Shirtliff M. E., Peters B. M. & Jabra-Rizk M. A. Cross-kingdom interactions: Candida albicans and bacteria. FEMS Microbiol. Lett. 299, 1–8 (2009).
    1. Bayles K. W. Bacterial programmed cell death: making sense of a paradox. Nat. Rev. Microbiol. 12, 63–69 (2014).
    1. Allocati N., Masulli M., Di llio C. & De Laurenzi V. Die for the community: an overview of programmed cell death in bacteria. Cell Death Dis. 6, e1609 (2015).
    1. Bamford C. V. et al. Streptococcus gordonii modulates Candida albicans biofilm formation through intergeneric communication. Infect. Immun. 77, 3696–3704 (2009).
    1. Schulz K. F., Altman D. G., Moher D. & CONSORT Group. CONSORT 2010 statement: updated guidelines for reporting parallel group randomized trials. Ann. Intern. Med. 152, 726–732 (2010).
    1. Haahr M. T. & Hróbjartsson A. Who is blinded in randomized clinical trials? A study of 200 trials and a survey of authors. Clin. Trials 3, 360–365 (2006).
    1. Ahlström B., Thompson R. A. & Edebo L. The effect of hydrocarbon chain length, pH, and temperature on the binding and bactericidal effect of amphiphilic betaine esters on Salmonella typhimurium. APMIS 107, 318–324 (1999).
    1. Oosterhof J. J., Buijssen K. J., Busscher H. J., van der Laan B. F. & van der Mei H. C. Effects of quaternary ammonium silane coatings on mixed fungal and bacterial biofilms on tracheoesophageal shunt prostheses. Appl. Environ. Microbiol. 72, 3673–3677 (2006).
    1. Chávez de Paz L. E. Image analysis software based on color segmentation for characterization of viability and physiological activity of biofilms. Appl. Environ. Microbiol. 75, 1734–1739 (2009).

Source: PubMed

Sponsors et collaborateurs

Les conditions médicales

Interventions en matière de drogue

3
S'abonner