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Biofilm on orthodontic retention wiresJongsma, Marije

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Biofilm on orthodontic retention wires- an in vitro and in vivo study -

Marije A. Jongsma

Publication of this thesis was sponsored by:- Prof. K.G. Bijlstrastichting- Nederlandse Vereniging van Orthodontisten (NVvO)- Ortholab B.V.- Dentsply Lomberg B.V.- Orthodontisch Laboratorium Friesland B.V. - Noord Negentig accountants en belastingadviseurs

Biofilm on orthodontic retention wires - an in vitro and in vivo study -

Door Marije Albertine JongsmaUniversitair Medisch Centrum Groningen, Rijksuniversiteit GroningenGroningen, NederlandCover and layout: MidasMentink.nlCopyright © 2015 by Marije A. JongsmaPrinted by: GildeprintISBN (printed version) 978-94-6108-923-6ISBN (electronic version) 978-94-6108-924-3

Biofilm on orthodontic retention wires

- an in vitro and in vivo study -

Proefschriftter verkrijging van de graad van doctor aan de

Rijksuniversiteit Groningen

op gezag van de

Rector Magnificus prof. dr. E. Sterken

en volgens besluit van het College van Promoties.

De openbare verdediging zal plaatsvinden op

woensdag 1 april 2015 om 14.30 uur

door

Marije Albertine Jongsmageboren op 3 februari 1986

te Den Helder

Promotores

Prof. dr. Y. Ren

Prof. dr. ir. H.J. Busscher

Prof. dr. H.C. van der Mei

Beoordelingscommissie

Prof. dr. S.K. Bulstra

Prof. dr. J.M. ten Cate

Prof. dr. H. He

Aan mijn ouders

Paranimfen

Leontine A. Jongsma

Monique J.M. Vink-Vos

CONTENTS

Chapter 1 General introduction and aim of the thesis 9

Chapter 2 Orthodontic treatment with fixed appliances and 15

biofilm formation - a potential public health threat?

Clinical Oral Investigations (2013) 17:1209-1218

Chapter 3 Biofilm formation on stainless steel and gold wires for 33

bonded retainers in vitro and in vivo and their susceptibility

to oral antimicrobials.


Chapter 4 In vivo biofilm formation on stainless steel bonded-retainers 53

during different regimens of oral health care.

International Journal of Oral Science (2015)

doi:10.1038/ijos.2014.69

Chapter 5 Stress relaxation analysis facilitates a quantitative approach 73

towards antimicrobial penetration into biofilms.

PLoS One (2013) 8:e63750

Chapter 6 Synergy of brushing mode and antibacterial use on in 95

vivo biofilm formation

Submitted to: Journal of Dentistry

Chapter 7 General discussion 109

Summary 115

Nederlandse samenvatting 121

Dankwoord (acknowledgements) 129

Curriculum Vitae 135

Chapter 1General introduction and aim of the thesis

10

General introductionChapter 1

Orthodontic treatment is very common amongst both juveniles and adults and the number of orthodontic patients is still increasing every year.1 During orthodontic treatment one of the greatest challenges is to prevent biofilm related complications such as gingivitis, gingival hyperplasia and white spot lesions.2-4 Orthodontic appliances provide extra retention sites for biofilm formation and make removal of the biofilm through natural cleansing and tooth brushing more difficult.5 Despite all efforts to prevent these biofilm related complications, they are still quite common: gingivitis occurs in almost all orthodontic patients2 and white spot lesions occur in about 60% of orthodontic patients.3,6

After active orthodontic treatment, some form of retention of the dentition is required to maintain the treatment result, since long-term stability cannot be guaranteed.7 Different types of retention methods can be applied, such as the use of removable acrylic plates, vacuum formed retainers or bonded retention wires. It is increasingly common to place permanent bonded retention wires behind the anterior teeth.8 This means that after a lengthy orthodontic treatment, a much longer phase of retention treatment follows. Bonded retention wires are generally very effective in preventing the teeth from relapsing to their pre-treatment position,9,10 but the drawback of these retainers is that biofilm and calculus accumulate along the wires,11 leading to a greater incidence of gingival recession, increased pocket depth and bleeding on probing.12,13 With a growing number of orthodontic patients, prevention of biofilm related complications becomes more and more important in patients both under active treatment as well as when in the retention phase of treatment.

Mechanical removal of the biofilm remains the most important way to establish oral hygiene. However, orthodontic appliances and retention wires provide many crevices and niches in which biofilm can grow out of reach for mechanical removal. In general, powered toothbrushes provide better biofilm removal than manual toothbrushes14 and they can mechanically disrupt a biofilm from a distance due to strong fluid flows,15 air bubble inclusion16 and acoustic energy transfer. Nevertheless in orthodontic patients the beneficial effect of powered brushing is much smaller, if even present.17 In both orthodontic as well as in non-orthodontic patients, 100% biofilm removal can never be achieved18 and a part of the biofilm will always be left behind at locations out of reach for mechanical removal.

Chemical control of oral biofilms is an approach, additional to mechanical biofilm control, in preventing biofilm related complications. Various oral antimicrobials are available in the form of toothpastes, gels and mouthrinses, such as chlorhexidine, cetylpiridium chloride, stannous fluoride, triclosan and essential oils.19,20 Planktonic bacteria are much more susceptible to antimicrobials than bacteria growing in a biofilm.21 In the oral cavity bacteria are mainly present in a biofilm mode of growth. Oral biofilms are diverse communities of microorganisms, embedded in a self-produced matrix of extracellular-polymeric-substances.22 The

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extracellular matrix acts not only as a glue for the biofilm, ensuring adhesion to a substratum and integrity of the biofilm itself,23 but also hampers penetration of antimicrobials into the biofilm to offer protection to organisms in a biofilm mode of growth.

Previous studies have shown that after a single self performed brushing, about 40% - 50% of the biofilm is left behind.24,25 This biofilm is potentially harmful, but once antimicrobials have penetrated the biofilm, it can also act beneficially as a reservoir for oral antimicrobials,26 ensuring their prolonged action. The antimicrobials absorbed in biofilm left behind can be released over time in effective amounts, preventing new biofilm formation.27 This demonstrates that penetration of antimicrobials into oral biofilm is very important for both direct and prolonged action in controlling the biofilm. By mechanically disrupting the biofilm and therewith simultaneously altering its structure and viscoelastic properties,18 absorption of antimicrobials will be enhanced. Due to the to the crevices and niches in orthodontic appliances and retention wires, mechanical disruption of the biofilm is difficult by manual brushing, but is likely to occur through non-contact brushing with a powered toothbrush.18

In this thesis we focus only on biofilms formed on orthodontic retention wires. Many different types of retention wires are available, as can be divided in two groups: single-strand wires and multi-strand wires. Multi-strand wires provide additional flexibility compared to single-strand wires, which allows physiologic movement of the bonded teeth instead of fixing them all as one unit. Therefore multi-strand wires are bonded to all front teeth, whereas single-strand wires are generally only bonded to the canines.28-30 From a clinical point of view, multi-strand wires are preferred, since their long-term effectiveness in preventing incisor irregularity is higher than that of single-strand wires.9,10

We hypothesise that the amount of biofilm formation is dependent on the wire type, since the crevices and niches in the multi-strand wires provide a protected environment for biofilm growth.31 For this same reason, we hypothesise that the effect of manual removal of the biofilm and chemical control through oral antimicrobials is reduced for multi-strand wires compared to single-strand wires. Furthermore we hypothesise that to improve antimicrobial penetration into the biofilm of the multi-strand wires, it is beneficial to mechanically disrupt the biofilm by powered toothbrushing that has been proved to provide the energy necessary for disrupting the structure of the biofilm.

The general aim of this thesis is to verify the above hypotheses through evaluating the factors that play a role on biofilm formation on orthodontic retention wires and to determine how biofilm formation and antimicrobial penetration into the biofilm can be influenced.

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REFERENCES

1. American Association of Orthodontists (2012) AAO Patient Census Surveys 1989-2010. Bull Am Assoc Orthod 2. Renkema AA, Dusseldorp JK, Middel B, Ren Y (2010) Enlargement of the gingiva during treatment with fixed orthodontic appliances. Ned Tijdschr Tandheelkd 117:507-512 3. Enaia M, Bock N, Ruf S (2011) White-spot lesions during multibracket appliance treatment: A challenge for clinical excellence. Am J Orthod Dentofacial Orthop 140:e17-e24 4. Hadler-Olsen S, Sandvik K, El-Agroudi MA, Øgaard B (2012) The incidence of caries and white spot lesions in orthodontically treated adolescents with a comprehensive caries prophylactic regimen—a prospective study. Eur J Orthod 34:633-639 5. Øgaard B (2008) White Spot Lesions During Orthodontic Treatment: Mechanisms and Fluoride Preventive Aspects. Semin Orthod 14:183-193 6. Hadler-Olsen S, Sandvik K, El-Agroudi MA, Ogaard B (2012) The incidence of caries and white spot lesions in orthodontically treated adolescents with a comprehensive caries prophylactic regimen--a prospective study. Eur J Orthod 34:633-639 7. Little RM (1999) Stability and relapse of mandibular anterior alignment: University of Washington Studies. Semin Orthod 5:191-204 8. Renkema AM, Hélène Sips ET, Bronkhorst E, Kuijpers-Jagtman AM (2009) A survey on orthodontic retention procedures in the Netherlands. Eur J Orthod 31:432-437 9. Renkema A, Al-Assad S, Bronkhorst E, Weindel S, Katsaros C, Lisson JA (2008) Effectiveness of lingual retainers bonded to the canines in preventing mandibular incisor relapse. Am J Orthod Dentofacial Orthop 134:179.e1-179.e8 10. Renkema A, Renkema A, Bronkhorst E, Katsaros C (2011) Long-term effectiveness of canine-to-canine bonded flexible spiral wire lingual retainers. Am J Orthod Dentofacial Orthop 139:614-621 11. Artun J (1984) Caries and periodontal reactions associated with long-term use of different types of bonded lingual retainers. Am J Orthod 86:112-118 12. Pandis N, Vlahopoulos K, Madianos

P, Eliades T (2007) Long-term periodontal status of patients with mandibular lingual fixed retention. Eur J Orthod 29:471-476 13. Levin L, Samorodnitzky-Naveh GR, Machtei EE (2008) The association of orthodontic treatment and fixed retainers with gingival health. J Periodontol 79:2087-2092 14. Yaacob M, Worthington HV, Deacon SA, Deery C, Walmsley AD, Robinson PG, Glenny AM (2014) Powered versus manual toothbrushing for oral health. Cochrane Database Syst Rev 6:CD002281 15. Van der Mei HC, Rustema-Abbing M, Bruinsma GM, Gottenbos B, Busscher HJ (2007) Sequence of oral bacterial co-adhesion and non-contact brushing. J Dent Res 86:421-425 16. Parini MR, Pitt WG (2006) Dynamic removal of oral biofilms by bubbles. Colloids Surf B Biointerfaces 52:39-46 17. Kaklamanos EG, Kalfas S (2008) Meta-analysis on the effectiveness of powered toothbrushes for orthodontic patients. Am J Orthod Dentofacial Orthop 133:187.e1-187.e14 18. Busscher HJ, Jager D, Finger G, Schaefer N, Van Der Mei HC (2010) Energy transfer, volumetric expansion, and removal of oral biofilms by non-contact brushing. Eur J Oral Sci 118:177-182 19. Stoeken JE, Paraskevas S, Van der Weijden GA (2007) The long-term effect of a mouthrinse containing essential oils on dental plaque and gingivitis: a systematic review. J Periodontol 78:1218-1228 20. Addy M, Moran J (2008) Chemical Supragingival Plaque Control. In: Lang NP, Lindhe J (eds) Clinical periodontology and implant dentistry, Vol 2 edn. Blackwell Munksgaard, pp 734-765 21. Van der Mei HC, White DJ, Atema-Smit J, Van de Belt-Gritter E, Busscher HJ (2006) A method to study sustained antimicrobial activity of rinse and dentifrice components on biofilm viability in vivo. J Clin Periodontol 33:14-20 22. Marsh PD (2010) Microbiology of Dental Plaque Biofilms and Their Role in Oral Health and Caries. Dent Clin North Am 54:441-454 23. Flemming HC, Wingender J (2010) The biofilm matrix. Nat Rev Microbiol 8:623-633 24. Van der Weijden GA, Echeverria JJ, Sanz

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M, Lindhe J (2008) Mechanical Supragingival Plaque Control. In: J. Lindhe, N. P. Lang, and T. Karring (ed) Clinical Periodontology and Implant Dentistry, 5th edn. Blackwell Munskgaard, Copenhagen, pp 705-733 25. Paraskevas S, Timmerman MF, Van der Velden U, Van der Weijden GA (2006) Additional effect of dentifrices on the instant efficacy of toothbrushing. J Periodontol 77:1522-1527 26. Otten MP, Busscher HJ, Abbas F, Van der Mei HC, Van Hoogmoed CG (2012) Plaque-left-behind after brushing: intra-oral reservoir for antibacterial toothpaste ingredients. Clin Oral Investig 16:1435-1442 27. Otten MP, Busscher HJ, Van der Mei HC, Abbas F, Van Hoogmoed CG (2010) Retention of antimicrobial activity in plaque and saliva following mouthrinse use in vivo. Caries Res 44:459-464

28. Artun J, Zachrisson B (1982) Improving the handling properties of a composite resin for direct bonding. Am J Orthod 81:269-276 29. Zachrisson BU (1982) The bonded lingual retainer and multiple spacing of anterior teeth. Swed Dent J Suppl 15:247-255 30. Bearn DR (1995) Bonded orthodontic retainers: a review. Am J Orthod Dentofacial Orthop 108:207-213 31. Al-Nimri K, Al Habashneh R, Obeidat M (2009) Gingival health and relapse tendency: a prospective study of two types of lower fixed retainers. Aust Orthod J 25:142-146

Chapter 2 Orthodontic treatment with fixed appliances and biofilm

formation – a potential public health threat?

Yijin Ren, Marije A. Jongsma, Li Mei, Henny C. van der Mei, Henk J. Busscher

Clinical Oral Investigations (2014) 18: 1711-1718

16

Challenges in orthodontic treatmentChapter 2

ABSTRACT

Objectives Orthodontic treatment is highly popular for restoring function and facial esthetics in juveniles and adults. As a downside, prevalence of biofilm-related complications is high. Objectives of this review are to (1)-identify special features of biofilm formation in orthodontic-patients and (2)-emphasize the need for strong concerted action to prevent biofilm-related complications during orthodontic treatment.

Materials and methods Literature on biofilm formation in the oral cavity is reviewed to identify special features of biofilm formation in orthodontic patients. Estimates are made of juvenile and adult orthodontic-patient-population sizes and biofilm-related complication rates are used to indicate the costs and clinical workload resulting from biofilm-related complications.

Results Biofilm formation in orthodontic patients is governed by similar mechanisms as common in the oral cavity. However, orthodontic-appliances hamper maintenance of oral hygiene and provide numerous additional surfaces, with properties alien to the oral cavity, to which bacteria can adhere and form a biofilm. Biofilm formation may lead to gingivitis and white spot lesions, compromising facial esthetics. Whereas gingivitis after orthodontic treatment is often transient, white spot lesions may turn into cavities requiring professional restoration. Complications requiring professional care develop in 15% of all orthodontic patients, implying an annual cost of over US$ 500,000,000 and a workload of 1000 fulltime dentists in the USA alone.

Conclusions Improved preventive measures and antimicrobial materials are urgently required to prevent biofilm-related complications of orthodontic treatment from overshadowing its functional and esthetic advantages.

Clinical relevance High treatment demand and occurrence of biofilm-related complications requiring professional care make orthodontic treatment a potential public health threat.

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INTRODUCTION

Orthodontic treatment for restoring function and facial esthetics is highly popular. Between 1982 and 2010 the number of orthodontic patients in North America has increased by 100% (Fig. 1). Together there are nearly four million juvenile, aged between 6-18 years, and more than one million adult patients in North America alone reported by the American Association of Orthodontists.1 The juvenile patients constitute about 7% of the total population,2 which is much lower than the number of juvenile patients with an objective orthodontic treatment need, estimated to be between 17-43%.3 When subjective treatment need is taken into consideration, 50-75% of the Western population could benefit from orthodontic treatment.1 Therefore, the number of potential orthodontic patients is much larger than currently treated and further increase in the number of orthodontic patients over the coming years can be expected with increasing self-awareness of dental esthetics, oral health related quality of life and affordability of orthodontic treatment.

However, the downside of orthodontic treatment has not been much addressed. The region of the tooth surface around brackets is prone to adhesion of oral bacteria and subsequent biofilm formation or “dental plaque”. Oral biofilms on dental hard and soft tissues are the main cause of dental diseases, including caries and periodontal disease and are difficult to remove. A single-time, self-performed manual brushing4 is often insufficient and known to leave biofilm behind in retention sites, such as fissures, interproximal spaces and gingival

Figure 1. Number of orthodontic patients in North America over the past three decades. For 1982-1986, no data are available about the percentage of adult patients. (source: American Association of Orthodontists: AAO Patient Census Surveys 1989-2010. Bull Am Assoc Orthod 2012)

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margins. Orthodontic appliances make effective biofilm removal even more difficult and brushing nearly always leaves biofilm behind at the vulnerable bracket-adhesive-enamel junction and the sensitive region between brackets and gingival margin (Fig. 2), therewith contributing to the occurrence of dental diseases.

In the current review, we identify the special features of biofilm formation in orthodontic patients, without aiming to fully describe mechanisms of oral biofilm formation in general, and provide an estimate of the occurrence of biofilm-related complications during orthodontic treatment, including consequences for dental health care in general.

Oral biofilm formationWhereas it is beyond the scope of this review, to fully describe mechanisms of oral biofilm formation in general, we will briefly outline some important features. Oral biofilms form on all surfaces exposed to the human oral cavity, most notably on all oral hard and soft surfaces. Oral biofilms formed on tooth surfaces cause demineralization of enamel, which in its mildest form yields white spot lesions, indicative of sub-surface decalcification. Biofilm formed below the gingival margin leads to inflammation of the gums, which in an extreme case can lead to periodontitis and tooth loss.

Oral biofilms are diverse communities of adhering microorganisms, embedded in a self-produced matrix of extracellular-polymeric-substances and possessing a complex, spatially heterogeneous and dynamic structure.5 The extracellular matrix acts not only as a glue for the biofilm, ensuring adhesion to a substratum and integrity of the biofilm itself,6 but also hampers

Figure 2. Orthodontic biofilm, visualized by staining with GUM red-cote, before (lower dentition) and after (upper dentition) removal of brackets. Stained areas, representing oral biofilm, can be clearly seen on the tooth surfaces around the area where the brackets have been bonded, around the brackets still present and along the gingival margins.

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penetration of antimicrobials into the biofilm to offer protection to organisms in a biofilm mode of growth. Although the bacterial diversity in the oral cavity is estimated to include at least 800 different species, consisting of a wide variety of Gram-positive and Gram-negative bacteria, oral biofilms accumulate through sequential and ordered colonization by different strains and species present in the oral cavity.7

Bacterial adhesion depends on the properties of the bacterial cell and substratum surfaces. Under clinical conditions, surface roughness is the overruling property of any material placed in the oral cavity with respect to bacterial adhesion and biofilm formation, especially in supra-gingival regions where sizeable detachment forces are operative during the day.8 Roughness is of less importance in relatively stagnant regions, such as in sub-gingival pockets and here substratum hydrophobicity plays a major role.

Oral biofilm in orthodontic patientsPlacement of an orthodontic appliance consisting of metals and polymers, is accompanied by the creation of surfaces with properties, alien to the those of the natural oral hard and soft surfaces. In addition, the number of retention sites is much larger in orthodontic patients. These special features not only increase the amount of biofilm, but also the prevalence of cariogenic bacteria such as mutans streptococci9 and periodontopathic bacteria such as Porphyromonas gingivalis, Prevotella intermedia, Prevotella nigrescens, Tannerella forsythia, and Fusobacterium species.10 Moreover, orthodontic appliances greatly reduce the efficacy of natural oral cleansing forces and of mechanical biofilm removal by toothbrushing.11

The variety of alien surfaces introduced by orthodontic intervention provides numerous additional surfaces to which microorganisms can adhere and form a biofilm. Banding induced more biofilm formation mostly at the gingival margin, periodontal inflammation and white spot lesions than bonding.12 Composite bonding resins are prone to bacterial adhesion at the vulnerable bracket-adhesive-enamel junction, especially since polymerization shrinkage may yield a gap at the contact interface where bacteria find themselves protected against oral cleansing forces and antibacterial agents.13 Moreover, bacterial adhesion forces to composite resin, often having a rougher surface than enamel or brackets, were stronger than to brackets or saliva-coated enamel.14

Initial biofilm formation in vivo has been observed on different bracket materials.15 Brackets placed maxillary or at labial surfaces harvested more biofilm than at mandibular or lingual ones.16 Although more anaerobic and aerobic organisms have been found in self-ligating than in conventional bracket sites,17 the occurrence of white spot enamel lesions and gingival inflammation was similar in both patient groups,18 indicating that biofilm formation on the brackets themselves is less harmful than when formed at the bracket-adhesive-enamel junction.

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No difference was found regarding biofilm weight or biofilm-related clinical indices between different ligating devices. However, use of elastomeric rings was related to a higher incidence of enamel demineralization.19 In general, complicated auxiliaries create areas difficult to clean and enhancing biofilm formation.11

Removable acrylic retainers stimulate early biofilm formation, harvesting different strains of streptococci and candida, and provide new retention sites favoring bacterial adhesion and growth.20 Fixed retainers in direct contact with the enamel surface cannot be removed for extensive cleaning and may yield extensive biofilm formation.21 No differences were found in the clinical plaque and gingivitis indices between fixed retainers made of multi-strand or single-strand wires, but more biofilm was isolated from the multi-strand wires having niches where biofilms can be easily form and are protected against environmental attacks22 (Fig. 3).

Complications arising from biofilms during orthodontic treatmentEnamel demineralization Enamel demineralization surrounding brackets is the most common side-effect in orthodontics and can range from white spot lesions to cavitation upon bracket removal (Fig. 4). This can occur on both vestibular and lingual surfaces, with the most affected sites being the bracket-adhesive-enamel junction on teeth at the esthetic region.14 Enamel remineralization of white spot lesions can be achieved spontaneously by saliva or actively by fluoride or calcium-phosphate-based remineralization.23 Whether complete remineralization occurs or not is related to the type and severity of the lesions.11 White spot lesions can develop rapidly in susceptible individuals within the first month of treatment, and can remain visible many years after debonding, or in severe cases appear as a permanent enamel

Figure 3. Scanning electron micrograph of a multi-strand wire used for fixed retainers. Biofilm formation in the niches between the wires is clearly visible.

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scar.11 Fast developing or soft lesions are mostly superficial enamel defects and may almost completely remineralize within a few weeks. In most patients, lesions develop gradually during treatment and remineralize extremely slowly. Micro-abrasion, in essence an invasive method removing sound as well as diseased tissue, is an effective professional, cosmetic measure to treat permanent enamel scarring,24 which may also take place spontaneously leading to a gradual regression of the white spot lesion. More severely, white spot lesions may turn into actual cavities and not seldom orthodontic appliances have to be removed before the treatment goal has been reached to prevent further demineralization. The long term presence of white spot lesions or of composite restorations at labial surfaces of teeth, with the potential to turn into cavities or discolor respectively, are the most prevalent biofilm-related complications in orthodontics, compromising facial esthetics after an often lengthy and costly orthodontic treatment.

Soft tissue inflammation Almost all orthodontic patients experience some degree of soft tissue inflammation (Fig. 4). Gingivitis during orthodontic treatment is often temporary and rarely progresses to periodontitis, although biofilms on retention sites increase the risk for periodontitis. Biofilms on temporary anchorage devices (Fig. 5), such as mini-screws, micro-implants, or mini-plates, can cause inflammation of surrounding soft tissues similar to peri-implantitis, especially on trans-gingival parts of the devices. These inflammations are associated with a 30% increase in failure rate of the devices.25 In addition, biofilms on the head of a temporary anchorage device may infect adjacent contacting mucosa resulting in aphthous ulceration forewarning a greater soft tissue inflammation.26 Treatment of gingivitis or peri-implantitis in orthodontics includes local cleaning, application of antimicrobial containing products, such as chlorhexidine, cetylpyridinium chloride or triclosan preferably combined with brushing with a fluoridated toothpaste.26

Figure 4. White spot lesions, cavities (upper dentition) and gingival inflammation (lower dentition) caused by orthodontic biofilms after removal of fixed orthodontic appliance.

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Other consequences of orthodontic biofilms Bacteremia caused by trauma during appliance placement or removal, is usually transient and occurs with an incidence of up to 10% during fixed appliance treatment27 and 30% at removal of fixed expansion appliances.28 Biofilms may also affect the appliance itself and cause pitting and crevice corrosion of metallic biomaterials, affect mechanical properties, surface roughness or topographies of composite adhesives.29 Increase in roughness of the appliance materials due to biofilm is especially troublesome, since rougher surfaces promote biofilm formation,30 providing protective niches against environmental challenges. Hence a vicious cycle develops in which biofilm formation amplifies itself and may eventually compromise the efficiency of clinical mechanics.31

Occurrence of biofilm-related complicationsTable 1 summarizes the occurrence of biofilm-related complications during orthodontic treatment. Noticeably, large differences exist in reported occurrences of the major complications possibly relating to the various patient compliances that will greatly affect the study outcome, but are not systematically recorded in all studies. In a study on the prevalence of white spot lesions in 19-year-olds, only 23% of all participants showed good compliance with oral hygiene instructions, while 77% had moderate or poor compliance.32

Based on Table 1, it can be concluded that white spot lesions are a very common biofilm-related complication during orthodontic treatment, with a conservative estimate of the occurrence of 60%. Severe lesions requiring professional attention develop in up to 15% of all patients.33

Figure 5. Biofilms on and around temporary anchorage devices causing soft tissue inflammation.(A): Gingival inflammation (black arrow) around a temporary anchorage device (see white arrow). (B) and (C): Scanning electron micrographs of biofilm formed on a temporary anchorage device at low (B) and high magnification (C).

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Interproximal caries development is not significantly different from untreated controls34 and periodontitis is virtually absent.35 Gingivitis, often combined with gingival hyperplasia, is very common after orthodontic treatment but normally requires no treatment because of its transient nature.36

Year Number of patients

WSL (%)

Severe WSL requiring treatment (%)a

Fluorideaddition

Evaluation method

Reference

1982 121 50 7 No Visual Gorelick et al.57

1982 269 84 Not reported No Visual Mizrahi et al.58

1986 60 59 Not reported Yes Visual Artun and Brobakken59

1988 34 34 5 Yes Visual Geiger60

1989 51 96 10 Yes Visual Øgaard32

2005 64 97 Not reported No QLFb Boersma et al.61

2007 53 94 3 Yes PAc Lovrov et al.62

2010 332 36 14 No PA Chapman et al.63

2011 72 46 Not reported No Visual Tufekci et al.64

2011 400 61 15 Yes PA Enaia et al.33

2012 40 60 0 Yes Visual Hadler-Olsen et al.34

2012 64 43 Not reported No Visual Lucchese et al.65

2013 885 23 Not reported No PA Julien et al.66

The number of biofilm-related complications developing during orthodontic treatment is high. Considering the size of the current patient population, the results of this review indicate that 3 million orthodontic patients in the US alone develop white spot lesions as a result of the treatment. Up to 750,000 of these patients require professional care after orthodontic treatment. We estimate that basic treatment of white spot lesion on teeth in the esthetic region costs at least US$ 650 per patient37 adding up to nearly US$ 500,000,000 for all patients requiring professional care after orthodontic treatment. Since at least 2-3 hours are needed per patient, the total amount of man hours involved in these restorative treatments is estimated to be around 2,000,000. This means that every year around 1000 dentists have to work full time in order to treat the consequences of biofilm-related complications after orthodontic treatment. Although most orthodontists are aware of these problems, effective preventive programs and focussed research efforts are lacking.

a percentage of total number of patients;b quantitative light-induced fluorescence; c photographic assessment.

Table 1. Overview of reported occurrences of white spot lesions (WSL) during orthodontic treatment, according to different studies over the past three decades.

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Traditional and current preventive measuresMechanical removal Effective manual or powered brushing and use of interdental brushes is still by far the most important measure for oral hygiene control in orthodontic patients. Manual toothbrushes with a special head design for orthodontics, such as staged, v-shaped, or triple-headed, are more efficient than brushes with a conventional planar bristle field.38 Powered toothbrushes for removing biofilms are difficult to compare because of the diversity of frequencies or types of vibration, areas or types of bristle, and criteria or methods for assessment,39 but are generally accepted to perform better than manual brushing. However, the use of powered toothbrushes demonstrating non-contact removal (“cleaning beyond the bristles”) of oral biofilm40 up to brushing distances of 6 mm, depending on the energy output and frequency of the brush,41 may be advisable for orthodontic patients, although a thorough evaluation of the use of such brushes has never been made.

Chemical biofilm control A variety of chemical biofilm control measures including incorporation of antimicrobials in toothpastes, mouthrinses, varnishes and adhesives are currently used. Chlorhexidine however, still remains the most effective antimicrobial in reducing biofilm-related complications in orthodontic patients,42 although compliance may not be optimal in many patients since long-term use of chlorhexidine is known to stain teeth and tongue and affect taste sensation. Cetylpyridinium chloride is also an effective oral antimicrobial, but in many formulations its bio-availability is low. The benefits of fluoride containing toothpastes and mouthrinses in preventing caries have been well established and besides aiding enamel remineralization, fluoride acts as a buffer to neutralize acids produced by bacteria and suppresses their growth.30 Stannous fluoride provides dual benefits with respect to caries and biofilm prevention by stannous ions.43 The combination of an aminefluoride/stannous fluoride containing toothpaste or mouthrinse showed greater inhibition of biofilms, less white spot lesions and gingivitis during orthodontic treatment than sodium fluoride containing products.11 Laser irradiation in addition to fluoride treatment has been suggested to prevent formation of white spot lesions both in vitro and in vivo.44

Recently, it has been demonstrated that oral biofilm left-behind after brushing, absorbs antibacterial components from mouthrinses used after brushing to act as a reservoir for antibacterial components, that are subsequently slowly released in bioactive concentrations.45 Importantly, biofilm is always left-behind where it appears most harmful to the enamel surface, in case of orthodontic treatment around brackets. Consequently, slow release of absorbed antibacterial components from biofilm left-behind occurs where it matters most.

Modification of orthodontic materials Fluoride has been incorporated into various orthodontic adhesives46 to yield a slow release system with direct, beneficial clinical effects on enamel de- and remineralization. Other fluoride applications, which have not yet found their way to

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extensive clinical use, include coating of brackets and wires e.g. titanium tetrafluoride or calcium fluoride,47 demonstrating sustained release of fluoride and associated reductions in lesion depths and total mineral loss around the bracket-adhesive-enamel junction. Fluoride-containing elastomeric rings have also been demonstrated to release significant amounts of fluoride with a concurrent clinical reduction in the degree of decalcification around brackets,48 although the number of Streptococcus mutans or anaerobic bacterial growth in saliva or biofilms surrounding the brackets remained the same.49

Incorporation of antimicrobial agents in adhesives is more directly aimed at biofilm prevention. Antimicrobial release kinetics depend on the solubility of the antimicrobial in water, while the build-up of sufficiently high concentrations preventing microbial growth in saliva may be impossible due to wash-out in vivo. The solubility of chlorhexidine and triclosan in water for instance, is low and their release from adhesives may be less than required to reach a minimal inhibitory concentration preventing microbial growth.50 The release of cetylpyridinium chloride in water from adhesives showed a burst release during the first two weeks, followed by a much lower tail-release and in vitro caused an inhibition zone on bacterially inoculated agar. Other antimicrobials as e.g. benzalkonium chloride are only effective for two weeks after an initial burst release. Silver nanoparticles and quaternary ammonium polyethylenimine nanoparticles mixed into adhesives with an antibacterial activity upon contact are preferred since they are long-lasting,51 but the safety of nanoparticles for human use is still a matter of controversy.

Efforts required to prevent biofilm-related complicationsOrthodontists should first of all inform patients adequately about the potential risks of treatment and emphasize preventive programs. Especially adult patients can be made aware, better than juveniles, of the importance of oral hygiene. As an essential part of a preventive program, patients should be encouraged towards a more intensive oral hygiene control and use of powered toothbrushes, in combination with fluoridated, antibacterial toothpastes and antimicrobially effective mouthrinses, not solely aimed at creating fresh breath. Efforts to determine the possible clinical importance of non-contact, powered brushing in orthodontic patients should be undertaken.

Materials-related efforts currently focus on the development of antimicrobial releasing adhesives to fix brackets to tooth surfaces which will protect the vulnerable bracket -adhesive- enamel junction against biofilm formation, but it is doubtful whether clinically the small volume of adhesive applied to fix a bracket will be an effective reservoir for any antimicrobial over the duration of an average orthodontic treatment. Considering the duration of orthodontic treatment, more permanent non-adhesive or antimicrobial coatings that kill bacteria upon contact are preferable. However, neither low surface free energy polytetrafluoroethylene

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coatings on brackets52 nor polymer brush-coatings53 to discourage bacterial adhesion or photocatalytic TiO2 on wires54 to discourage bacterial growth have yet found their way toward clinical application. Alternative directions include modified composites with antimicrobial surface properties that kill bacteria upon adhesion.55 Recently, polymerization of antimicrobial cross-linked quaternary ammonium polyethylenimine nanoparticles into composite matrix has been demonstrated to significantly prevent oral biofilm formation in vivo and exhibit a potent broad spectrum antibacterial activity against salivary bacteria.56 Contact-killing coatings may have greater potential for the future than antimicrobial-release coatings as their efficacy is not hampered over time by a reduced release rate of antimicrobials from a reservoir with a limited volume.

CONCLUSIONS

The number of patients at risk of biofilm-related complications, including white spot lesions, caries and gingivitis has increased tremendously over the past two decades as a result of the success of orthodontic intervention to restore function and facial esthetics and now encompasses sizeable juvenile and adult populations. Based on this study, a conservative estimate of 60% of all orthodontic patients acquires one or more biofilm-related complications as a result of orthodontic treatment. Fixed braces and other orthodontic appliances hamper the maintenance of oral hygiene and provide numerous additional surfaces in the oral cavity to which bacteria can adhere and form a biofilm. With the growing demand for orthodontic treatment and a high occurrence of oral biofilm-related complications requiring professional care, orthodontic treatment is at risk of becoming a public health threat requiring improved preventive measures, including information for patients, effective personal oral care products like powered toothbrushes demonstrating non-contact removal of biofilms, pastes and rinses and the development of antimicrobial materials, preferentially contact-killing rather than materials relying on limited release of antimicrobials overtime. Only through concerted action, we will be able to prevent biofilm-related complications during orthodontic treatment from overshadowing it’s obvious advantages.

ACKNOWLEDGMENTS

This study has been supported by the Departments of Orthodontics and BioMedical Engineering, University Medical Centre Groningen, University of Groningen, the Netherlands.

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CONFLICT OF INTEREST:

H.J. Busscher is also director of a consulting company, SASA BV (GN Schutterlaan 4, 9797 PC Thesinge, The Netherlands). The authors declare no potential conflicts of interest with respect to authorship and/or publication of this article. Opinions and assertions contained herein are those of the authors and are not construed as necessarily representing views of the funding organizations or their respective employers.

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Dentofacial Orthop 140:e17-e24 34. Hadler-Olsen S, Sandvik K, El-Agroudi MA, Ogaard B (2012) The incidence of caries and white spot lesions in orthodontically treated adolescents with a comprehensive caries prophylactic regimen--a prospective study. Eur J Orthod 34:633-639 35. Bollen A, Cunha-Cruz J, Bakko DW, Huang GJ, Hujoel PP (April 2008) The effects of orthodontic therapy on periodontal health: A systematic review of controlled evidence. J Am Dent Assoc 139:413-422 36. Renkema AA, Dusseldorp JK, Middel B, Ren Y (2010) Enlargement of the gingiva during treatment with fixed orthodontic appliances. Ned Tijdschr Tandheelkd 117:507-512 37. Tariefbeschikking Tandheelkundige zorg. Nederlandse Zorg Autoriteit. http://www.nza.nl/98174/139255/654366/TB-CU-7042-02.pdf 38. Rafe Z, Vardimon A, Ashkenazi M (2006) Comparative study of 3 types of toothbrushes in patients with fixed orthodontic appliances. Am J Orthod Dentofacial Orthop 130:92-95 39. Schätzle M, Sener B, Schmidlin PR, Imfeld T, Attin T (2010) In vitro tooth cleaning efficacy of electric toothbrushes around brackets. Eur J Orthod 32:481-489 40. Schmidt JC, Zaugg C, Weiger R, Walter C (2013) Brushing without brushing?--a review of the efficacy of powered toothbrushes in noncontact biofilm removal. Clin Oral Investig 17:687-709 41. Busscher HJ, Jager D, Finger G, Schaefer N, Van der Mei HC (2010) Energy transfer, volumetric expansion, and removal of oral biofilms by non-contact brushing. Eur J Oral Sci 118:177-182 42. Sari E, Birinci I (2007) Microbiological evaluation of 0.2% chlorhexidine gluconate mouth rinse in orthodontic patients. Angle Orthod 77:881-884 43. Wiegand A, Bichsel D, Magalhães AC, Becker K, Attin T (2009) Effect of sodium, amine and stannous fluoride at the same concentration and different pH on in vitro erosion. J Dent 37:591-595 44. Noel L, Rebellato J, Sheats RD (2003) The effect of argon laser irradiation on demineralization resistance of human enamel adjacent to orthodontic brackets: an in vitro study. Angle Orthod 73:249-258 45. Otten MP, Busscher HJ, Van der Mei HC, Abbas F, Van Hoogmoed CG (2010) Retention of antimicrobial activity in plaque and saliva following mouthrinse use in vivo. Caries Res 44:459-464

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46. Chin MYH, Sandham A, Rumachik EN, Ruben JL, Huysmans MDNJM (2009) Fluoride release and cariostatic potential of orthodontic adhesives with and without daily fluoride rinsing. Am J Orthod Dentofacial Orthop 136:547-553 47. Lee S, Kim H, Kong Y, Kim H, Lee S, Chang Y (2005) Fluoride coatings on orthodontic wire for controlled release of fluorine ion. J Biomed Mater Res Part B: Applied Biomaterials 75B:200-204 48. Banks P, Chadwick S, Asher-McDade C, Wright J (2000) Fluoride-releasing elastomerics - a prospective controlled clinical trial. Eur J Orthod 22:401-407 49. Benson PE, Douglas CWI, Martin MV (2004) Fluoridated elastomers: Effect on the microbiology of plaque. Am J Orthod Dentofacial Orthop 126:325-330 50. Sehgal V, Shetty VS, Mogra S, Bhat G, Eipe M, Jacob S, Prabu L (2007) Evaluation of antimicrobial and physical properties of orthodontic composite resin modified by addition of antimicrobial agents—an in vitro study. Am J Orthod Dentofacial Orthop 131:525-529 51. Mei L, Ren Y, Loontjens TJ, Van der Mei HC, Busscher HJ (2012) Contact-killing of adhering streptococci by a quaternary ammonium compound incorporated in an acrylic resin. Int J Artif Organs 35:854-863 52. Demling A, Elter C, Heidenblut T, Bach F, Hahn A, Schwestka-Polly R, Stiesch M, Heuer W (2010) Reduction of biofilm on orthodontic brackets with the use of a polytetrafluoroethylene coating. Eur J Orthod 32:414-418 53. Roosjen A, De Vries J, Van der Mei HC, Norde W, Busscher HJ (2005) Stability and effectiveness against bacterial adhesion of poly(ethylene oxide) coatings in biological fluids. J Biomed Mater Res Part B: Applied Biomaterials 73B:347-354 54. Chun MJ, Shim E, Kho EH, Park KJ, Jung J, Kim JM, Kim B, Lee KH, Cho DL, Bai DH, Lee SI, Hwang HS, Ohk SH (2007) Surface modification of orthodontic wires with photocatalytic titanium oxide for its antiadherent and antibacterial properties. Angle Orthod 77:483-488 55. Tiller JC, Liao C, Lewis K, Klibanov AM (2001) Designing surfaces that kill bacteria on contact. Proc Natl Acad Sci 98:5981-5985

56. Beyth N, Yudovin-Farber I, Perez-Davidi M, Domb AJ, Weiss EI (2010) Polyethyleneimine nanoparticles incorporated into resin composite cause cell death and trigger biofilm stress in vivo. Proc Natl Acad Sci 107:22038-22043 57. Gorelick L, Geiger AM, Gwinnett AJ (1982) Incidence of white spot formation after bonding and banding. Am J Orthod 81:93-98 58. Mizrahi E (1982) Enamel demineralization following orthodontic treatment. Am J Orthod 82:62-67 59. Årtun J, Brobakken BO (1986) Prevalence of carious white spots after orthodontic treatment with multibonded appliances. Eur J Orthod 8:229-234 60. Geiger AM, Gorelick L, Gwinnett AJ, Griswold PG (1988) The effect of a fluoride program on white spot formation during orthodontic treatment. Am J Orthod Dentofacial Orthop 93:29-37 61. Boersma JG, Van der Veen MH, Lagerweij MD, Bokhout B, Prahl-Andersen B (2005) Caries prevalence measured with QLF after treatment with fixed orthodontic appliances: influencing factors. Caries Res 39:41-47 62. Lovrov S, Hertrich K, Hirschfelder U (2007) Enamel demineralization during fixed orthodontic treatment - Incidence and correlation to various oral-hygiene parameters. J Orofac Orthop 68:353-363 63. Chapman JA, Roberts WE, Eckert GJ, Kula KS, González-Cabezas C (2010) Risk factors for incidence and severity of white spot lesions during treatment with fixed orthodontic appliances. Am J Orthod Dentofacial Orthop 138:188-194 64. Tufekci E, Dixon JS, Gunsolley JC, Lindauer SJ (2011) Prevalence of white spot lesions during orthodontic treatment with fixed appliances. Angle Orthod 81:206-210 65. Lucchese A, Gherlone E (2012) Prevalence of white-spot lesions before and during orthodontic treatment with fixed appliances. Eur J Orthod doi:10.1093/ejo/cjs07066. Julien KC, Buschang PH, Campbell PM (2013) Prevalence of white spot lesion formation during orthodontic treatment. Angle Orthod 83:641-647

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Chapter 3Biofilm formation on stainless steel and gold wires

for bonded retainers in vitro and in vivo and their

susceptibility to oral antimicrobials

Marije A. Jongsma and Floris D.H. Pelser, Henny C. van der Mei, Jelly Atema-Smit, Betsy van de Belt-Gritter, Henk J. Busscher, Yijin Ren.


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Biofilm formation on retention wiresChapter 3

ABSTRACT

Objective Bonded retainers are used in orthodontics to maintain treatment result. Retention wires are prone to biofilm formation and cause gingival recession, bleeding-on-probing and increased pocket depths near bonded retainers. In this study we compare in vitro and in vivo biofilm formation on different wires used for bonded retainers and the susceptibility of in vitro biofilms to oral antimicrobials.

Materials and Methods Orthodontic wires were exposed to saliva and in vitro biofilm formation was evaluated using plate counting and live-dead staining, together with effects of exposure to toothpaste slurry alone or followed by antimicrobial mouthrinse application. Wires were also placed intra orally for 72 h in human volunteers and undisturbed biofilm formation was compared by plate counting and live-dead staining as well as by Denaturing Gradient Gel Electrophoresis for compositional differences in biofilms.

Results Single-strand wires attracted only slightly less biofilm in vitro than multi-strand wires. Biofilms on stainless-steel single-strand wires however, were much more susceptible to antimicrobials from toothpaste slurries and mouthrinses than on single-strand gold wires and biofilms on multi-strand wires. Also in vivo significantly less biofilm was found on single-strand than on multi-strand wires. Microbial composition of biofilms was more dependent on the volunteer involved than on wire type.

Conclusions Biofilms on single-strand stainless steel wires attract less biofilm in vitro and are more susceptible to antimicrobials than on multi-strand wires. Also in vivo, single-strand wires attract less biofilm than multi-strand ones.

Clinical Significance Use of single-strand wires is preferred over multi-strand wires, not because they attract less biofilm, but because biofilms on single-strand wires are not protected against antimicrobials as in crevices and niches as on multi-strand wires.

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INTRODUCTION

In the last decades, an increasing number of patients are being treated with orthodontic appliances. After an active orthodontic treatment, patients are often given a fixed retainer to prevent teeth from relapsing to their pre-treatment positions. Before the 1970s, fixed retainers were normally banded to the lower canines, but in the early 1970s the first report was published on the use of an acid-etching technique to bond retainers to the lingual surfaces of the lower canines.1 Since then, plain stainless steel round or rectangular retention wires have been used as bonded fixed retainers.1,2 In the early 1980s, the use of multi-strand wires was described. First, these retention wires were bonded only to the canines,3 while later multi-strand wires were bonded to all six front teeth.4 The twist in the multi-strand wires provided additional flexibility which allowed physiologic movement of the bonded teeth instead of fixing them all as one unit, and also provided undercut areas for mechanical retention for the composite bonding material.3-5

Despite the advantage of retainers in preventing teeth from relapsing to their pre-treatment position, the general drawback of retainers is that biofilm and calculus accumulate along the wires of lingually bonded retainers,6 yielding a greater incidence of gingival recession, increased pocket depth and bleeding on probing.7,8 Commonly used preventive measures, including toothbrushing, the use of antibacterial toothpastes, possibly supplemented with the use of antibacterial mouthrinses are generally not enough to adequately clean retainer sites, which is despite the generally favourable effects of antibacterial toothpastes and mouthrinses on plaque inhibition in vivo.9-12

Oral biofilm formation depends on the surface characteristics of the substratum surfaces, but also on the amount of surface area exposed to the oral environment. Multi-strand retention wires have crevices and therewith possess a larger surface area than single-strand wires, which can be expected to yield increased biofilm formation. Thick oral biofilms have been found on gold surfaces in vivo, but these were barely viable.13 Therefore the use of gold-coated wires for fixed bonded retainers has been advocated over the use of stainless steel wires.14 However, controversial results exist in the literature with respect to biofilm formation on different types of bonded retainers.6,15-17 This may be related to the fact that in previously published in vivo studies biofilm formation was not evaluated on the retention wires themselves but on the tooth surface surrounding the wires. However, a standardized in vitro study on biofilm formation on wires themselves should clarify this controversy.

The aim of this study was to compare in vitro and in vivo biofilm formation on different gold or stainless steel wires with different numbers of strands used for orthodontic bonded retainers and the susceptibility of in vitro formed biofilms on these retainers for chemical plaque

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control measures, i.e. exposure to toothpaste slurry, possibly followed by exposure to an antimicrobial mouthrinse.

MATERIALS AND METHODS

Retainers, toothpaste and mouthrinseFive types of orthodontic wires used for bonded retainers were evaluated in this study, as summarized in Table 1. Lengths of three cm were cut out of each wire type and sterilized with 70% ethanol. For plaque control, a NaF-sodium lauryl sulphate containing toothpaste without antibacterial claims was commercially obtained and 25 wt% slurries were prepared in sterilized distilled water after centrifugation to remove abrasion particles. Cool Mint Listerine® was also commercially purchased for use as an antimicrobial mouthrinse (Johnson and Johnson, New Jersey, USA).

Table 1. Overview of the orthodontic retention wires used in this study.Wire type Diameter Material Filament Manufacturer

Forestanit® 0.020 inch(0.5080 mm)

Stainless steel

single-strand Forestadent, Pforzheim, Germany

RW028 0.028 inch(0.7112 mm)

Gold single-strand Gold’n Braces, Inc., Palm Harbor, Florida, USA

Wildcat® 0.0175 inch(0.4445 mm)

Stainless steel

triple-strand Dentsply GAC Int., Bohemia, New York, USA

Quadcat®(rectangular)

0.016 x 0.022 inch(0.4064 x 0.5588 mm)

Stainless steel

triple-strand PG Supply, Inc., Avon, Connecticut, USA

Pentacat® 0.0175 inch(0.4445 mm)

Stainless steel

six-strands Dentsply GAC Int., Bohemia, New York, USA

Saliva collection and biofilm formation in vitroHuman whole saliva from five healthy volunteers of both sexes was collected into ice-chilled beakers after stimulation by chewing Parafilm. The saliva was pooled and sonicated on ice-chilled water for three times 10 s with 30 s intervals. All volunteers gave their informed consent to saliva donation, in agreement with the rules set out by the Ethics Committee at the University Medical Centre Groningen (February 6th, 2009).

A schematic protocol of the experiment is shown in Fig. 1. In one experiment four samples of each wire type were first placed in a sterile plastic tube containing 4 mL fresh pooled human saliva to allow bacterial adhesion to the wire surface. The tubes were incubated for 4 h at 37°C in an aerobic incubator while shaking at 60 rpm. After 4 h, samples were removed from the saliva and rinsed in sterile water, while one sample was kept for bacterial enumeration. Three samples were individually placed in sterile plastic tubes with 6 mL Tryptone Soya Broth (TSB) and left to incubate under shaking for 48 h. After 48 h, the three samples were

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removed from the TSB and rinsed in sterile water, while again retaining one for bacterial enumeration. The two remaining samples were exposed to either a tooth paste slurry (2 min) or a tooth paste slurry followed by exposure to a mouthrinse (30 s) and rinsed once again. For reference, ground and polished enamel samples (surface roughness 7 nm, as determined by atomic force microscopy) were included as a reference. All in vitro experiments were done in four-fold for each wire type.

Biofilm formation in vivoFour stainless steel wires (Forestanit®, Wildcat®, Quadcat® and Pentacat®) were bonded on the palatal and buccal side of the first molar and the second premolar (see also Fig. 1) of eight healthy volunteers in agreement with the rules set out by the Ethics Committee at the University Medical Centre Groningen (June 23rd, 2011). Different types of retention wires

Figure 1. (A) Schematic description of the experimental protocol for biofilm growth in vitro and in vivo. All in vitro experiments were carried out in four-fold, while in vivo experiments were done in eight human volunteers. (B) Buccal placement of retainer wires for in vivo biofilm growth

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were randomly attached to the right and left side of the maxillary arch. Wires were pre-bend on a plaster model of the volunteers dentition and had a length of 1 cm between the points of attachment to the teeth and were sterilized in 70% ethanol before use. Volunteers were instructed not to brush or touch the wires with an interdental cleaning aid, while brushing the remainder of their dentition with a commercially obtained NaF-sodium lauryl sulphate containing toothpaste without antibacterial claims. No additional oral hygiene products were allowed. Wires were removed after 72 h and oral biofilm was collected from the buccal and palatal enamel, together with a saliva sample. The wires and biofilm collected were stored in an Eppendorf tube containing 1.0 mL filter sterile reduced transport fluid (RTF). Saliva samples were stored on ice.

Evaluation of in vitro and in vivo biofilmsFor enumeration, retention wires with adhering biofilm formed in vitro or in vivo, and oral biofilm collected from enamel and saliva samples in human volunteers were sonicated three times for 10 s with 30 s intervals in Eppendorf tubes containing 1.5 mL filter sterile reduced transport fluid (RTF) on ice chilled water, to disperse the adhering bacteria. Bacteria were enumerated in a Bürker-Türk counting chamber and ten-fold serial dilutions were prepared in RTF for each wire type and condition and 100 μL was plated onto non-selective blood agar plates. After seven days of anaerobic incubation at 37°C, the total numbers of colony forming units (CFU’s) were counted and expressed per unit wire length. In addition, the percentage viability of the biofilms was evaluated after live/dead staining (BacLightTM, Bacterial Vitality Kit, Molecular Probes Europe BV) of dispersed biofilms. Live/dead stain was prepared by adding 3 μL of SYTO®9/Propidium iodide (1:3) to 1 mL of sterile, demineralized water. 15 μL of the stain was added to 10 μL of the undiluted biofilm dispersion. After 15 min incubation in the dark, the number of live and dead bacteria were counted using a fluorescence microscope (Leica DM4000B, Leica Microsystems Heidelberg GmbH, Heidelberg, Germany) and expressed as a percentage viability. Scanning electron micrographs of in vitro and in vivo biofilms on wires were taken, as described below.

DGGE analysis of in vivo biofilmsAll samples of in vivo formed biofilms and saliva were stored at -80°C until use for PCR- Denaturing Gradient Gel Electrophoresis (DGGE) in order to compare the microbial compositions of the biofilms. For extraction of DNA, samples were thawed centrifuged for 5 min at 13,000 g (Eppendorf Centrifuge 5415D, Hamburg, Germany) and subsequently washed and vortexed with 200 μL TE-buffer (10 mM Tris-HCl, 1 mM EDTA pH 7.4), again followed by centrifuging for 5 min at 13,000 g. Next, the supernatant was removed and the pellet was subsequently placed in a microwave (500 W, 5 min), after which it was suspended in 50 μL TE-buffer, vortexed and placed on ice. The quality and quantity of DNA samples

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were measured with a NanoDrop® spectrophotometer (ND-1000, NanoDrop Technologies, Inc, Wilmington, DE, USA) at 230 nm. The final concentration of each DNA sample was adjusted to 100 ng DNA for PCR amplifications.

PCR was performed with a Tgradient thermocycler (Bio-rad I-cycler, GENOtronics BV, USA). For amplification of the 16S rRNA gene, the following bacterial primers were used: F357-GC (forward primer, 5’-GC clamp-TACGGGAGGCAGCAG-3’)18 containing a GC clamp (5’- CGCCCGCCGCGCCCCGCGCCCGGCCCGCCGCCCCCGCCCC-3’)19 to make it suitable for DGGE, and R-518 (reverse primer, 5’-ATTACCGCGGCTGCTGG- 3’).20 Twentyfive μL of each PCR mixture contained 12.5 μL PCR Master Mix (0.05 units/μL Taq DNA polymerase in reaction buffer, 4 mM MgCl2, 0.4 mM dATP, 0.4 mM dCTP, 0.4 mM dGTP, 0.6 mM dTTP (Fermentas Life Sciences)), 1 μL of both forward and reverse primer (1 μM), and 100 ng DNA (in a volume of 10.5 μL). The temperature profile included an additional denaturing step of 5 min at 94°C, followed by a denaturing step at 94°C for 45 s, a primer annealing step at 58°C for 45 s, an extension step at 72°C for 1 min and a final extension step of 72°C for 5 min. PCR products were analyzed by electrophoresis on a 2.0% agarose gel containing 0.5 μg/mL ethidium bromide.

DGGE of PCR products generated with the F357-GC/R-518 primer set was performed as described by Muyzer et al.,21 using system PhorU (INGENY, Goes, The Netherlands). The PCR products were applied on 8% (w/v) polyacrylamide gel in 0.5 X TAE buffer (20 mM Tris acetate, 10 mM sodium acetate, 0.5 mM EDTA, pH 8.3). The denaturing gradient consisted of 30 to 80% denaturant (100% denaturant equals 7 M urea and 37% formamide). Gels were poured using a gradient mixer. A 10 mL stacking gel without denaturant was added on top. Electrophoresis was performed overnight at 120 V and 60°C. Gels were stained with silver nitrate.19 Each DGGE gel was normalized according to a marker consisting of 7 reference species comprising common bacterial species associated with oral health and disease,20,22 and stored at 4°C. The reference strains were Lactobacillus sp., Streptococcus oralis ATCC 35037, Streptococcus mitis ATCC 9811, Streptococcus sanguinis ATCC 10556, Streptococcus salivarius HB, Streptococcus sobrinus ATCC 33478 and Steptococcus mutans ATCC 10449.23

Scanning electron microscopy Topography of the wires, in absence and presence of both in vitro and in vivo formed biofilms, were visualized using scanning electron microscopy (SEM). Wires were fixed overnight in 2% glutaraldehyde and post-fixed for 1 h with 1% osmiumtetroxide. After dehydration through a water-ethanol series, wires were incubated in tetramethylsilane and air–dried, the samples that contained biofilm were sputter-coated with a gold-palladium alloy, after which they were fixed on SEM-stub-holders using double-sided sticky carbon tape and visualized in a field

40


emission scanning electron microscope (FE-SEM), type 6301F (JEOL Ltd., Tokyo, Japan) at 2 kV with a working distance of 39 mm and a small spot size.

Statistical analysisData were analyzed with the Statistical Package for Social Sciences (Version 16.0, SPSS Inc., Chicago, IL, USA). A one-way analysis of variance (ANOVA) was used to compare the number of CFUs found and the percentage biofilm viability. A Bonferroni test was used for post-hoc multiple comparisons. Statistical significance was set at p < 0.05.

DGGE gel images were converted and transferred into a microbial database with GelCompar II, version 6.1 (Applied Maths). The similarities in bacterial composition of the different biofilms were analysed using a band based similarity coefficient and a non-weighted pair group method with arithmetic averages was used to generate dendograms indicating similarities in composition.24

RESULTS

Scanning electron micrographs of the five wires are shown in Fig. 2 and clearly show crevices and niches formed by the multi-strand wires that are absent on the single-strand wires. Furthermore, it can be seen that the roughness of the single-strand gold wire is higher than that of the single-strand stainless steel wire.

The numbers of colony forming units on the different wires formed in vitro are summarized in Table 2. There was no significant difference between the number of CFUs adhering to the wires after 4 h incubation in saliva, while both single-strand stainless steel and gold wires showed less biofilm formation after 48 h compared to the three stainless steel multi-strand wires. For the stainless steel single-strand wire, this difference was significant compared to all three multi-strand stainless steel wires (p < 0.05), but for the gold single-strand wire there was only a significant difference compared to the six-strands stainless steel wire (p < 0.05). There was no statistically significant difference in the amount of biofilm formation on the stainless steel versus the gold single-strand wires, neither were there any statistically significant differences between the three multi-strand stainless steel wires. All wires attract highly viable biofilms, with less than 20% dead bacteria. For comparison, we carried out a similar experiment on enamel surfaces, and found a similarly high viability of 74.0% ± 8.5% (note that the amount of biofilm formed on enamel could not be expressed in units allowing comparison with the amount of biofilm formed per cm wire length).

The single-strand wires attract a differently structured biofilm than the multi-strand wires (Fig. 3). On the multi-strand wires, biofilm is mostly located in the crevices between strands,

41

3


while on the single-strand wires bacteria are present as a thin film (compare Figs. 3A and B with Fig. 3C). Comparison of the biofilms on single-strand stainless steel versus gold wires gives the impression of a higher degree of clustering of the biofilm on gold, possibly as a result of its larger roughness (compare Fig. 3A with Fig 3B). In vitro results show similar viability for gold and stainless steel as well as for enamel after 48 h of biofilm formation.

Figure 2. Scanning electron micrographs of the different wire types prior to biofilm formation; magnification 75x, bar marker indicates 100 µm.(A) Forestanit® (single-strand, stainless steel), (B) RW028 (single-strand, gold),(C) Wildcat® (triple-strand, stainless steel),(D) Quadcat® (triple-strand, stainless steel) and (E) Pentacat® (six-strands, stainless steel).

42


Figure 3. Scanning electron micrographs of 48 h old biofilms formed in vitro on selected wire types; magnification 750x, bar marker indicates 10 µm. (A) Forestanit® (single-strand, stainless steel): biofilm is present as a thin, scattered film,(B) RW028 (single-strand, gold): scattered clusters of biofilm are formed,(C) Quadcat® (triple-strand, stainless steel): biofilm is mostly located in the crevices between strands.

The number of CFUs cultured from 48 h old in vitro biofilms on multi-strand wires was not significantly affected by exposure to toothpaste slurries, nor by exposure to toothpaste slurries followed by exposure to an antimicrobial mouthrinse, although significant drops in viability were observed. Oppositely, 48 h old biofilms on stainless steel and gold single-strand wires showed significantly reduced numbers of CFUs after exposure to the toothpaste supernatant concurrent with a drop in viability, while further reductions in amount of biofilm and viability could be achieved by subsequent exposure to the antimicrobial mouthrinse for single-strand stainless steel wires. Interestingly, biofilms on single-strand stainless steel wires were much more susceptible to chemical plaque control than biofilms formed on gold wires. For comparison, for biofilms formed on enamel surfaces, viability decreased from 74.0% ± 8.5% to 59.5% ± 0.7% upon exposure to a toothpaste slurry, dropping further down to 19.5% ± 7.7% upon subsequent exposure to an antimicrobial mouthrinse.

43

3


Tabl

e 2

The

num

ber o

f CFU

s (lo

g un

its ±

SD

ove

r fou

r exp

erim

ents

with

sep

arat

ely

cultu

red

bact

eria

) and

the

viab

ility

foun

d in

4 h

and

48

h ol

d bi

ofilm

s fo

rmed

in v

itro

from

fres

h, h

uman

who

le s

aliv

a on

1 c

m le

ngth

s of

the

diffe

rent

wire

s in

volv

ed in

this

stu

dy a

nd th

e ef

feac

ts o

f exp

osur

e to

a 2

5w%

to

othp

aste

slu

rry

alon

e, o

r fol

low

ed b

y an

add

ition

al e

xpos

ure

to a

mou

thrin

se.

Wire

type

4 h

biofi

lm48

h b

iofil

m48

h b

iofil

m e

xpos

ed to

to

othp

aste

slu

rry

48 h

bio

film

exp

osed

to

toot

hpas

te s

lurry

and

mou

thrin

se

CFU

s%

Live

CFU

s%

Live

CFU

s%

Live

CFU

s%

Live

Fore

stan

it®4.

1 ±

0.3

> 95

5.7

± 0.

387

.0 ±

8.4

1.6

± 1.

4c,d

10.0

± 4

.1d

0.3

± 0.

4c,d

0.0

± 0.

0d,e

RW02

84.

5 ±

0.2

> 95

6.0

± 0.

486

.7 ±

7.6

4.5

± 1.

2a,d

43.7

± 8

.0a,

d4.

9 ±

1.2a

20.0

± 1

1.3a,

d,e

Wild

cat®

4.2

± 0.

3>

956.

5 ±

0.3a,

c90

.5 ±

4.9

6.2

± 0.

1a,b,

c56

.0 ±

1.4

a,d

6.3

± 1.

2a,c

42.0

± 5

.7a,

b,d

Qua

dcat

®4.

7 ±

0.5

> 95

6.6

± 0.

8a,c

89.0

± 1

.46.

2 ±

0.2a,

b,c

45.5

± 1

0.6a,

d6.

2 ±

0.3a,

b,c

27.5

± 3

.5a,

d,e

Pent

acat

®4.

6 ±

0.2

> 95

6.9

± 0.

2a,b,

c82

.5 ±

7.8

6.5

± 0.

2a,b,

c,d

56.5

± 0

.7a,

d6.

6 ±

0.1a,

b,c

46.5

± 0

.7a,

b,d

a sig

nific

antly

diff

eren

t fro

m F

ores

tani

t® (s

ingl

e-st

rand

sta

inle

ss s

teel

)b s

igni

fican

tly d

iffer

ent f

rom

RW

028

(sin

gle-

stra

nd g

old)

c sig

nific

antly

diff

eren

t fro

m 4

hd s

igni

fican

tly d

iffer

ent f

rom

48

h w

ithou

t exp

osur

e to

toot

hpas

te s

lurry

or m

outh

rinse

e sig

nific

antly

diff

eren

t fro

m 4

8 h

with

exp

osur

e to

toot

hpas

te s

lurry

44


The number of CFUs in biofilms formed in vivo are summarized in Table 3. Buccally placed wires collected more biofilm than palatally placed wires, regardless of the wire type, while no significant differences were found between the different wires placed on the buccal side. Significantly less biofilm had grown on the single-strand stainless steel wire placed palatally compared to other palatally placed wires, while all multi-strand wires on the palatal side collected similar amounts of biofilm. All in vivo biofilms formed on the different retention wires contained a similar percentage of live bacteria (see Table 3), while oral biofilm collected from enamel surfaces in vivo was slightly less viable (64.2% ± 6.8% and 64.0% ± 6.4% for bucally and palatally sampled oral biofilm).

Table 3. The number of CFUs in and the viability of biofilms formed in vivo (log units ± SD over eight different volunteers) on 1 cm lengths of the different wires involved in this study*.

Wire typeBuccally placed Palatally placed

CFUs % Live CFUs % Live

Forestanit® 7.4 ± 0.3a 73.6 ± 6.9 6.7 ± 0.5 73.0 ± 13.1

Wildcat® 7.5 ± 0.2a 70.9 ± 14.5 7.2 ± 0.4a 75.0 ± 6.7

Quadcat® 7.6 ± 0.3a 73.5 ± 8.4 7.3 ± 0.5a 74.9 ± 9.9

Pentacat® 7.6 ± 0.1a 75.6 ± 6.5 7.3 ± 0.4a 74.2 ± 10.2* RW028 became unavailable during the course of the study and no in vivo data are availablea significantly different from Forestanit® placed palatally.

Scanning electron micrographs (Fig. 4) show that also in vivo the single-strand wires attract a differently structured biofilm than the multi-strand wires. On the multi-strand wires, biofilm is mostly located in the crevices between strands, while on the single-strand wires bacteria are present as a thin film. There is also a clear difference between wires placed buccally compared to wires placed palatally. Biofilm on buccally placed wires covers the entire wire surface, whereas smooth surfaces of palatally placed wires are either clean or only covered with a thin organic film. Biofilm on the multi-strand, palatally placed wires is almost entirely located in crevices and niches, while palatally placed single-strand wires collect biofilm mostly on the side of the wire facing the tooth surface and thus out of reach by the tongue.

Microbial composition of the in vivo formed biofilms on enamel was equally variable among volunteers (Fig. 5) as the variation in the composition of biofilms formed on different wire types. Microbial compositions of saliva from different volunteers had a tendency to cluster, but the composition of biofilms formed on the retainer wires, including enamel surfaces could not be related with a specific material or wire type.

45

3


Figure 4. Scanning electron micrographs of 48 h biofilms formed in vivo on selected wire types: magnification 75x, bar marker indicates 100 µm and 750x, bar marker indicates 10 µm.(A) Forestanit® (single-strand, stainless steel) buccally placed: biofilm is present as a thick fully covering film,(B) Forestanit® (single-strand, stainless steel) palatally placed: biofilm is present as a thin, scattered film, (C) Quadcat® (multi-strand, stainless steel) buccally placed: biofilm is present in the crevices between strands as well as on the smooth surfaces,(D) Quadcat® (multi-strand, stainless steel) palatally placed: biofilm is mostly located in the crevices between strands.

46


Figure 5. Dendograms of biofilms formed on different wire types and enamel and saliva, showing clustering of biofilms with a similar microbial composition. Numbers denote different volunteers.(A) buccal samples(B) palatal samples.

47

3


DISCUSSION

Although the use of the bonded retainers to prevent teeth from relapsing back to their original, pre-treatment position is generally accepted in orthodontics, the tendency of these retainers to collect oral biofilm and calculus is considered a disadvantage. It has long been suggested, that the increased numbers of retention areas in crevices and niches of multi-strand wires do not yield higher biofilm attraction than on single-strand wires.6 The present study for the first time confirms that there is indeed little difference in biofilm formation in vitro on single- versus multi-strand wires and differences were only statistically significant after 48 h of biofilm formation. Highly interesting, biofilms formed in vitro on multi-strand wires appear less susceptible to oral antimicrobials than biofilms on single-strand wires, probably because of their protected growth in crevices and niches on multi-strand wires. Likely, the protected growth in crevices and niches is the reason why in vivo more biofilm accumulated at the undercut areas of the multi-strand wires and on the surrounding lingual tooth surfaces than with single-strand wire retainers.17 On tooth surfaces, minute irregularities have been demonstrated to protect microorganisms and stimulate biofilm accumulation.25 Biofilm formation on surgical meshes and suture materials also demonstrate more biofilm formation on multi- than on mono-filament structures,26-28 with aerobic and anaerobic bacteria being isolated in nearly equal numbers of viable bacteria from monofilament sutures made of different materials used in intraoral dentoalveolar surgery.29

The lack of a significant difference in in vitro numbers of viable bacteria on single-strand retention stainless steel and gold wires in the current study indicates that the influence of the material on initial biofilm formation is low, which is in agreement with literature, stating that roughness is the dominant factor in biofilm adhesion.30 Five-days-old oral biofilms on gold surfaces in vivo are known to be thick and fully covering the substratum surfaces though with a viability less than 2%.13 Possibly, full coverage by a relatively thick biofilm hampers the supply of nutrients to the biofilm, leading to a low viability extending to the deeper layers of the biofilm,31 while allowing antimicrobials to remain active on the outer layer.32,33 The present in vitro study, though confined to 48 h, shows a larger clustering of bacteria on gold than on stainless steel, which may be considered as the on-set of a thick and fully covering biofilm. The difference can probably be attributed to the a higher surface roughness of gold wires compared to stainless steel single-strand wires (compare Figs. 3A and 3B), since a surface roughness above a threshold of 2 µm is already known to facilitate biofilm formation on restorative materials.34 The larger clustering of bacteria on gold probably offers protection against oral antimicrobials. Positive effects of antimicrobials, such as NaF, sodium lauryl sulphate in toothpastes and essential oil in mouthrinses on biofilm inhibition have been extensively described for oral biofilms on smooth surfaces.9-12 In the present study

48


however, exposure to toothpaste supernatant reduced biofilm formation only on both single-strand wires. Additional effects of the antimicrobial mouthrinse compared to the toothpaste supernatant were observed for the stainless steel single-strand wires. This supports the above suggestion that the increased roughness of gold wires compared with stainless steel ones as well the crevices and niches in multi-strand wires protect oral biofilm organisms against chemical challenges. Unfortunately, gold wires became unavailable during the course of this study, impeding inclusion of gold wires in our in vivo analysis. The current in vitro study has been carried out using biofilms grown from human whole saliva. Therewith, a larger number of strains can be grown from more controlled experiments using clinical isolates and the chances of bacteria to adhere are therefore increased.25 Moreover, biofilms grown from saliva are more representative of in vivo biofilms, whereas at the same time it may be considered a disadvantage that we had a lower control of the biofilm composition than when using single strains of bacteria.35

Despite differences between salivary protein adsorption in vitro and in vivo36 and possible differences in the selective growth of bacteria from saliva in vitro and in vivo, our in vivo comparison of biofilm formation on different retention wires confirms that on the palatal side less biofilm is formed on single-strand than on multi-strand retention wires, likely due to the protection offered by growth in crevices and niches of multi-strand wires against mechanical removal (brushing) and oral antimicrobials. Although volunteers did not brush the wire itself with toothpaste and used a toothpaste without antibacterial claims, it cannot be avoided that antimicrobials are involved in in vivo biofilm formation on the wires. The toothpaste used contains fluoride and sodium lauryl sulphate, both known to be antimicrobial, that spread through the oral cavity during brushing. Moreover, saliva contains several antimicrobial peptides and proteins that affect biofilm formation.37 In vivo protection against mechanical removal is furthermore implicated by the fact that no significant differences were observed between buccally placed wire types, but only for palatally placed ones, within reach of frictional removal forces exerted by the tongue.30 These results imply that for this type of research, buccal placement, though preferred by volunteers, is to be avoided in comparative studies on oral biofilm formation on retention wires, since differences only become evident under clinical conditions when wires are placed palatally.

Microbial compositions of saliva from different volunteers obtained using DGGE cluster more strongly than the compositions of the adhering biofilms in different volunteers. Moreover, no clustering is observed for the composition of biofilms formed on different retention wires, including enamel in different volunteers. This demonstrates that inter-individual differences control the composition of the oral microflora,38 for instance through dietary influences, difficult to standardize in any clinical study.24,39

49

3


CONCLUSIONS

Recent studies showed an increased incidence of lingual gingival recession, biofilm retention and bleeding on probing of teeth with bonded retainers.7,8 Based on the current results, it is concluded that single-strand wires attract only slightly less biofilm in vitro than multi-strand wires, with no significant difference between single-strand stainless steel and gold wires. In vivo however, single-strand, palatally placed stainless steel wires attracted significantly less biofilm compared to the other wires, indicating that with respect to biofilm formation and its prevention, single-strand stainless steel wires should be the first choice. Single-strand stainless steel wires attract less biofilm in vivo, not because they are less adhesive to oral biofilm, but because biofilms on single-strand retention wires is less protected by growth in crevices and niches against oral antimicrobials than when formed on multi-strand wires.

ACKNOWLEDGMENTS

The authors would like to extend their gratitude to Mr. Jeroen Kuipers from the Centre for Medical Electron Microscopy of the University Medical Centre Groningen for his assistance with the SEM analysis, Dr. James Huddleston Slater from Department of Oral and Maxillofacial Surgery of the University Medical Centre Groningen for his assistance with the statistical analysis and Ortho=solutions BV (Wijk bij Duurstede, The Netherlands), Dentsply Lomberg (Soest, The Netherlands) and Orthotec B.V. (Zeist, The Netherlands) for kindly providing the different wires used in this study.

50


REFERENCES

1. Knierim RW (1973) Invisible lower cuspid to cuspid retainer. Angle Orthod 43:218-220 2. Carter RN (1978) Simplified direct-bonded retainer. J Clin Orthod 12:221 3. Artun J, Zachrisson B (1982) Improving the handling properties of a composite resin for direct bonding. Am J Orthod 81:269-276 4. Zachrisson BU (1982) The bonded lingual retainer and multiple spacing of anterior teeth. Swed Dent J Suppl 15:247-255 5. Bearn DR (1995) Bonded orthodontic retainers: a review. Am J Orthod Dentofacial Orthop 108:207-213 6. Artun J (1984) Caries and periodontal reactions associated with long-term use of different types of bonded lingual retainers. Am J Orthod 86:112-118 7. Pandis N, Vlahopoulos K, Madianos P, Eliades T (2007) Long-term periodontal status of patients with mandibular lingual fixed retention. Eur J Orthod 29:471-476 8. Levin L, Samorodnitzky-Naveh GR, Machtei EE (2008) The association of orthodontic treatment and fixed retainers with gingival health. J Periodontol 79:2087-2092 9. Riep BG, Bernimoulin JP, Barnett ML (1999) Comparative antiplaque effectiveness of an essential oil and an amine fluoride/stannous fluoride mouthrinse. J Clin Periodontol 26:164-168 10. Arweiler NB, Auschill TM, Reich E, Netuschil L (2002) Substantivity of toothpaste slurries and their effect on reestablishment of the dental biofilm. J Clin Periodontol 29:615-621 11. Stoeken JE, Paraskevas S, Van der Weijden GA (2007) The long-term effect of a mouthrinse containing essential oils on dental plaque and gingivitis: a systematic review. J Periodontol 78:1218-1228 12. Pizzo G, La Cara M, Licata ME, Pizzo I, D’Angelo M (2008) The effects of an essential oil and an amine fluoride/stannous fluoride mouthrinse on supragingival plaque regrowth. J Periodontol 79:1177-1183 13. Auschill TM, Arweiler NB, Brecx M, Reich E, Sculean A, Netuschil L (2002) The effect of dental restorative materials on dental biofilm. Eur J Oral Sci

110:48-53 14. Zachrisson BU, Buyukyilmaz T (2005) Bonding in orthodontics. In: Graber TM, Vanarsdall RLJ, Vig KWL (eds) Orthodontics: Current Principles and Techniques, 4th edn. Elsevier Mosby, St. Louis, pp 579-659 15. Artun J, Spadafora AT, Shapiro PA, McNeill RW, Chapko MK (1987) Hygiene status associated with different types of bonded, orthodontic canine-to-canine retainers. A clinical trial. J Clin Periodontol 14:89-94 16. Artun J, Spadafora AT, Shapiro PA (1997) A 3-year follow-up study of various types of orthodontic canine-to-canine retainers. Eur J Orthod 19:501-509 17. Al-Nimri K, Al Habashneh R, Obeidat M (2009) Gingival health and relapse tendency: a prospective study of two types of lower fixed retainers. Aust Orthod J 25:142-146 18. Di Cagno R, Rizzello CG, Gagliardi F, Ricciuti P, Ndagijimana M, Francavilla R, Guerzoni ME, Crecchio C, Gobbetti M, De Angelis M (2009) Different fecal microbiotas and volatile organic compounds in treated and untreated children with celiac disease. Appl Environ Microbiol 75:3963-3971 19. Zijnge V, Welling GW, Degener JE, Van Winkelhoff AJ, Abbas F, Harmsen HJ (2006) Denaturing gradient gel electrophoresis as a diagnostic tool in periodontal microbiology. J Clin Microbiol 44:3628-3633 20. Marsh PD (1994) Microbial ecology of dental plaque and its significance in health and disease. Adv Dent Res 8:263-271 21. Muyzer G, De Waal EC, Uitterlinden AG (1993) Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 59:695-700 22. Marsh PD (2006) Dental plaque as a biofilm and a microbial community - implications for health and disease. BMC Oral Health 6 Suppl 1:S14 23. Otten MPT, Busscher HJ, Abbas F, Van der Mei HC, Van Hoogmoed CG (2012) Plaque-left-behind after brushing: intra-oral reservoir for antibacterial toothpaste ingredients. Clin Oral Investig DOI 10.1007/s00784-011-0648-2 24. Signoretto C, Bianchi F, Burlacchini G, Sivieri F, Spratt D, Canepari P (2010) Drinking habits are

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associated with changes in the dental plaque microbial community. J Clin Microbiol 48:347-356 25. Lang PL, Mombelli A, Attström R (1997) Dental plaque and calculus. In: Lindhe J, Karring T, Lang PL (eds) Clinical periodontology and implant dentistry, 3rd edn. Munksgaard, Copenhagen, pp 102-137 26. Engelsman AF, Van Dam GM, Van der Mei HC, Busscher HJ, Ploeg RJ (2010) In vivo evaluation of bacterial infection involving morphologically different surgical meshes. Ann Surg 251:133-137 27. Engelsman AF, Van der Mei HC, Ploeg RJ, Busscher HJ (2007) The phenomenon of infection with abdominal wall reconstruction. Biomaterials 28:2314-2327 28. Masini BD, Stinner DJ, Waterman SM, Wenke JC (2011) Bacterial adherence to suture materials. J Surg Educ 68:101-104 29. Otten JE, Wiedmann-Al-Ahmad M, Jahnke H, Pelz K (2005) Bacterial colonization on different suture materials - A potential risk for intraoral dentoalveolar surgery. J Biomed Mater Res Part B: Appl Biomater

74B:627-635 30. Quirynen M, Bollen CM (1995) The influence of surface roughness and surface-free energy on supra- and subgingival plaque formation in man. A review of the literature. J Clin Periodontol 22:1-14 31. Busscher HJ, Rinastiti M, Siswomihardjo W, Van der Mei HC (2010) Biofilm formation on dental restorative and implant materials. J Dent Res 89:657-665 32. Marsh PD (2004) Dental plaque as a microbial biofilm. Caries Res 38:204-211 33. Ten Cate JM (2006) Biofilms, a new approach to the microbiology of dental plaque. Odontology 94:1-9 34. Teughels W, Van Assche N, Sliepen I, Quirynen M (2006) Effect of material characteristics and/or surface topography on biofilm development. Clin Oral Implants Res 17 Suppl 2:68-81 35. Fournier A, Payant L, Bouclin R (1998) Adherence of Streptococcus mutans to orthodontic brackets. Am J Orthod Dentofacial Orthop 114:414-417 36. Carlen A, Borjesson AC, Nikdel K, Olsson J (1998) Composition of pellicles formed in vivo on tooth

Chapter 4In vivo biofilm formation on stainless steel bonded-

retainers during different regimens of oral health care

Marije A. Jongsma, Henny C. van der Mei, Jelly Atema-Smit,

Henk J. Busscher, Yjijn Ren.

International Journal of Oral Science (2015) doi:10.1038/ijos.2014.69

54

Effects of antimicrobials on oral biofilmsChapter 4

ABSTRACT

Permanent bonded retention wires to anterior teeth are used after orthodontic treatment to prevent teeth from relapsing to pre-treatment positions. A drawback of bonded retainers is biofilm accumulation along the wires, yielding greater incidence of gingival recession, increased pocket depth and bleeding-on-probing.

This study compares in vivo biofilm formation on single-strand and multi-strand retention wires during different regimens of oral healthcare. Two-cm wires were placed in brackets bonded to the buccal side of first molars and second premolars in the upper arches of 22 volunteers. Volunteers used a selected toothpaste with or without additional use of an essential-oils containing mouthrinse. Brushing was performed manually. Regimens were maintained for one week, after which wires were removed and oral biofilm was collected for enumeration of the number of organisms and their viability, microbial composition and electron microscopic visualization. Six weeks wash-out was applied in between regimens. Less biofilm was formed on single-strand wires than on multi-strand wires, on which bacteria were observed adhering in between strands. Use of antibacterial toothpastes marginally reduced the amount of biofilm on both wire types, but viability of biofilm organisms was significantly reduced by use of antibacterial toothpastes. No significant effects were observed on amount or viability of biofilms upon additional use of the mouthrinse.

However, major shifts in biofilm composition were induced by combining a stannous-fluoride or triclosan containing toothpaste with the essential-oils containing rinse. Tentatively, these shifts are attributed to small changes in bacterial cell surface hydrophobicity after adsorption of toothpaste components, that stimulate bacterial adhesion to hydrophobic oil, as illustrated for a Streptococcus mutans strain.

55

4


INTRODUCTION

A major challenge in orthodontics is to retain treatment results after removal of orthodontic appliances. Long-term results of orthodontic treatment show relapse of crowding without use of retention devices.1 To prevent relapse, permanent retention wires are often bonded to the anterior teeth.2 Different types of retention wires can be used, including single-strand retainers, bonded only to the canines, or multi-strand retainers that are bonded to all six anterior teeth.3,4 The downside of placing retention wires is that biofilm and calculus accumulate along the wires, which may cause a greater incidence of gingival recession, increased pocket depth and bleeding on probing.5,6

Previous in vitro results have shown that wire morphology has an influence on the number of viable organisms in biofilm formed on retention wires.7 Biofilms pre-formed on single-strand wires harvested less viable organisms than biofilms formed on multi-strand wires after a single exposure to a NaF-sodium lauryl sulphate containing toothpaste slurry and an essential-oils containing mouthrinse, demonstrating that biofilms on multi-strand wires are less susceptible to oral antimicrobials than biofilms formed on single-strand wires. The biofilm mode of growth is indeed known to protect its inhabitants against penetration of antimicrobials agents,8 an effect that may be enhanced when the biofilm is formed in crevices and niches of a retention wire.9 It is unknown however, how these differences in the susceptibility of oral biofilms pre-formed on different wire morphologies in vitro, translate to biofilm formation in vivo during the use of antibacterial health care products, such as toothpastes or mouthrinses with antibacterial claims.

In the great majority of the population not all biofilm is removed by mechanical means, and as a consequence, despite the difficulty for antimicrobials to penetrate a biofilm, oral antimicrobials generally have a favourable effect on biofilm inhibition in vivo.10-13 Biofilm-left-behind after brushing, either dead or alive, can play an important role in making an antimicrobial action substantive as it can absorb antimicrobials to become released over time in antimicrobially effective amounts.14 It is unknown however, whether this is a mechanism that is clinically operative to a degree that it yields measurable effects on biofilm formation.

The aim of this study is to compare biofilm formation in vivo on both single-strand and multi-strand retention wires during different regimens of oral health care and to evaluate whether use of oral antimicrobials affects the composition of the biofilm. Regimens included manual brushing. Two different toothpastes with antibacterial claims15 were used, containing either stannous fluoride or triclosan or a fluoridated toothpaste without antibacterial claims. Toothpastes were employed with or without the additional use of an essential-oils containing mouthrinse.12

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Retainers, volunteers and inclusion criteriaTwo different types of retainers were evaluated in this study, a single-strand wire (Forestanit®, Forestadent, Pforzheim, Germany) and a multi-strand wire (Quadcat®, PG Supply, Inc., Avon, USA). Brackets (SPEED System Orthodontics, Cambridge, Canada) were bonded to the buccal side of the first molar and the second premolar in the upper arch of 22 healthy volunteers in agreement with the rules set out by the Ethics Committee at the University Medical Centre Groningen (letter June 23rd, 2011). Wires had a length of 2 cm between the brackets in which they were placed. The wires were sterilized in 70% ethanol before use and stayed in situ for one week during which the volunteers were instructed to brush for 2 min twice a day with a manual toothbrush (Lactona iQ X-Soft, Lactona Europe B.V., Bergen op Zoom, The Netherlands) and use a toothpaste with antibacterial claims (Oral-B Pro Expert®, Procter & Gamble, Cincinnati, USA or Colgate Total®, Colgate-Palmolive Company, Piscataway, USA) or a toothpaste without antibacterial claims that contains only NaF-sodium lauryl sulphate (Prodent Softmint®, Sara Lee Household & Bodycare, Exton, USA). Toothpastes were used either without additional oral hygiene measures or in combination with an essential-oils containing mouthrinse (Cool Mint Listerine®, Pfizer Consumer Healthcare, Morris Plains, NJ, USA).

In between regimens, a washout period of 6 weeks was applied during which only the NaF-sodium lauryl sulphate containing toothpaste without antibacterial claims was used. The duration of the washout period was based on the results of a pilot study which indicated that the composition of the oral biofilm returned to base line values within 5 weeks after use of an antibacterial toothpaste.

Volunteers were included in the study, provided that they had a healthy and complete dentition, no bleeding upon probing, did not use any medication and did not smoke. All volunteers granted a written informed consent. After inclusion, volunteers were randomly divided into two groups. The first group successively used 3 different types of toothpaste, the second group combined the same toothpastes with an antimicrobial mouthrinse (see Figure 1).

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Regimens were maintained for 1 week, after which wires were removed and oral biofilm was collected from the buccal enamel surfaces for reference using a cotton swab, while also unstimulated salivary samples were taken. The wires, collected enamel biofilms and salivary samples were stored in an Eppendorf tube containing 1.0 mL filter sterile reduced transport fluid.16 Saliva samples were stored on ice.

For enumeration of the numbers of organisms, retention wires with adhering biofilm, cotton swabs with oral biofilm collected from enamel, both stored in Eppendorf tubes containing 1.0 mL filter sterile reduced transport fluid and saliva samples were separately sonicated three times for 10 s with 30 s intervals on ice chilled water, to disperse bacteria. Bacteria were then enumerated in a Bürker-Türk counting chamber. In addition, the percentage viability of the biofilms was evaluated after live/dead staining (BacLightTM, Invitrogen, Breda, The Netherlands) of dispersed biofilms. Live/dead stain was prepared by adding 3 μL of SYTO®9/propidium iodide (1:3) to 1 mL of sterile, demineralised water. 15 μL of the stain was added to 10 μL of the undiluted biofilm dispersion. After 15 min incubation in the dark, the number of live and dead bacteria were counted using a fluorescence microscope (Leica DM4000B, Leica Microsystems Heidelberg GmbH, Heidelberg, Germany) and expressed as a percentage viability. Scanning electron micrographs of the biofilms on wires were taken, as described below.

DGGE analysis of in vivo biofilmsAll samples of in vivo formed biofilms and saliva were stored at -80°C until use for PCR- Denaturing Gradient Gel Electrophoresis (DGGE) in order to compare their microbial

Figure 1. Schematic description of the two experimental groups, each consisting of 11 volunteers. Toothpastes were randomly assigned and included: - Toothpaste without antibacterial claims (Prodent Softmint, Sara Lee Household & Bodycare, Exton, USA).- Stannous fluoride containing toothpaste (Oral-B Pro Expert, Procter & Gamble, Cincinnati, USA).- Triclosan containg toothpaste (Colgate Total, Colgate-Palmolive Company, Piscataway, USA).The mouthrinse that was used is Cool Mint Listerine® (Pfizer Consumer Healthcare, Morris Plains, NJ, USA)

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5415D, Hamburg, Germany) and subsequently washed and vortexed with 200 μL TE-buffer (10 mM Tris-HCl, 1 mM EDTA pH 7.4), again followed by centrifugation for 5 min at 13,000 g. Next, the supernatant was removed and the pellet was placed in a microwave (500 W, 5 min), after which it was suspended in 50 μL TE-buffer, vortexed and placed on ice. The quality and quantity of DNA samples were measured with a NanoDrop® spectrophotometer (ND-1000, NanoDrop Technologies, Inc, Wilmington, DE, USA) at 230 nm. The final concentration of each DNA sample was adjusted to 100 ng DNA for PCR amplifications.

PCR was performed with a Tgradient thermocycler (Bio-rad I-cycler, GENOtronics BV, USA). For amplification of the 16S rRNA gene, the following bacterial primers were used: F357-GC (forward primer, 5’-GC clamp-TACGGGAGGCAGCAG-3’)17 containing a GC clamp (5’- CGCCCGCCGCGCCCCGCGCCCGGCCCGCCGCCCCCGCCCC-3’)18 to make it suitable for DGGE, and R-518 (reverse primer, 5’-ATTACCGCGGCTGCTGG- 3’).19 Twenty-five μL of each PCR mixture contained 12.5 μL PCR Master Mix (0.05 units/μL Taq DNA polymerase in reaction buffer, 4 mM MgCl2, 0.4 mM dATP, 0.4 mM dCTP, 0.4 mM dGTP, 0.6 mM dTTP (Fermentas Life Sciences)), 1 μL of both forward and reverse primer (1 μM), and 100 ng DNA (in a volume of 10.5 μL). The temperature profile included an additional denaturing step of 5 min at 94°C, followed by a denaturing step at 94°C for 45 s, a primer annealing step at 58°C for 45 s, an extension step at 72°C for 1 min and a final extension step of 72°C for 5 min. PCR products were analyzed by electrophoresis on a 2.0% agarose gel containing 0.5 μg/mL ethidium bromide.

DGGE of PCR products generated with the F357-GC/R-518 primer set was performed, as described by Muyzer et al.,20 using system PhorU (INGENY, Goes, The Netherlands). The PCR products were applied on 8% (w/v) polyacrylamide gel in 0.5 X TAE buffer (20 mM Tris acetate, 10 mM sodium acetate, 0.5 mM EDTA, pH 8.3). The denaturing gradient consisted of 30 to 80% denaturant (100% denaturant equals 7 M urea and 37% formamide). Gels were poured using a gradient mixer. A 10 mL stacking gel without denaturant was added on top. Electrophoresis was performed overnight at 120 V and 60°C. Gels were stained with silver nitrate.18 Each DGGE gel was normalized according to a marker consisting of 7 reference species comprising common bacterial species associated with oral health and disease21 and stored at 4°C. The reference strains included Lactobacillus sp., Streptococcus oralis ATCC 35037, Streptococcus mitis ATCC 9811, Streptococcus sanguinis ATCC 10556, Streptococcus salivarius HB, Streptococcus sobrinus ATCC 33478 and S. mutans ATCC 10449.14

Scanning electron microscopyBiofilms on the different wires were visualized using scanning electron microscopy (SEM). Wires were fixed overnight in 2% glutaraldehyde and post-fixed for 1 h with 1%

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osmiumtetroxide. After dehydration through a water-ethanol series, wires were incubated in tetramethylsilane, air–dried and sputter-coated with a gold-palladium alloy, after which they were fixed on SEM-stub-holders using double-sided sticky carbon tape and visualized in a field emission scanning electron microscope (FE-SEM), type 6301F (JEOL Ltd., Tokyo, Japan) at 2 kV with a working distance of 39 mm and a small spot size.

Statistical analysisData were analyzed with the Statistical Package for Social Sciences (Version 16.0, SPSS Inc., Chicago, IL, USA). A one-way analysis of variance (ANOVA) was used to compare the number of bacteria and their percentage viability. A Bonferroni test was used for post-hoc multiple comparisons. Statistical significance was set at p < 0.05.

DGGE gel images were converted and transferred into a microbial database with GelCompar II, version 6.1 (Applied Maths N.V, Sint-Martens-Latem, Belgium). Similarities in bacterial composition of the different biofilms and salivary samples were analysed using a band based similarity coefficient and a non-weighted pair group method with arithmetic averages was used to generate dendograms indicating similarities in composition.22and a non-weighted pair group method with arithmetic averages was used to generate dendograms indicating similarities in composition.22

RESULTS

The total number of bacteria collected from the multi-strand wire was slightly but significantly higher than from single-strand, regardless of the oral health care regimen applied (p < 0.01, Table 1). The percentage viability of the bacteria adhering to the different types of wires was significantly higher on single-strand wires compared to multi-strand wires (p < 0.05) and buccal enamel surfaces (p < 0.001) when using a standard, fluoridated toothpaste without antibacterial claims, regardless of the additional use of an essential-oils containing mouthrinse.

The use of antibacterial toothpastes not complemented with the mouthrinse, hardly affected the total number of bacteria retrieved from the wires but their viability was significantly reduced (p < 0.001). The viability on the wires remained higher than on buccal enamel surfaces. The combined use of a triclosan containing toothpaste with the mouthrinse yielded the lowest number and viability of adhering bacteria on either wire.

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Microbial composition of biofilms adhering to the different wires and buccal enamel surfaces and of the salivary microbiome are compared in a cluster tree (Figure 2A), combining the different oral hygiene regimens. The composition of the salivary microbiome separates from the composition of the different adhering biofilms, mainly through a higher prevalence of S. salivarius and a lower prevalence of S. mutans in saliva (Table 2). Biofilms adhering on the wires have a higher prevalence of Lactobacilli and S. sobrinus than biofilms adhering on buccal enamel surfaces (Table 2).

When combining results for the different biofilms, an influence of the oral health care regimens becomes evident (Figure 2B). Regimens involving only the triclosan containing toothpaste, and the different individual toothpastes combined with the mouthrinse, form clear clusters. The regimen involving only the stannous fluoride toothpaste, yields a decrease in prevalence of Lactobacilli, S. oralis/S. mitis and S. sanguinis in comparison with the toothpaste without antibacterial claims and this decrease continues when the stannous fluoride containing paste is used together with the mouthrinse. In the latter, combined regime, also S. salivarius is found less prevalent (Table 2). The prevalence of S. sobrinus and S. mutans in biofilms adhering to wires and buccal enamel surfaces is similar as for the paste without antibacterial

Number of bacteria (Log-units)

% live bacteria

Single strand

Multi strand Single strand

Multi strand Enamel

Toothpaste without antibacterial claims

7.5 ± 0.2a 8.0 ± 0.2 68 ± 10a,d 51 ± 19 38 ± 14

Toothpaste without antibacterial claims + mouthrinse

7.5 ± 0.2a 70.9 ± 14.5 7.2 ± 0.4a 75.0 ± 6.7 46 ± 11

Stannous fluoride containing toothpaste

7.3 ± 0.1a 7.8± 0.3 25 ± 8e 36 ± 10e 20 ± 12e


7.0 ± 0.1a,b 7.5 ± 0.3b 24 ± 10e 32 ± 11e 22 ± 10e

Triclosan containing toothpaste

7.1 ± 0.2a,b 7.7 ± 0.3b 27 ± 8e 30 = 4e 17 ± 8e

Tricosan containing toothpaste + mouthrinse

6.6 ± 0.2a,b,c 7.4 ± 0.2b 23 ± 7e 28 = 4e 19 ± 4e

a Significantly different from multi-strand wireb Significantly different from a toothpaste without antibacterial claimsc Significantly different from toothpaste onlyd Significantly different from enamel e Significantly different from a toothpaste without antibacterial claims, with or without the use of mouthrinse

Table 1. The number of bacteria retrieved from 1 cm retainer wires treated with the different toothpastes and with or without the essential-oils containing mouthrinse and their viability. For reference, the viabilities on buccal enamel surfaces is also provided, but because of experimental reasons no comparative data could be given on the total numbers of adhering bacteria. All data represent averages ± standard deviations over 11 different volunteers.

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claims, regardless of whether complemented with the use of the mouthrinse. The triclosan containing toothpaste yields major increases in the prevalence of adhering S. oralis/S. mitis, S. sanguinis and S. mutans. However, when combining the triclosan containing toothpaste with the essential-oils containing mouthrinse, we see the lowest prevalences of Lactobacilli, S. sobrinus and S. mutans developing over the different regimens applied.

Scanning electron micrographs (Figure 3) show the protected location of bacteria adhering to multi-strand wires. On the multi-strand wires, bacteria are mostly located in the crevices between strands, while on the single-strand wires bacteria are present as a thin scattered film, attached mainly to irregularities on the wire surface.

STRAINS COMBINING ORAL HEALTH CARE REGIMENS

Single-strand wire Multi-strand wire Enamel Saliva

Lactobacillus 20 25 13 22S. oralis/mitis 55 56 57 65S. sanguinis 63 60 65 57S. salivarius 16 21 20 57S. sobrinus 45 52 39 46S. mutans 57 48 57 35

STRAINS COMBINING BIOFILMS ADHERING TO WIRES AND BUCCAL ENAMEL SURFACES

Toothpaste without antibacterial claims

Toothpaste without antibacterial claims + mouthrinse


Stannous fluoride containing toothpaste + mouthrinse


Triclosan containing toothpaste + mouthrinse

Lactobacillus 30 45 21 5 11 5S. oralis/mitis 53 95 29 20 86 71S. sanguinis 67 45 50 10 82 95S. salivarius 23 35 31 10 32 38S. sobrinus 39 80 43 70 34 33S. mutans 30 70 43 85 68 5

Table 2. Prevalence of marker strains in microbial samples from biofilms adhering to the different wires and buccal enamel surfaces and taken from the salivary microbiome for different oral health care regimens and different volunteers.

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Figure 2. Clustering trees describing the bacterial compositions of the microbial samples taken from the different volunteers included in this study. The corresponding circles in Figures 2A and 2B represent the same sample. (A) Colours indicate different locations of microbial sampling, i.e. enamel, retention wires or saliva. (B) Colours indicate the use of different oral health care regimens.

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Figure 3. Scanning electron micrographs of 1 week old biofilms formed in vivo, during use of a toothpaste without antibacterial claims. (A), (B), (C) Single-strand wire. (D), (E), (F) Multi-strand wire.

DISCUSSION

Biofilm formation in vivo on both single-strand and multi-strand retention wires during the use of antibacterial toothpastes and a mouthrinse was evaluated. Although statistically significant differences were found in the numbers of bacteria adhering to retention wires upon use of different toothpastes with and without antibacterial claims, and when complemented or not with the use of an essential-oils containing mouthrinse, these differences are likely too small to be of clinical significance. This coincides with results of clinical studies, showing that antibacterial toothpastes, including the two included in this study, yield reduced amounts of oral biofilm formed.23, 24 Clinical studies also confirm a small, if any effect of the additional use of an essential-oils containing mouthrinse on oral biofilm formation.12, 25, 26 More interestingly from a clinical perspective, the use of antibacterial toothpastes reduced the percentage viability of the adhering organisms. Statistically significant, but likely clinically irrelevant differences in the number of bacteria adhering to single- and multi-strand wires were found too, but more importantly antibacterial regimens caused a stronger drop in the viability of the adhering organisms on single- than on multi-strand wires, indicating better penetration of antimicrobials in biofilms forming on single-strand wires. This coincides with a higher viability of biofilms forming on single-strand wires compared to multi-strand ones during use of a

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toothpaste without antibacterial claims. This is probably caused by the fact that, similar to antimicrobials, also nutrients have better access to bacteria adhering on single-strand than on multi-strand wires.27

Biofilms on both types of retention wires have roughly the same microbial composition (Table 2), with some differences with respect to the composition of oral biofilm on enamel surfaces. Adhering biofilms have a very different composition than the salivary microbiome. These substratum-dependent microbial compositions confirm recent work28 that the surface dictates the composition of the biofilm it attracts through differential adhesion forces exerted on different strains of bacteria. The largest differences in microbial composition in biofilms adhering to retention wires and enamel surfaces are seen after regimens of antibacterial toothpastes combined with the essential-oils containing rinse. Most strikingly and of clinical importance, a regimen comprising the use of a triclosan containing toothpaste complemented with an essential-oils containing mouthrinse yielded a reduction in the prevalence of S. mutans from 30% to 5%. Other combination regimens, increased the prevalence of S. mutans in retainer biofilms. It is intriguing why the combination of a triclosan containing toothpaste with an essential-oils containing mouthrinse causes such a drastic shift in the composition of the oral microbiome into a direction that could be perceived as being less cariogenic, i.e. comprising less S. mutans. Oil containing mouthrinses have the ability to remove bacteria from the oral cavity through adhesion to the hydrophobic oil, which requires a certain degree of hydrophobicity of the bacterial cell surface.29 Moreover, certain concentrations of cationic antibacterial agents such as cetylpyridinium chloride and chlorhexidine have been demonstrated to promote binding of oral microorganisms to oil droplets.30

Hypothetically, exposure to the non-polar triclosan31 could make S. mutans cell surfaces more hydrophobic which would facilitate their removal by hydrophobic oils. In order to verify this hypothesis, we exposed a S. mutans strain used in this study to supernatants of the different toothpastes, and examined its removal by a hexadecane in the so-called kinetic MATH assay,32 as described in the Supplementary information. S. mutans possessed a low removal rate by hexadecane (Figure S1), classifying its surface as very little hydrophilic (Table S1, Supplementary information). Only exposure to the triclosan containing toothpaste supernatant however, increased the removal by hexadecane of S. mutans (see also Figure S1 and Table S1). This finding supports our hypothesis that exposure to triclosan can make S. mutans cell surfaces more hydrophobic facilitating their removal by oil containing mouthrinses and corresponds with the observation that S. mutans strains grown in the presence of triclosan formed more extensive biofilms.33 However, the authors of the latter paper ruled out effects of triclosan on streptococcal cell surface hydrophobicity, probably because they did not use the MATH assay in its more sensitive kinetic mode as advocated by Ligtenberg et al.32 Pathways to influence the composition of oral biofilms towards a “healthy” composition are

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still in its infancy. Adsorption of toothpaste components to create more hydrophobic surfaces of selected oral pathogens and making use of hydrophobic oil-containing mouthrinses for their subsequent removal from the oral cavity seems like a clinically feasible approach to this end. The present results warrant more research into components that alter the cell surface hydrophobicity of selected oral bacterial strains.

At this stage it is impossible to say whether the compositional changes observed have any beneficial clinical effect. However, it has been shown that the use of an antibacterial toothpaste containing sodium lauryl sulphate and stannous fluoride or triclosan, increases the pH of oral biofilm and decreases its viability34, 35 to yield a less cariogenic biofilm. Clearly such changes in biofilm properties may be taken as an indication of an altered microbial composition if not of a reduced prevalence of S. mutans and Lactobacilli in the biofilm. Most studies on oral biofilm composition, including the present one, make use of a control regimen, like in our study the use of a NaF-sodium lauryl sulphate containing toothpaste with mint flavour. This paste was chosen as a control, because it has no antimicrobial claims, but at the same time it cannot be ruled out that it affects both oral biofilm viability as well as composition. Both fluoride, sodium lauryl sulphate as well as mint flavouring agents are known to have antibacterial properties, 15, 36, 37 while fluoride is known to inhibit calcium-bridging between co-adhering pairs of oral bacteria.38

In summary, oral biofilm formation in vivo is slightly less on single-strand retention wires than on multi-strand wires. Orthodontic patients with a fixed bonded retainer benefit from use of an appropriate regimen of an antibacterial toothpaste and mouthrinse, not so much through reduction of the amount of biofilm formed, but rather through reduction of its viability. Moreover, appropriate regimens may make the selected members of the oral microbiome more hydrophobic through adsorption of non-polar components from toothpastes to subsequently enhance their removal by oil containing mouthrinses, yielding less pathogenic biofilms. This pathway to restoring a healthy oral microbiome needs to be explored further though.

ACKNOWLEDGEMENTS

This study was entirely funded by UMCG, Groningen, and The Netherlands. The authors would like to extend their gratitude to the volunteers who participated in this study, Mr. Jeroen Kuipers from the Centre for Medical Electron Microscopy of the University Medical Centre Groningen for his assistance with the SEM analysis, DENTSPLY Lomberg (Soest, The Netherlands) and Orthotec B.V. (Zeist, The Netherlands) for kindly providing the different wires used in this study. HJB is also director of a consulting company SASA BV, Thesinge, The Netherlands.

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CONFLICT OF INTERESTS

The authors declare no potential conflicts of interest with respect to authorship and/or publication of this article. Opinions and assertions contained herein are those of the authors and are not construed as necessarily representing views of the companies who donated the different wires or their respective employers.

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GA (2007) The long-term effect of a mouthrinse containing essential oils on dental plaque and gingivitis: a systematic review. J Periodontol 78:1218-1228 13. Pizzo G, La Cara M, Licata ME, Pizzo I, D’Angelo M (2008) The effects of an essential oil and an amine fluoride/stannous fluoride mouthrinse on supragingival plaque regrowth. J Periodontol 79:1177-1183 14. Otten MP, Busscher HJ, Abbas F, Van der Mei HC, Van Hoogmoed CG (2012) Plaque-left-behind after brushing: intra-oral reservoir for antibacterial toothpaste ingredients. Clin Oral Investig 16:1435-1442 15. Addy M, Moran J (2008) Chemical Supragingival Plaque Control. In: Lang NP, Lindhe J (eds) Clinical periodontology and implant dentistry, Vol 2 edn. Blackwell Munksgaard, pp 734-765 16. Syed SA, Loesche WJ (1972) Survival of human dental plaque flora in various transport media. Appl Microbiol 24:638-644 17. Di Cagno R, Rizzello CG, Gagliardi F, Ricciuti P, Ndagijimana M, Francavilla R, Guerzoni ME, Crecchio C, Gobbetti M, De Angelis M (June 15, 2009) Different Fecal Microbiotas and Volatile Organic Compounds in Treated and Untreated Children with Celiac Disease. Appl Environ Microb 75:3963-3971 18. Zijnge V, Welling GW, Degener JE, Van Winkelhoff AJ, Abbas F, Harmsen HJ (2006) Denaturing gradient gel electrophoresis as a diagnostic tool in periodontal microbiology. J Clin Microbiol 44:3628-3633 19. Marsh PD (1994) Microbial ecology of dental plaque and its significance in health and disease. Adv Dent Res 8:263-271 20. Muyzer G, De Waal EC, Uitterlinden AG (1993) Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microb 59:695-700 21. Marsh PD (2006) Dental plaque as a biofilm and a microbial community - implications for health and disease. BMC Oral Health 6 Suppl 1:S14 22. Signoretto C, Bianchi F, Burlacchini G, Sivieri F, Spratt D, Canepari P (2010) Drinking habits are associated with changes in the dental plaque microbial community. J Clin Microbiol 48:347-356 23. Riley P, Lamont T (2013) Triclosan/

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SUPPLEMENTARY INFORMATION: MICROBIAL ADHESION TO

HYDROCARBONS (MATH)

S. mutans ATCC 10449 grown on blood agar plates from a frozen stock, was used to inoculate 10 mL Tryptone Soya Broth (TSB) and cultured for 24 h at 37ºC. This culture was used to inoculate 100 mL TSB, which was grown overnight. Bacteria were harvested by centrifugation and washed twice with potassium phosphate buffer (pH 7.0) and suspended to an optical density A0 (at 600 nm) of between 0.4 and 0.6. Next, half of the suspension was mixed with the supernatant of a toothpaste slurry (25% by weight) in water used after centrifugation, 5 min at 10,000 g to remove particulate matter for 2 min, centrifuged, washed and resuspended in potassium phosphate buffer to an optical density A0 (at 600 nm) of between 0.4 and 0.6. In order to measure the hydrophobicity of the bacterial cell surfaces before and after exposure to a toothpaste supernatant, 150 µL hexadecane was added to 3 mL of each suspension and the suspension was vortexed for 10 s, allowed to settle for 10 min for phase separation and finally the optical density At of the aqueous phase was measured. This was repeated 6 times and log (At/A0 x100) was plotted against the vortexing time (Figure S1). Initial removal rates R0 (min-1) were calculated as the slopes of the tangent of the curves obtained and used to compare effects of adsorption of toothpaste components on the hydrophobicity of the streptococcal cell surface (Table S1).

Supplementary Table S1. Cell surface hydrophobicity of S. mutans ATCC 10449 before and after exposure to slurries of the different toothpastes involved in this study, as measured by the kinetic MATH assay and expressed in terms of their initial removal rates. All data represent the average ± SD of three experiments with

Toothpaste used Initial removal rate (min-1)

None 0.01 ± 0.01Toothpaste without antibacterial claims 0.0 ± 0.0Stannous fluoride containing toothpaste 0.0 ± 0.0Triclosan containing toothpaste 0.05 ± 0.011a

a Significantly different from all other data at p<0,000 (A One-Way ANOVA was used with a Bonferroni test for post-hoc multiple comparisons. Statistical significance was set at p< 0.05.

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Figure S1. Optical density log (At/A0 x100) as a function of the vortexing time for the removal of S. mutans ATCC 10449 prior to or after its exposure to a toothpaste slurry by hexadecane. Each data point represents the average over three experiments with different bacterial cultures. Standard deviations are smaller than the data points.

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Chapter 5Stress relaxation analysis facilitates a quantitative

approach towards antimicrobial penetration into biofilms

Yan He, Brandon W. Peterson, Marije A. Jongsma, Yijin Ren, Prashant K. Sharma, Henk J. Busscher, and Henny C. van der Mei.

PLos One (2013) 8:e63750

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Visco-elasticity of biofilms and chlorhexidine Chapter 5

ABSTRACT

Biofilm-related infections can develop everywhere in the human body and are rarely cleared by the host immune system. Moreover, biofilms are often tolerant to antimicrobials, due to a combination of inherent properties of bacteria in their adhering, biofilm mode of growth and poor physical penetration of antimicrobials through biofilms. Current understanding of biofilm recalcitrance toward antimicrobial penetration is based on qualitative descriptions of biofilms. Here we hypothesize that stress relaxation of biofilms will relate with antimicrobial penetration. Stress relaxation analysis of single-species oral biofilms grown in vitro identified a fast, intermediate and slow response to an induced deformation, corresponding with outflow of water and extracellular polymeric substances, and bacterial re-arrangement, respectively. Penetration of chlorhexidine into these biofilms increased with increasing relative importance of the slow and decreasing importance of the fast relaxation element. Involvement of slow relaxation elements suggests that biofilm structures allowing extensive bacterial re-arrangement after deformation are more open, allowing better antimicrobial penetration. Involvement of fast relaxation elements suggests that water dilutes the antimicrobial upon penetration to an ineffective concentration in deeper layers of the biofilm. Next, we collected biofilms formed in intra-oral collection devices bonded to the buccal surfaces of the maxillary first molars of human volunteers. Ex situ chlorhexidine penetration into two weeks old in vivo formed biofilms followed a similar dependence on the importance of the fast and slow relaxation elements as observed for in vitro formed biofilms. This study demonstrates that biofilm properties can be derived that quantitatively explain antimicrobial penetration into a biofilm.

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INTRODUCTION

In the 17th century the Dutch fabric merchant Antonie van Leeuwenhoek started to construct his own microscopes in order to be able to better examine the quality of the fabrics he bought and sold. He examined more than just his fabrics and after utilizing one of his own microscopes in 1684 to look at the accumulation of matter on his teeth, he remarked in a report to the Royal Society of London: “The number of these animalcules in the scurf of a man’s teeth are so many that I believe they exceed the number of men in a kingdom”. This was not enough however, to satisfy the curiosity of the fabric merchant, who would become one of the most famous microbiologists of all times, and he furthermore discovered “that the vinegar with which I washt my Teeth, kill’d only those Animals which were on the outside of the scurf, but did not pass thro the whole substance of it”.

Translated to one of the important topics in modern microbiology, Van Leeuwenhoek was referring to the biofilm mode of growth of bacteria adhering on a surface,1 embedding themselves in a matrix of extracellular polymeric substances (EPS)2 that not only offers physical protection against antimicrobial penetration but can also yield bacterial properties that are different from their planktonic counterparts. Bacteria in their adhering, biofilm mode of growth can become inherently resistant to antimicrobials through mutation,3 formation of antibiotic degrading enzymes,4 endogenous oxidative stress,5 phenotypic changes,6 and low metabolic activities.7 Despite extensive studies over many centuries, prevention of biofilm formation remains a prime challenge in many industrial and biomedical applications. In industrial applications, biofilms inflict major damage when formed on processing equipment or in pipes used to transport resources.8 In the biomedical field, biofilm-related infections can develop everywhere in the human body from head (oral biofilms9) to toe (infected diabetic foot ulcers10). Biofilm-related infections are rarely cleared by the host immune system and especially infections that arise after implantation of biomaterial implants (e.g. prosthetic hips and knees) or devices (e.g. pace makers) are known to be persistent and difficult to treat, since the antimicrobial tolerance of bacteria in their biofilm mode of growth extends to many antibiotics used in modern medicine.11 Moreover, dental caries and periodontal diseases, the most wide-spread infectious diseases in the world, are due to biofilms that Van Leeuwenhoek tried to eliminate by using vinegar as an antimicrobial mouthrinse.12

Although the microscopes used nowadays are more sophisticated than the ones Van Leeuwenhoek employed, our understanding of the recalcitrance of biofilms toward antimicrobial penetration is still based on qualitative description of biofilms,13 using expressions as “water channels”, “mushroom structures”, “whiskers” and “streamers”.14,15 This raises the question whether quantifiable properties of biofilms exist that would relate with antimicrobial penetration into a biofilm. As for polymeric materials, structural and

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compositional properties of biofilms, should be reflected in their viscoelastic properties. Viscoelastic properties of oral biofilms depend on the degree of compaction during formation, the absence or presence of flow during growth, their architecture and microbial composition.16,17 The viscoelastic properties of oral biofilms can be determined by evaluating their relaxation after deformation during external loading. Stress relaxation during external loading is a time-dependent process and can be separated into a number of responses, each with a characteristic time-constant.18 Although Maxwell analysis of stress-relaxation to derive the characteristic time-constants of the various relaxation processes that occur in a biofilm under external loading has been done before,19 results have been regarded mainly from a mathematical perspective and the details of the relaxation-structure-composition relation in biofilms and the physical processes associated with the different time-constants, are mostly neglected. Stress relaxation may involve a number of processes, like the outflow of water and EPS from the biofilm and re-arrangement of the bacteria in the biofilm.20 Since penetration of an antimicrobial into a biofilm depends on diffusion21 and therewith on its structural and compositional features, like the presence of water-filled channels in the biofilm or EPS-containing spaces, we here hypothesize that the penetration of an antimicrobial into a biofilm may relate with stress relaxation and its underlying processes.

The aim of this study is to gain evidence in support of this hypothesis. To this end, single-species biofilms of two oral bacterial strains, Streptococcus oralis and Actinomyces naeslundii were grown in a parallel plate flow chamber (PPFC)22 and in a constant depth film fermenter (CDFF).23 Subsequently, we measured their viscoelastic properties using a low load compression tester, as well as the penetration of chlorhexidine into the biofilms. Following Van Leeuwenhoek, we chose to collect support for our hypothesis based on oral biofilms, because the human oral cavity is highly accessible and also allows for sampling of in vivo formed biofilm. Therefore, in order to not only gain in vitro evidence in support of our hypothesis, an intra-oral biofilm collection device was developed to grow oral biofilms in situ, in absence of mechanical perturbation. In vivo formed biofilms in the devices worn by human volunteers were examined ex situ with respect to their viscoelastic properties and chlorhexidine penetration and results and conclusions compared with those obtained for in vitro formed oral biofilms. Chlorhexidine is known to be the most effective oral antimicrobial to date24 and surprisingly, despite its extensive use, inherent bacterial resistance against chlorhexidine has hardly or never been reported as compared to antibiotic resistance of many bacterial pathogens. This makes chlorhexidine an ideal antimicrobial to separate a possible inherent tolerance of biofilm bacteria for the antimicrobial from the physical protection offered by the biofilm mode of growth and study its penetration through a biofilm.

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Bacterial strains and growth conditions S. oralis J22 and A. naeslundii T14V-J1 grown on blood agar plates, were used to inoculate 10 ml modified Brain Heart Infusion broth (BHI, Oxoid Ltd., Basingstoke, UK) (37.0 g/l BHI, 5.0 g/l yeast extract, 0.4 g/l NaOH, 1.0 g/l hemin, 0.04 g/l vitamin K1, 0.5 g/l L-cysteine, pH 7.3) and were cultured for 24 h at 37°C in ambient air for S. oralis J22 and anaerobically for A. naeslundii T14V-J1. These cultures were used to inoculate 200 ml modified BHI and grown for 16 h. Bacteria were harvested by centrifugation at 870 g, 10°C for 5 min and washed twice in sterile adhesion buffer (50 mM potassium chloride, 2 mM potassium phosphate, 1 mM calcium chloride, pH 6.8). The bacterial pellet was suspended in 10 ml adhesion buffer and sonicated intermittently in an ice-water bath for 3 × 10 s at 30 W (Vibra cell model 375, Sonics and Materials Inc., Newtown, CT, USA) to break bacterial chains and clusters, after which bacteria were resuspended in adhesion buffer. A concentration of 3 × 108 bacteria/ml was used for PPFC experiments, while a concentration of 9 × 108 bacteria/ml was used in CDFF experiments.

Biofilm formation in a PPFC and CDFFBiofilms were grown on glass slides (water contact angle 7 ± 3 degrees) and hydroxyapatite discs (water contact angle 34 ± 8 degrees) in a PPFC and a CDFF, respectively after adsorption of a salivary conditioning film from reconstituted human whole saliva for 14 h at 4°C under static conditions. Reconstituted human whole saliva was obtained from a stock of human whole saliva from at least 20 healthy volunteers of both genders, collected into ice-cooled beakers after stimulation by chewing Parafilm®, 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. Freeze-dried saliva was dissolved in adhesion buffer (1.5 g/l). All volunteers, gave their verbal informed consent to saliva donation according to a fixed written protocol and were registered in order to document the gender, age and health status of the volunteers, in agreement with the guidelines set out by the Medical Ethical Committee at the University Medical Center Groningen, Groningen, The Netherlands (letter 06-02-2009). Written consent was not required since saliva collection was entirely non-invasive, saliva’s were pooled prior to use and the study was not aimed towards measuring properties of the saliva. Rather saliva was used to lay down an adsorbed protein film prior to biofilm formation studies.

For biofilm formation in the PPFC, 200 ml bacterial suspension was circulated at a shear rate of 15 s-1 in a sterilized PPFC till a bacterial surface coverage of 2 × 106 cm-2 was achieved on a saliva-coated glass bottom plate (for details see16). Subsequently, adhesion buffer

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was flowed at the same shear rate of 15 s-1 for 30 min in order to remove non-adhering bacteria from the tubes and flow chamber. Next, growth medium (20% modified BHI and 80% adhesion buffer) was perfused through the system at 37°C for 48 h, also at a shear rate of 15 s-1.

Biofilms were grown in a sterile CDFF (for details see23) on saliva coated hydroxyapatite discs by introducing 200 ml bacterial suspension in the fermenter during 1 h, while the table with the sample holders was rotating at 1 rpm. Then, rotation was stopped for 30 min to allow bacteria to adhere before growth medium was introduced and rotation resumed. The biofilm was grown for 96 h at 37°C under continuous supply of a mixture of adhesion buffer and modified BHI at a rate of 80 ml/h. The system was equipped with 15 sample holders and each sample holder contained 5 saliva coated hydroxyapatite discs, recessed to a depth of 100 µm.

Oral biofilm collection in vivo The intra-oral biofilm collection device (Fig. 1) was made of medical grade stainless steel 316, and is composed of two parts: a base (5×3×2 mm) that is fixed to the center of the buccal surface of the upper first molars and a replaceable cover plate (4×3×0.2 mm). Biofilms formed on the inner side of the replaceable cover plate in the absence of mechanical perturbations, were considered for this study.

Five volunteers (aged 26 to 29 years) were included in this study. Volunteers all had a complete dentition with maximally one restoration, no bleeding upon probing and were not using any medication. Each volunteer was assigned a random number between 1 and 5 used for later data processing. The study was approved according to the guidelines of the Medical Ethics Committee of the University Medical Center Groningen, Groningen, The Netherlands (letter 28-9-2011), including the written informed consent by the volunteers and the tenets of the Declaration of Helsinki.

A base device was fixed to buccal surfaces of the upper first molars of the volunteers (see also Fig. 1) after mild etching of the tooth surface using light cure adhesive paste (Transbond™ XT, 3M Unitek, USA), a procedure similar to the one used for the bonding of orthodontic brackets. Prior to bonding, the base and cover plate of the device were brushed using a rubber cup and cleaner paste (Zircate® Prophy Paste, Densply, Caulk, USA) at low speed (less than 2,500 rpm/min) and autoclaved. Subsequently, the base surface was coated with a thin layer of primer and bonding agent (CLEARFIL SE BOND, Kurary Medical Inc., Japan). The stainless steel cover plate was inserted using a pair of tweezers and kept in place using Light Cure Adhesive Paste (Transbond™ XT, 3M Unitek, USA). Volunteers were asked to wear the device for a total of eight weeks during which they were requested

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to perform manual brushing with a standard fluoridated toothpaste (Prodent Softmint®, Sara Lee Household & Bodycare, Exton, USA) according to their habitual oral hygiene but to refrain from the use of an additional mouthrinse.

The cover plates could be removed with a dental explorer, after which cover plates with biofilm were placed in a moisturized petri dish for transport from the dental clinic to the laboratory. In a separate pilot study, it was established that two weeks of intra-oral biofilm formation in the device yielded biofilm thicknesses that were similar to the ones obtained in vitro. Therewith, in vivo biofilms could be collected four times from each volunteer. After each experiment, cover plates were sanded to remove biofilm and other residuals, prior to autoclaving. After the experiments, the base of the device was removed from the tooth surface with a debracketing plier and residual adhesive was grinded off the tooth surface with a low speed hand piece. A base device was only used once in each volunteer. The tooth surface was polished and cleaned with rubber cup and cleaner paste. No signs of gingival inflammation were observed in any volunteer after removal of the base device.

Figure 1. Intra-oral biofilm collection device.A: The stainless steel base and cover plate of the device. B: The base of the intra-oral biofilm collection device fixed to the center of the buccal surface of a maxillary first molar. C: Side view of the intra-oral biofilm collection device, showing the open spacing in which undisturbed biofilm growth to the cover plate occurred.D: Top view of the closed intra-oral biofilm collection device in situ, showing the hole in the cover plate used for its removal with a dental explorer.

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Low load compression testing The thickness and stress relaxation of the biofilms were measured with a low load compression tester, described before.16 Stress relaxation was monitored after inducing 10, 20, and 50% deformation of the biofilms within 1 s and held constant for 100 s, while monitoring the stress relaxation (see Fig. 2A). Each deformation was induced three times at different locations on the same biofilm.

Stress relaxation as a function of time was analyzed using a generalized Maxwell model containing three elements (see Fig. 2B) according to

(1)

in which E(t) is the total stress exerted by the biofilm divided by the strain imposed, expressed as the sum of three Maxwell elements with a spring constant Ei, and characteristic decay time, ti (see also Fig. 2B). For calculating E(t), deformation was expressed in terms of strain, e, according to the large strain model using

(2)

where Dh is the decrease in height and h is the un-deformed height of the biofilm. The model fitting for Ei and ti values of the three elements was done by minimizing the chi-squared value using the Solver tool in Microsoft Excel 2010. Fitting to three Maxwell elements yielded the lowest chi-squared values and increasing the number of Maxwell elements only yielded minor decreases in chi-squared values of less than 3%. The elements derived were rather arbitrarily named fast, intermediate or slow based on their t values, i.e. t1 < 5 s, 5 s < t2 < 100 s and t3 > 100 s, respectively (see also Fig. 2B). Relative importance of each element, based on the value of its spring constant Ei, was expressed as the percentage of its spring constant to the sum of all elements’ spring constants at t = 0.

Penetration of chlorhexidine into biofilmsIn vitro and in vivo formed biofilms were all exposed in vitro to a 0.2 wt% chlorhexidine-containing mouthrinse (Corsodyl®, SmithKline Beecham Consumer Brands B.V., Rijswijk, The Netherlands) for 30 s and subsequently immersed in adhesion buffer for 5 min. After exposure to chlorhexidine, biofilms were stained for 30 min with live/dead stain (BacLight™, Invitrogen, Breda, The Netherlands) and CLSM (Leica TCS-SP2, Leica Microsystems Heidelberg GmbH, Heidelberg, Germany) was used to record a stack of images of the biofilms with a 40× water objective lens. Images were analyzed with Leica confocal software to visualize live and dead bacteria in the biofilms. The ratio of the intensity of red (dead

E(t) = E1e−tτ1 +E2e

−tτ 2 +E3e

−tτ3

ε = ln(1+ Δhh)

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bacteria) to green (live bacteria), R/G, was plotted versus the biofilm thickness (see Fig. 3). The biofilm thickness where the ratio R/G became less than 1.5 was taken as the thickness of the dead band. Next, a penetration ratio was calculated according to

(3)

Penetration ratios were calculated for three different, randomly chosen locations on the biofilms and presented as averaged over the different locations.

Figure 2. Measurement and Maxwell model of the viscoelasticity of biofilms.A: Stress versus time diagram for relaxation of a compressed biofilm. B: Schematic of a three element Maxwell model: Ei represent the spring constants and τi the relaxation time constants, which are equal to hi/Ei.

Penetration ratio = dead band thicknesstotal biofilm thickness

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Statistical analysis Statistical analysis was performed with SigmaPlot software (version 11.0, systat software, Inc., California, USA). Differences in biofilm thickness and viscoelasticity were evaluated after testing for normal distribution and equal variance of the data. If data failed one of these tests, a Mann-Whitney Rank Sum test was used to determine statistical significance, otherwise a Student t-test was applied. Pearson Product Moment Correlation test was used to disclose relations between the penetration of chlorhexidine into and the relaxation of biofilms.

Figure 3. Chlorhexidine penetration into in vitro and in vivo formed oral biofilms and the calculation of the penetration ratio.I. Representative CLSM-images (cross sectional view) of the penetration of chlorhexidine (0.2 wt%) during 30 s into oral biofilms grown in vitro and in vivo (exposure to chlorhexidine was done in vitro). A: S. oralis J22 biofilm grown under flow in a PPFC. B: S. oralis J22 biofilm grown under compaction in a CDFF. C: A. naeslundii T14V-J1 biofilm grown under flow in a PPFC.D: A. naeslundii T14V-J1 biofilm grown under compaction in a CDFF.E and F: two weeks old, in vivo formed oral biofilm. Scale bar represents 75 μm. II. Red to green intensity ratio (R/G), denoting the ratio of dead to live organisms in a biofilm versus the thickness of the biofilm. a is the dead band thickness and b is the total biofilm thickness. R/G = 1.5 was taken as the cut-off for the thickness of the dead band.

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RESULTS

Biofilms of coccal-shaped S. oralis J22 and rod-shaped A. naeslundii T14V-J1 grown in the PPFC reached a thickness of 131 ± 15 μm and 109 ± 26 μm, respectively (Table 1). The biofilm thickness in the CDFF for S. oralis J22 was 119 ± 6 μm and 125 ± 9 μm for A. naeslundii T14V-J1. There were no significant differences (p > 0.05, Student t-test) in thickness between biofilms grown under flow and in the CDFF. Also differences in biofilms thickness across strains were not statistically significant (p > 0.05, Student t-test).

The penetration of chlorhexidine in biofilms grown in the PPFC was significantly different (p < 0.05, Mann-Whitney Rank Sum test) for S. oralis J22 and A. naeslundii T14V-J1, and the penetration ratio amounted 0.33 ± 0.09 and 0.56 ± 0.08, respectively (see also Table 1 and Fig. 3). On the other hand, there were no significant strain-dependent differences in penetration of chlorhexidine into biofilms grown in the CDFF, showing penetration ratios of 0.48 ± 0.04 and 0.39 ± 0.06 in biofilms of S. oralis J22 and A. naeslundii T14V-J1, respectively (p > 0.05, Mann-Whitney Rank Sum test). Interestingly, whereas biofilms offered a clear physical protection against chlorhexidine, bacteria dispersed from biofilms grown either in the PPFC or in the CDFF were highly susceptible to chlorhexidine (Fig. 4), confirming that the absence of bacterial killing in the deeper layers of the biofilms are not due to changes in inherent properties of the bacteria in their biofilm mode of growth, but solely to difficulties encountered by the antimicrobial in penetrating to the deeper layers. Note that a similar conclusion has been drawn for three days old in vivo grown oral biofilms, after dispersal and exposure to chlorhexidine.25

Total stress relaxation (Fig. 2A) of biofilms grown in the PPFC were different for both strains and S. oralis J22 biofilms showed significantly (p < 0.05, Mann-Whitney Rank Sum test) more stress relaxation than biofilms of A. naeslundii T14V-J1, especially after 10% and 20% induced deformation (Table 1). There were no significant differences (p > 0.05, Mann-Whitney Rank Sum test) in stress relaxation between biofilms of the coccal and rod-shaped organisms when grown in the CDFF. Interestingly, the penetration ratio of chlorhexidine decreased with increasing stress relaxation of the biofilms, regardless of the induced deformation (Fig. 5).

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Tabl

e 1.

The

thic

knes

s, p

enet

ratio

n ra

tio, t

otal

str

ess

rela

xatio

n an

d th

e re

lativ

e im

port

ance

of t

he th

ree

Max

wel

l ele

men

ts o

f in

vivo

an

d in

vitr

o fo

rmed

ora

l bio

film

s.#

Thic

knes

s (μ

m)

Pene

tratio

n ra

tio10

% d

efor

mat

ion

(%)

20%

def

orm

atio

n (%

)50

% d

efor

mat

ion

(%)

Rela

xatio

nE 1

E 2E 3

Rela

xatio

nE 1

E 2E 3

Rela

xatio

nE 1

E 2E 3

In v

ivo

121±

86a

0.20

±0.1

*60

±14*

21±1

6*27

±13

52±1

6*58

±15*

24±1

4*15

±13

61±1

7*65

±11

43±1

6a14

±943

±16

In v

itro

aver

age

120±

520.

46±0

.182

±14

44±2

028

±15

28±1

979

±15

54±2

218

±10

28±1

672

±19

54±2

613

±634

±24

PPFC

av

erag

e10

9-13

1b0.

33b -

0.56

64-9

7b17

-60b

35b -

404b -

4357

-92b

25-7

3b11

b -26

16b -

4943

-76b

18-6

5b11

b -15

24b -

68

CDFF

av

erag

e11

9b -12

50.

39-0

.48b

83b -

8347

b -49

10b -

2526

-43b

80b -

8456

-60b

10b -

2717

-30b

74b -

9059

b -75

10b -

1411

-31b

# In v

ivo

data

refe

r to

aver

ages

± S

D o

btai

ned

in fi

ve v

olun

teer

s, w

hile

in v

itro

data

are

ave

rage

s ov

er a

ll sin

gle-

spec

ies

biofi

lms

form

ed in

the

PPFC

and

CD

FF

by c

occa

l and

rod-

shap

ed o

rgan

ism

s. In

add

ition

, in v

itro

data

are

ave

rage

d as

form

ed in

the

PPFC

and

CD

FF b

y co

ccal

and

rod-

shap

ed o

rgan

ism

s.* i

ndic

ates

p <

0.0

5.

a in

dica

tes

the

com

paris

on w

as c

arrie

d ou

t by

Man

n-W

hitn

ey R

ank

Sum

test

.

b in

dica

tes

data

for S

. ora

lis J

22.

85

5


Figure 4. Tolerance and intolerance of biofilm organisms to chlorhexidine prior to and after their dispersal.Fluorescence images of dispersed S. oralis J22 and A. naeslundii T14V-J1, treated with chlorhexidine for 30 s in their biofilm mode of growth prior to dispersal and treated immediately after dispersal. Live (green)–dead (red) staining was used to show the viability of bacteria.A: S. oralis J22 grown in the PPFC and treated in its biofilm mode of growth. B: S. oralis J22 grown in the PPFC and treated in its dispersed state. C: A. naeslundii T14V-J1 grown in the CDFF and treated in its biofilm mode of growth..D: A. naeslundii T14V-J1 grown in the CDFF and treated in its dispersed state.Scale bar represents 10 μm.

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Total stress relaxation was subsequently resolved in a fast, intermediate and slow component (Fig. 2B). Since bacteria in a biofilm constitute the heaviest masses, their re-arrangement upon an induced deformation will be slow, and we associate the relative importance of the slow Maxwell element with bacterial re-arrangement in a biofilm. On the other hand, water has the smallest viscosity in a biofilm, and therefore the fast Maxwell element is associated with the flow of water through a biofilm, which leaves an association between the behavior of EPS with the intermediate Maxwell element. Analysis of the stress relaxation according to a three element Maxwell model revealed that penetration increased with increasing relative importance of the slow relaxation component and decreasing importance of the fast component (Fig. 6). This confirms the existence of a relaxation-structure-composition relation that may facilitate a quantitative approach towards antimicrobial penetration in biofilms.

In order to confirm that a relaxation-structure-composition relation facilitates understanding of antimicrobial penetration also for in vivo grown biofilms, we first developed an intra-oral biofilm collection device (Fig. 1). The average thickness of the oral biofilms formed in vivo over a time period of two weeks was 121 ± 86 μm, comparable to the thickness of in vitro biofilms (p > 0.05, Mann-Whitney Rank Sum test), as can be seen in Table 1.

Figure 5. Penetration of chlorhexidine and stress relaxation of differently grown biofilms in vitro. (A) The schematics of parallel plate flow chamber and constant depth film fermenter. (B) Penetration ratio of chlorhexidine as a function of relaxation of different biofilms for 10%, 20% and 50% induced deformation. Dashed lines indicate 95% confidence intervals.

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Total stress relaxation of in vivo biofilms upon 10% and 20% deformation were more comparable to the stress relaxation observed for in vitro biofilms grown in the PPFC than in the CDFF, as averaged over both bacterial strains (Table 1). On the other hand, upon inducing a deformation of 50%, stress relaxation of in vivo biofilms became more comparable to the one of in vitro biofilms grown in the CDFF. On average, in vitro biofilms showed higher total stress relaxation than in vivo formed biofilms, although this difference was only significant (p < 0.05, Student t-test) for 10% and 20% induced deformations (Fig. 7).

In vivo formed biofilms furthermore distinguished themselves significantly from in vitro averages by a smaller importance of the fast component (E1) and larger importance of the slow component (E3) (p < 0.05, Student t-test; Table 1) for induced deformations of 10% and 20%. At 50% induced deformation however, differences in the importance of the different relaxation parameters had disappeared (see also Fig. 7). The importance of the intermediate component (E2) was relatively similar across the different biofilms (Table 1).

Figure 6. Chlorhexidine penetration and Maxwell analyses of in vitro grown biofilms.Penetration ratio as a function of the relative importance of the three Maxwell elements E1, E2 and E3, denoting the fast, intermediate and slow relaxation components, respectively for different biofilms after 10%, 20% and 50% induced deformation. All data points refer to single experiments, while symbols are explained in Fig. 5. Dashed lines represent 95% confidence intervals.

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The chlorhexidine penetration ratio for in vivo formed biofilms was smaller than the average penetration into in vitro biofilms (p < 0.05, Student t-test; Table 1). Similarly as observed for in vitro biofilms, penetration decreased with increasing importance of the fast (E1) component

Figure 7. Stress relaxation properties of intra-orally grown oral biofilms.Relaxation properties of oral biofilms formed in vivo, obtained in five volunteers as indicated by different colors in comparison with the average relaxation properties of different single-species biofilms formed in a PPFC and CDFF, falling within the black rectangles.

Figure 8. Chlorhexidine penetration and Maxwell analyses of intra-orally grown biofilms.Penetration ratio of chlorhexidine as a function of the relative importance of the fast, intermediate and slow Maxwell elements E1, E2 and E3 for in vivo biofilms formed in different volunteers after 10%, 20% and 50% induced deformation. All data points refer to single experiments in one volunteer. Different volunteers are indicated by the same color codes as used in Fig. 7. Dashed lines represent 95% confidence intervals.

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and increased with the importance of the slow component (E3) (Fig. 8). No relation was observed with the importance of the intermediate component (E2), as was also lacking for in vitro biofilms.

DISCUSSION

The recalcitrance of oral biofilm toward penetration of antimicrobials is known ever since Van Leeuwenhoek wrote in the 17th century that “the vinegar with which I washed my teeth killed only those animals which were on the outside of the scurf, but did not pass through the whole substance of it”. Over recent years, the limited penetration of antimicrobials into a biofilm has been attributed to reduced solute diffusion in water, the presence of bacterial cells, EPS, abiotic particles or gas bubbles trapped in a biofilm.21 Interestingly, whereas the influence of the chemistry and biology of biofilms on diffusion have been amply described and reviewed,21,26,27 antimicrobial penetration has never been related with quantifiable, physical properties of a biofilm. This study demonstrates for the first time since Van Leeuwenhoek his observation of the poor penetration of vinegar into an oral biofilm, that through a relaxation-structure-composition relation, biofilm properties can be derived that facilitate explanation of antimicrobial penetration into a biofilm on basis of quantitative biofilm properties. Incidentally, not only antimicrobials have difficulty penetrating a biofilm, but also nutrients may have difficulty penetrating a biofilm, causing reduced viability of organisms residing in deeper layers of biofilms.28

The bacteria in a biofilm constitute the heaviest masses, and their re-arrangement during stress relaxation upon an induced deformation will thus be slow, which associates the relative importance of the slow Maxwell element with bacterial re-arrangement. Furthermore, the positive correlation between penetration and the importance of the slow Maxwell element confirms that organisms arranged in a more open, water-filled structure, allow easier penetration of antimicrobials. Different from the role of water-filled channels in diffusion,21 we found that water itself had a negative influence on the efficacy of antimicrobials during penetration. Since water has the smallest viscosity in a biofilm, the fast Maxwell element may be associated with the outflow of water through and its presence in biofilms. Consequently, dilution of antimicrobials after penetration into a biofilm to an ineffective concentration in deeper layers is evidenced by the negative correlation between the relative importance of the fastest Maxwell element and the penetration ratio. At this point, it must be emphasized that in our study chlorhexidine might have penetrated beyond the dead bands, as visible in Fig. 3, but clearly to a concentration insufficient to yield bacterial killing. Arguably, this raises the issue that penetration not only depends on possible physical difficulties of an antimicrobial in penetrating a biofilm, but moreover on the time allowed for penetration and antimicrobial concentration. In many clinical situations however, time and concentration cannot be increased at will. In the oral case highlighted here, the time most people allow themselves for an antimicrobial mouthrinse to be active in the oral cavity is

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30 s utmost, while concentrations of chlorhexidine higher than 0.12 w% rapidly cause severe soft tissue damage and discolorations of teeth.29 Equilibration of a biofilm with an antimicrobial as can be achieved in vitro is thus often impossible for the in vivo situation. Clearly, similar types of limitations with respect to time and/or concentration exist everywhere in the human body where antimicrobials are applied to combat biofilm-related infections, emphasizing the importance of good penetration in biofilm control through the use of antimicrobials.

The importance of a relaxation-structure-composition relation for biofilms and its role in understanding antimicrobial penetration was established both for in vivo grown biofilms as well as in two distinctly different model systems to grow biofilms in vitro. In the CDFF, there is a constant turn-over of bacterial growth, death and biofilm removal by the scraper blades23 in addition to compaction by the blades. Whereas similar turn-over, death and removal by fluid flow can be expected in a PPFC, compaction is absent in a PPFC. In this respect, it is interesting that there was no difference in stress relaxation of biofilms formed by coccal or rod-shaped organisms in the CDFF, presumably because biofilms in the CDFF are mechanically compacted during formation (see Table 1). In the absence of mechanical compaction like in the PPFC, rod-shaped organisms have more difficulties in spontaneously forming a dense structure, as this requires organisms to take a favorable orientation with respect to one another. This becomes especially evident at the larger deformation induced of 50% and explains why biofilms formed by rod-shaped organisms in the PPFC had a different stress relaxation than coccal organisms, but not in the CDFF.

The two model systems to grow biofilms used in this study represent two extreme situations that may occur in the oral cavity. Highly compacted biofilms may be expected in fissures due to mastication, while compaction occurs less on interproximal biofilms. In addition, biofilm-left-behind in interproximal spaces inaccessible to contact-brushing will be in a more “fluffed-up” state,30 resembling biofilms grown in a PPFC. Indeed, biofilms grown in our intra-oral biofilm collection device, inaccessible to contact toothbrushing, are more fluffed up than in in vitro formed biofilms (compare Figs. 3E and F with Figs. 3A-D). Accordingly, stress relaxation characteristics after 10% and 20% deformation of biofilms formed in the PPFC more closely resemble those of in vivo formed biofilms than biofilms formed in the CDFF. This is especially so for 10% and 20% induced deformations, yielding information on the relaxation-structure-composition of the outermost surface of the biofilms, opposite to data derived upon inducing 50% deformation that invokes the deeper layers of the biofilms. This being true for the images selected, it must be realized that it is difficult if not impossible by human nature to obtain confocal laser scanning microscopic (CLSM) images of biofilms in an unbiased, observer-independent way. This is why conclusions on biofilm structure from quantitative, observer-independent stress relaxation analysis of larger sections of a biofilm than can ever be obtained

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microscopically, are to be preferred. Interestingly, upon increasing the induced deformation to 50%, a better resemblance between in vivo and CDFF grown biofilms appears. This is probably because biofilms formed in vivo are compacted more than when formed in a PPFC through the presence of multiple strains and species that can more easily arrange themselves spontaneously through their differences in size and shape to a compact mass, even in the absence of external compaction or mechanical perturbations. For single-species biofilms grown in a CDFF, this compaction is achieved by continuously scraping off the biofilm by a rotating blade. Therefore it can be expected that oral biofilm in fissures and interproximal spaces, left behind multiple times after brushing, will eventually become compacted and better resemble biofilms formed in the CDFF than oral biofilms freshly formed, for which the PPFC may be the preferred model system.

The in vivo relations between relaxation characteristics and chlorhexidine penetration have larger 95% confidence intervals than the in vitro ones, partly due to the limited power of the study that was confined to five volunteers. More importantly however, it is intrinsically impossible to obtain the same narrow confidence intervals for in vivo biofilms as found for in vitro biofilms, that were all single-species. In our analyses, we employ chlorhexidine killing as an indicator of its penetration. In vivo formed biofilms contain a large number of different strains and species, that all have their own susceptibility to chlorhexidine not only within one volunteer, but also among volunteers. This inevitably affects the penetration as indicated by bacterial killing of chlorhexidine, making the in vivo relation less significant than the one obtained for in vitro biofilms.

In summary, this study is the first to demonstrate a role of viscoelastic properties of oral biofilm on antimicrobial penetration through a relaxation-structure-composition relationship. Herewith, biofilm viscoelasticity becomes an important quantifiable physical property of biofilms next to qualitative, observer-dependent CLSM-imaging of structure, with respect to advancing our understanding of antimicrobial penetration in biofilms. Although the current study was performed on oral biofilms, its applicability will extend to biofilms formed in other industrial and biomedical applications. Especially in the biomedical field, understanding the factors that control the penetration of antibiotics into biofilms is of utmost importance, as difficult to treat biofilm-related infections occur across all medical sub-disciplines causing large patients morbidity and mortality and inflicting huge costs to the health care system.

ACKNOWLEDGEMENTS

We thank Mr. Yun Chen for his help in data processing and Mrs. Jelly Atema-Smit for the help with CLSM.

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REFERENCES

1. Hall-Stoodley L, Costerton JW, Stoodley P (2004) Bacterial biofilms: From the natural environment to infectious diseases. Nat Rev Microbiol 2: 95-108.Flemming HC, Wingender J (2010) The biofilm matrix. Nat Rev Microbiol 8: 623-633.2. Driffield K, Miller K, Bostock JM, O’Neill AJ, Chopra I (2008) Increased mutability of Pseudomonas aeruginosa in biofilms. J Antimicrob Chemother 61: 1053-1056.3. Hoiby N, Bjarnsholt T, Givskov M, Molin S, Ciofu O (2010) Antibiotic resistance of bacterial biofilms. Int J Antimicrob Agents 35: 322-332.4. Boles BR, Singh PK (2008) Endogenous oxidative stress produces diversity and adaptability in biofilm communities. Proc Natl Acad Sci U S A 105: 12503-12508.5. Post JC, Stoodley P, Hall-Stoodley L, Ehrlich GD (2004) The role of biofilms in otolaryngologic infections. Curr Opin Otolaryngol Head Neck Surg 12: 185-190.6. Brown MR, Allison DG, Gilbert P (1988) Resistance of bacterial biofilms to antibiotics: A growth-rate related effect? J Antimicrob Chemother 22: 777-780.Tiirola M, Lahtinen T, Vuento M, Oker-Blom C (2009) Early succession of bacterial biofilms in paper machines. J Ind Microbiol Biotechnol 36: 929-937.7. Wolff D, Frese C, Maier-Kraus T, Krueger T, Wolff B (2012) Bacterial biofilm composition in caries and caries-free subjects. Caries Res 47: 69-77.Davis SC, Martinez L, Kirsner R (2006) The diabetic foot: The importance of biofilms and wound bed preparation. Curr Diab Rep 6: 439-445.8. Busscher HJ, Van der Mei HC, Subbiahdoss G, Jutte PC, Van den Dungen JJ, et al. (2012) Biomaterial-associated infection: Locating the finish line in the race for the surface. Sci Transl Med 4: 153rv10.9. Sambunjak D, Nickerson JW, Poklepovic T, Johnson TM, Imai P, et al. (2011) Flossing for the management of periodontal diseases and dental caries in adults. Cochrane Database Syst Rev 12: CD008829. Available: http://www.thecochranelibrary.com. Accessed 28 October 2011.10. Zaura-Arite E, Van Marle J, Ten Cate JM (2001) Confocal microscopy study of undisturbed and chlorhexidine-treated dental biofilm. J Dent Res 80:

1436-1440.11. Massol-Deya AA, Whallon J, Hickey RF, Tiedje JM (1995) Channel structures in aerobic biofilms of fixed-film reactors treating contaminated groundwater. Appl Environ Microbiol 61: 769-777.12. Picioreanu C, Van Loosdrecht MC, Heijnen JJ (1998) Mathematical modeling of biofilm structure with a hybrid differential-discrete cellular automaton approach. Biotechnol Bioeng 58: 101-116.13. Paramonova E, Kalmykowa OJ, Van der Mei HC, Busscher HJ, Sharma PK (2009) Impact of hydrodynamics on oral biofilm strength. J Dent Res 88: 922-926.14. Purevdorj B, Costerton JW, Stoodley P (2002) Influence of hydrodynamics and cell signaling on the structure and behavior of Pseudomonas aeruginosa biofilms. Appl Environ Microbiol 68: 4457-4464.15. Lau PC, Dutcher JR, Beveridge TJ, Lam JS (2009) Absolute quantitation of bacterial biofilm adhesion and viscoelasticity by microbead force spectroscopy. Biophys J 96: 2935-2948.16. Guelon T, Mathias J, Stoodley P (2011) Advances in biofilm mechanics. In: Flemming HC, Wingender J, Szewzyk U, editors. Biofilm Highlights. Heidelberg: Springer. pp. 111-140.17. Peterson BW, Busscher HJ, Sharma PK, Van der Mei HC (2012) Environmental and centrifugal factors influencing the visco-elastic properties of oral biofilms in vitro. Biofouling 28: 913-920.18. Stewart PS (2003) Diffusion in biofilms. J Bacteriol 185: 1485-1491.19. Busscher HJ, Van der Mei HC (2006) Microbial adhesion in flow displacement systems. Clin Microbiol Rev 19: 127-141.20. Hope CK, Wilson M (2006) Biofilm structure and cell vitality in a laboratory model of subgingival plaque. J Microbiol Methods 66: 390-398.21. Corbin A, Pitts B, Parker A, Stewart PS (2011) Antimicrobial penetration and efficacy in an in vitro oral biofilm model. Antimicrob Agents Chemother 55: 3338-3344.22. Van der Mei HC, White DJ, Atema-Smit J, Van de Belt-Gritter E, Busscher HJ (2006) A method to study sustained antimicrobial activity of rinse and dentifrice components on biofilm viability in vivo. J Clin

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Periodontol 33: 14-20.23. Takenaka S, Trivedi HM, Corbin A, Pitts B, Stewart PS (2008) Direct visualization of spatial and temporal patterns of antimicrobial action within model oral biofilms. Appl Environ Microbiol 74: 1869-1875.24. Lau PC, Lindhout T, Beveridge TJ, Dutcher JR, Lam JS (2009) Differential lipopolysaccharide core capping leads to quantitative and correlated modifications of mechanical and structural properties in Pseudomonas aeruginosa biofilms. J Bacteriol 191: 6618-6631.25. Sjollema J, Rustema-Abbing M, Van der Mei HC, Busscher HJ (2011) Generalized relationship between numbers of bacteria and their viability in biofilms. Appl Environ Microbiol 77: 5027-5029.

26. Hope CK, Wilson M (2004) Analysis of the effects of chlorhexidine on oral biofilm vitality and structure based on viability profiling and an indicator of membrane integrity. Antimicrob Agents Chemother 48: 1461-1468.27. Busscher HJ, Jager D, Finger G, Schaefer N, Van der Mei HC (2010) Energy transfer, volumetric expansion, and removal of oral biofilms by non-contact brushing. Eur J Oral Sci 118: 177-182.

Chapter 6Synergy of brushing mode and antibacterial use on in vivo

biofilm formation

Marije A. Jongsma and Marieke van de Lagemaat, Henk J. Busscher, Gesinda I. Geertsema-Doornbusch, Jelly Atema-Smit, Henny C. van der Mei,

Yijin Ren.

Submitted to Journal of Dentistry

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Effects of oral antimicrobials on brushed biofilmsChapter 6

ABSTRACT

Objectives: Orthodontic, multi-strand retention-wires are used as a generalized model for oral retention sites to investigate whether biofilm left-behind after powered toothbrushing in-vivo enabled better penetration of antibacterials as compared with manual brushing.

Methods: 2-cm multi-strand, stainless-steel retention-wires were placed in brackets bonded bilaterally in the upper arches of 10-volunteers. Volunteers used NaF-sodium-lauryl-sulphate-containing toothpaste and antibacterial, triclosan-containing toothpaste supplemented or not with an essential-oils containing mouthrinse. Opposite sides of the dentition including the retention-wires, were brushed manually or with a powered toothbrush. Health-care-regimens were maintained for 1-week, after which wires were removed and oral biofilm was collected.

Results: When powered toothbrushing was applied, slightly less bacteria were collected than after manual brushing, regardless whether an antibacterial-regimen was used or not. Powered-toothbrushing combined with antibacterial-regimens yielded lower biofilm viability than manual brushing, indicating better antibacterial penetration into biofilm left-behind after powered brushing. Major shifts in biofilm composition, with a decrease in prevalence of both cariogenic species and periodontopathogens, were induced after powered brushing using an antibacterial-regimen.

Conclusion: Oral biofilm left-behind after powered brushing in-vivo enabled better penetration of antibacterials than after manual brushing.

Clinical significance: Mechanical removal of oral biofilm is important for prevention of dental pathologies, but biofilm is always left-behind, such as in fissures, buccal pits, interproximal areas and gingival margins and around orthodontic appliances. Use of antibacterial toothpastes or mouthrinses can contribute to removal or killing of biofilm bacteria, but biofilm structure hampers antibacterial penetration. A synergy between brushing mode and antibacterial-regimen applied exists with clinically demonstrable effects.

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INTRODUCTION

Amount, viability and composition of oral biofilm play a major role in the development of oral pathologies, such as caries, gingivitis and periodontitis. Prevention of biofilm-related oral pathologies can be achieved either by mechanical or chemical removal of biofilm, changing its composition or preventing its formation.1 Mechanical biofilm removal by powered toothbrushing has been demonstrated to be superior to manual brushing.2 However, complete biofilm removal can never be achieved and after a single self-performed brushing, the amount of oral biofilm can only be reduced by 50-60%,3,4 leaving biofilm behind at locations out of reach for mechanical removal such as fissures, buccal pits, posterior interproximal areas and gingival margins, where oral pathologies mostly develop.5 In orthodontic patients, the number of locations out of reach of mechanical removal is even higher, making orthodontic patients more prone to oral pathologies than non-orthodontic patients.6

The use of antibacterial containing toothpastes or mouthrinses can be a valuable addendum to mechanical biofilm control in order to reduce the viability of biofilm left-behind after brushing.1 However, the general structure and composition of oral biofilm hampers penetration of oral antibacterials through the depth of an entire biofilm.7 Oral biofilm consists of a large variety of adhering bacteria embedded in an extracellular-polymeric-matrix that acts both as a glue for bacteria as well as a barrier against penetration of antibacterials.8,9 Powered toothbrushing of in vitro oral biofilm has been demonstrated to impact the structure of biofilm left-behind to create a more open structure, more amenable to antibacterial penetration,10 especially when the bristles of the brush have not been able to touch the biofilm and remove it.11 This more open structure is caused by a high energy transfer from a powered toothbrush into the biofilm through strong fluid flows,12 air bubble inclusion13 and acoustic waves.11 Accordingly it has been demonstrated in vitro that due to this more ‘fluffed-up’, open biofilm structure chlorhexidine and cetylpyridinium-chloride penetrate and kill bacteria to a greater depth into biofilm left-behind after powered brushing.14 Also, once oral antibacterials have penetrated the biofilm, the biofilm left-behind might act as a reservoir for the oral antibacterial agent ensuring a prolonged action of the agent.15 The impact of these in vitro findings for the clinical situation has never been demonstrated and could only be speculated upon, however.

In order to determine whether the improved penetration of antibacterial agents into biofilm left-behind after powered brushing as observed in vitro, also yields clinical benefits, we here aim to compare biofilm formation and composition in vivo on orthodontic, multi-strand retention wires after manual versus powered toothbrushing using a control, NaF- sodium lauryl sulphate containing toothpaste or an antibacterial, triclosan-containing toothpaste supplemented or not with the use of an essential-oils containing mouthrinse. Orthodontic,

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multi-strand retention wires are known to be difficult to clean16,17 and were employed as a generalized model for oral retention sites. Different regimens of oral health care were maintained for 1-week in a group of volunteers, equipped with multi-strand, stainless steel retention wires, after which oral biofilm left-behind after different modes of brushing was evaluated.

MATERIALS & METHODS

Retention wires, volunteers, inclusion criteria and oral hygiene regimens In this study, biofilm growth was evaluated on multi-strand, stainless steel retention wires (Quadcat®, PG Supply, Inc., Avon, USA), serving as a model for oral sites that are difficult to reach with a toothbrush. In addition, retention wires are easily removable for evaluation of biofilm formed. Brackets (SPEED System Orthodontics, Cambridge, Canada) were bonded to the buccal side of the first molar and the second premolar bilaterally in the upper arch of 10 healthy volunteers (5 male, 5 female) in agreement with the rules set out by the Ethics Committee at the University Medical Centre Groningen (letter June 23rd, 2011). Volunteers were included in the study, provided that they had a healthy and complete dentition, no bleeding upon probing and did not use any medication. All volunteers granted a written informed consent. Wires with a length of 2 cm were placed between the brackets. The wires were sterilized in 70% ethanol before use and stayed in situ for one week during which the volunteers were instructed to brush twice a day for 2 min with a manual toothbrush (Lactona iQ X-Soft, Lactona Europe B.V., Bergen op Zoom, The Netherlands) on one side of the dentition or with a powered toothbrush (Sonicare DiamondClean®, Philips Nederland B.V., Eindhoven, The Netherlands) on the other side. Volunteers were furthermore instructed to use a NaF-sodium lauryl sulphate (NaF-SLS) containing toothpaste without antibacterial claims (Prodent Softmint®, Sara Lee Household & Bodycare, Exton, USA), or a triclosan-containing toothpaste (Colgate Total®, Colgate-Palmolive Company, Piscataway, USA) with antibacterial claims. In addition, the use of the triclosan containing toothpaste was supplemented with the use of an essential-oils containing mouthrinse (Cool Mint Listerine®, Pfizer Consumer Healthcare, Morris Plains, NJ, USA). The order in which the regimens were applied in the different volunteers was determined at random. In between regimens and before the start of the experiment, a washout period of 6 weeks was applied during which only the NaF-SLS containing toothpaste was allowed to be used. The duration of the washout period was based on the results of a pilot study that indicated that the composition of the oral biofilm returned to base line values within 5 weeks after use of an antibacterial toothpaste.

Regimens were maintained for 1 week, after which wires were removed and oral biofilm was collected from the wires and the buccal enamel surfaces surrounding the brackets using a

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cotton swab. Wires were removed in the morning after breakfast and regular brushing by the volunteers. Wires and enamel biofilms collected were stored in an Eppendorf tube containing 1.0 mL filter sterile reduced transport fluid (RTF)18 for transportation from the orthodontic clinic to the laboratory.

Upon arrival in the laboratory, retention wires with adhering biofilm and biofilm collected from enamel surfaces were separately sonicated three times for 10 s with 30 s intervals in Eppendorf tubes containing 1.0 mL RTF on ice chilled water, to disperse bacteria. Part of the bacterial dispersions were stored at -80°C until use for PCR- Denaturing Gradient Gel Electrophoresis (DGGE), while another part was used to determine bacterial number and viability. For enumeration of the numbers of adhering bacteria, bacteria were enumerated in a Bürker-Türk counting chamber, while the percentage viability of the biofilms was evaluated after live/dead staining (BacLightTM, Invitrogen, Breda, The Netherlands) of the dispersed biofilms. Live/dead stain was prepared by adding 3 μL of SYTO®9/propidium iodide (1:3) to 1 mL of sterile, demineralised water. Fifteen μL of the stain was added to 10 μL of the undiluted bacterial dispersion. After 15 min incubation in the dark, the number of live and dead bacteria were counted using a fluorescence microscope (Leica DM4000B, Leica Microsystems Heidelberg GmbH, Heidelberg, Germany) and expressed as a percentage viability. Note that strictly speaking, live/dead staining is not a measure of microbial killing but of membrane damage.19,20 The membrane of live bacteria is permeable to SYTO9, staining both live and dead organisms and yielding green fluorescence. Propidium-iodide can only enter through damaged membranes, where it replaces SYTO9, yielding red fluorescence of dead or damaged cells.

DGGE analysis of in vivo biofilmsAfter all dispersed biofilms were collected, PCR-DGGE was carried out in order to compare their bacterial composition, as described previously.17 Briefly, for extraction of DNA, frozen bacterial dispersions were thawed, centrifuged for 5 min at 13,000 g, washed and vortexed with 200 μL TE-buffer (10 mM Tris-HCl, 1 mM EDTA pH 7.4) and again centrifuged. After DNA extraction, PCR was performed on 100 ng DNA with a T-gradient thermocycler for PCR amplifications. PCR products were analyzed by electrophoresis on a 2.0% agarose gel containing 0.5 μg/mL ethidium bromide. DGGE of PCR products generated with the F357-GC/R-518 primer set was performed, as described by Muyzer et al..21 The PCR products were applied on 0.08 g/mL polyacrylamide gel in 0.5 X TAE buffer (20 mM Tris acetate, 10 mM sodium acetate, 0.5 mM EDTA, pH 8.3). The denaturing gradient consisted of 30 to 80% denaturant (100% denaturant equals 7 M urea and 37% formamide). A 10 mL stacking gel without denaturant was added on top. Electrophoresis was performed overnight at 120 V and 60°C. Gels were stained with silver nitrate.22 Each DGGE gel was normalized according

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to a marker consisting of 7 reference species comprising common bacterial species associated with oral health and disease.23 The reference strains included Streptococcus oralis ATCC 35037, Streptococcus mitis ATCC 9811, Streptococcus sanguinis ATCC 10556, Streptococcus salivarius HB, Actinomyces naeslundii ATCC 51655, Lactobacillus sp., Streptococcus sobrinus ATCC 33478, Streptococcus mutans ATCC 10449, Porphyromonas gingivalis ATCC 33277 and Prevotella intermedia ATCC 49046.15

Statistical analysisData were analyzed with the Statistical Package for Social Sciences (Version 16.0, SPSS Inc., Chicago, IL, USA). A one-way analysis of variance (ANOVA) was used to compare the number of bacteria and their percentage viability. A Bonferroni test was used for post-hoc multiple comparisons. Statistical significance was set at p < 0.05.

DGGE gel images were converted and transferred into a microbial database with GelCompar II, version 6.1 (Applied Maths N.V, Sint-Martens-Latem, Belgium). Similarities in bacterial composition of the different biofilms were analysed using a band based similarity coefficient and a non-weighted pair group method with arithmetic averages was used to generate dendrograms indicating similarities in composition.24

RESULTS

When powered toothbrushing was applied, slightly less bacteria were collected from retention wires than after manual brushing, while enamel surfaces harvested insufficient amounts of biofilm for enumeration, providing a validation for the use of orthodontic, multi-strand retention wires as a model for oral retention sites, Within the regimens involving manual brushing, there were no significant differences (p < 0.05) in the numbers of bacteria collected from retention wires after use of a NaF-SLS-containing toothpaste and the use of an antibacterial, triclosan-containing toothpaste, regardless of supplementation with an essential-oils containing mouthrinse (Table 1). When powered toothbrushing was applied however, significantly less bacteria (p < 0.01) were collected when using the antibacterial, triclosan-containing toothpaste whether or not supplemented with an essential-oils containing mouthrinse, than when using the NaF-SLS-toothpaste.

More strikingly, viability of retention wire biofilm was significantly lower (p < 0.001) after the use of the antibacterial, triclosan-containing toothpaste whether or not combined with an essential-oils containing mouthrinse, when compared to the use of a NaF-SLS-containing toothpaste regardless of the brushing method. Moreover, in case of an antibacterial regimen, biofilm viability was lower after brushing with a powered toothbrush than after manual brushing.

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Table 1. Number and viability of bacteria retrieved from 1 cm stainless steel retainer wires after manual or powered toothbrushing with a NaF-SLS and an antibacterial, triclosan-containing toothpaste supplemented or not with the use of an essential-oils containing mouthrinse. All data represent averages ± standard deviations over 10 different volunteers.

Oral health care regimen

Number of bacteria(Log-units)

%live bacteria

Manual brushing

Powered brushing

Manual brushing

Powered brushing

NaF-SLS toothpaste without antibacterial claims

7.9 ± 0.1 7.6 ± 0.1a 68 ± 12 a 60 ± 7 a


7.6 ± 0.2 7.3 ± 0.3 42 ± 8 28 ± 9b

Triclosan containing toothpaste + mouthrinse

7.5 ± 0.2 7.0 ± 0.2 37 ± 5 16 ± 4b

a different from other regimens with the same brushing modeb different from the other brushing mode within the same regimens

Bacterial composition of biofilms formed on retention wires and enamel under the influence of the different oral hygiene regimens and brushing modes are compared in cluster trees (Figures 1A and 1B). Mode of brushing has no influence on the clustering of bacterial composition data, neither on retention wires (Figure 1A) nor on enamel surfaces (Figure 1B). However, the antibacterial regimens clearly separate from the NaF-SLS regimen although this is more clear on the retention wires than on enamel surfaces.

These changes in bacterial composition can further be exemplified from the prevalence of the marker strains applied (see Table 2), although it is difficult to find consistent patterns in effects of manual versus powered brushing. However, powered brushing yields a consistent decrease in the prevalence of P. gingivalis, both for biofilm collected from retention wires and enamel. Also the prevalence of S. sanguinis is consistently lower in case of powered brushing, but this is only the case for biofilm collected from retention wires. On the other hand, the prevalence of S. oralis/S. mitis increases after the use of a powered toothbrush compared to a manual toothbrush. In general, stronger effects of antibacterial regimens on the prevalence of marker stains are seen on retention wires than on enamel surfaces. S. salivarius, Lactobacillus, S. mutans and P. gingivalis decrease in prevalence on retention wires after use of the antibacterial, triclosan-containing toothpaste and these decreases become more pronounced when use of the antibacterial toothpaste is supplemented with an essential-oils containing mouthrinse. Prevalence of S. oralis/S. mitis on retention wires increases after the use of an antibacterial regimen.

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Figure 1. Clustering trees describing the bacterial compositions of biofilm samples taken from stainless steel retention wires (A) or enamel surfaces (B) in different volunteers using manual or powered toothbrushing in combination with different healthcare regimens.

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Tabl

e 2.

Pre

vale

nce

of m

arke

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in b

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m s

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100

% in

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wire

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amel

wire

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S. o

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s20

7040

5020

5050

8080

4070

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san

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8080

2070

4060

3070

6030

4050

S. s

aliv

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2030

1010

3010

200

1010

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nae

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00

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2020

2020

1030

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00

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3010

3030

3070

3030

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S. m

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1050

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2010

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lis30

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020

1010

00

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00

00

1010

00

00

00

104


DISCUSSION

Stress-relaxation analysis of mechanically compressed biofilms has pointed out that the structure and water content of in vitro biofilm-left behind after powered brushing changes into a direction that makes it more amenable to penetration of chlorhexidine and cetylpiridinium chloride than after manual brushing.14 Here we demonstrate the clinical impact of these in vitro findings. Clinical impact involves a reduction in the viability of in vivo formed biofilms left-behind after powered brushing on retention sites upon the use of an antibacterial triclosan-containing toothpaste with or without supplementation with an essential-oils containing mouthrinse. Thus also clinically, a synergy between mode of brushing and antibacterial-regimen applied exists.

We chose to study in vivo biofilms as formed on orthodontic retention wires after different 1-week regimens of oral health care, as especially multi-strand retention wires possess multiple sites where biofilm is sheltered from mechanical and chemical attack.25 Therewith retention wires can be considered as a generalized model for biofilm-retention sites in the oral cavity, with as an additional advantage that they are easily replaceable. Biofilm will be more readily left-behind on such retention sites after brushing and in this respect it is telling that in accordance with literature,3,4 biofilm could be collected from retention wires both after manual as well as after powered brushing (see Table 1), but hardly from smooth enamel surfaces. Powered toothbrushing generates a larger energy input into a biofilm than manual toothbrushing.26 Since biofilms have visco-elastic properties, biofilm will first expand during powered brushing after which it will detach.27-29 However, biofilm left-behind will remain in its expanded, more open state enabling better antibacterial penetration, which explains why in the current study we observe a greater reduction in biofilm viability upon application of antibacterial regimens when using a powered brush versus a manual brush. Note that the use of either one of the brushing methods without the use of an oral antibacterial regimen hardly affected the viability of the biofilm compared to an unbrushed biofilm.25 This indicates that the decrease in viability is solely attributed to the oral antibacterial agents, and not to toothbrushing itself.30 Therewith this is the first time to show the existence of a synergy between mode of toothbrushing and antibacterial action with clinically demonstrable effects.

Also other clinical studies, not geared towards demonstrating a synergy between mode of brushing and antibacterial use, have shown that oral biofilm formation is reduced after the use of antibacterial toothpastes,31,32 with minor effects of the supplemental use of an essential-oils containing mouthrinse.33-35 However, we saw sizeable further reduction of biofilm viability after supplemental use of an essential-oils containing rinse (Table 1), along with changes in bacterial composition of the biofilm (Figure 1) that we earlier attributed to adsorption of triclosan to bacterial cell surfaces altering their cell surface hydrophobicity to

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stimulate removal by hydrophobic ligands.17

DGGE analysis shows that the composition of biofilm formed on stainless steel retention wires differs from biofilm formed on enamel (Table 2). Atomic force microscopy has pointed out that bacterial adhesion forces to different materials used in orthodontics, including stainless steel, differ from the ones exerted by enamel surfaces in a strain-specific fashion.36 Accordingly this explains37 why biofilms on different materials have a different bacterial composition, including the enamel and stainless steel surfaces as involved here. Furthermore, the biofilm taken from retention wires will be more mature than biofilm taken from smooth enamel surfaces, as more biofilm will be left-behind after brushing on retention wires than on smooth enamel surfaces on which biofilm has to develop newly after each brushing. The composition of a newly formed biofilm as regularly developing on smooth enamel is thus different than that from a mature biofilm as in interproximal areas and fissures,38 the latter likely being comparable with biofilm found on the retention wires.

CONCLUSIONS

This is the first study to show that a synergy exists between powered toothbrushing and antibacterial regimen with clinically demonstrable effects, most notably on the viability of biofilm left-behind after brushing. Enhancing this synergy may be a goal of further research, either by changing the design of powered toothbrushes or use of different oral antibacterials. Since oral sites where biofilm is most frequently left-behind are also most susceptible to disease, this approach may proof to have major impact on oral health.

ACKNOWLEDGEMENTS

This study was entirely funded by UMCG, Groningen, The Netherlands. The authors would like to extend their gratitude to the volunteers who participated in this study and Ortholab B.V. (Doorn, The Netherlands) for kindly providing the different wires used in this study.

CONFLICT OF INTERESTS

The authors declare no potential conflicts of interest with respect to authorship and/or publication of this article. HJB is also director of a consulting company SASA BV, Thesinge, The Netherlands. Opinions and assertions contained herein are those of the authors and are not construed as necessarily representing views of the companies who donated the different wires or their respective employees.

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REFERENCES

1. Marsh PD (2012) Contemporary perspective on plaque control. Br Dent J 212:601-606 2. Yaacob M, Worthington HV, Deacon SA, Deery C, Walmsley AD, Robinson PG, Glenny AM (2014) Powered versus manual toothbrushing for oral health. Cochrane Database Syst Rev 6:CD002281 3. Paraskevas S, Timmerman MF, Van der Velden U, Van der Weijden GA (2006) Additional effect of dentifrices on the instant efficacy of toothbrushing. J Periodontol 77:1522-1527 4. Van der Weijden GA, Echeverria JJ, Sanz M, Lindhe J (2008) Mechanical Supragingival Plaque Control. In: J. Lindhe, N. P. Lang, and T. Karring (ed) Clinical Periodontology and Implant Dentistry, 5th edn. Blackwell Munskgaard, Copenhagen, pp 705-733 5. Sheiham A, Sabbah W (2010) Using universal patterns of caries for planning and evaluating dental care. Caries Res 44:141-150 6. Ren Y, Jongsma MA, Mei L, Van der Mei HC, Busscher HJ (2014) Orthodontic treatment with fixed appliances and biofilm formation-a potential public health threat? Clin Oral Investig 18:1711-17187. Van Leeuwenhoek AP (1684) Containing some microscopical observations, about animals in the scrurf of the teeth. Philosophical Transact Royal Soc London 14:568-574 8. Marsh PD (2010) Microbiology of dental plaque biofilms and their role in oral health and caries. Dent Clin North Am 54:441-454 9. Flemming HC, Wingender J (2010) The biofilm matrix. Nat Rev Microbiol 8:623-633 10. He Y, Peterson BW, Jongsma MA, Ren Y, Sharma PK, Busscher HJ, Van der Mei HC (2013) Stress relaxation analysis facilitates a quantitative approach towards antibacterial penetration into biofilms. PLoS One 8:e63750 11. Busscher HJ, Jager D, Finger G, Schaefer N, Van Der Mei HC (2010) Energy transfer, volumetric expansion, and removal of oral biofilms by non-contact brushing. Eur J Oral Sci 118:177-182 12. Van der Mei HC, Rustema-Abbing M, Bruinsma GM, Gottenbos B, Busscher HJ (2007) Sequence of oral bacterial co-adhesion and non-contact brushing. J Dent Res 86:421-425 13. Parini MR, Pitt WG (2006) Dynamic removal

of oral biofilms by bubbles. Colloids Surf B Biointerfaces 52:39-46 14. He Y, Peterson BW, Ren Y, Van der Mei HC, Busscher HJ (2014) Antibacterial penetration in a dual-species oral biofilm after noncontact brushing: an in vitro study. Clin Oral Investig 18:1103-1109 15. Otten MP, Busscher HJ, Abbas F, Van der Mei HC, Van Hoogmoed CG (2012) Plaque-left-behind after brushing: intra-oral reservoir for antibacterial toothpaste ingredients. Clin Oral Investig 16:1435-1442 16. Levin L, Samorodnitzky-Naveh GR, Machtei EE (2008) The association of orthodontic treatment and fixed retainers with gingival health. J Periodontol 79:2087-2092 17. Jongsma MA, Van der Mei HC, Atema-Smit J, Busscher HJ, Ren Y (2014) In vivo biofilm formation on stainless steel bonded-retainers during different regimens of oral health care. Int J Oral Science doi:10.1038/ijos.2014.69 18. Syed SA, Loesche WJ (1972) Survival of human dental plaque flora in various transport media. Appl Microbiol 24:638-644 19. Shi L, Gunther S, Hubschmann T, Wick LY, Harms H, Muller S (2007) Limits of propidium iodide as a cell viability indicator for environmental bacteria. Cytometry A 71:592-598 20. Netuschil L, Auschill TM, Sculean A, Arweiler NB (2014) Confusion over live/dead stainings for the detection of vital microorganisms in oral biofilms--which stain is suitable?. BMC Oral Health 14:2-6831-14-2 21. Muyzer G, De Waal EC, Uitterlinden AG (1993) Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl Environ Microbiol 59:695-700 22. Zijnge V, Welling GW, Degener JE, Van Winkelhoff AJ, Abbas F, Harmsen HJ (2006) Denaturing gradient gel electrophoresis as a diagnostic tool in periodontal microbiology. J Clin Microbiol 44:3628-3633 23. Marsh PD (2006) Dental plaque as a biofilm and a microbial community - implications for health and disease. BMC Oral Health 6 Suppl 1:S14 24. Signoretto C, Bianchi F, Burlacchini G, Sivieri F, Spratt D, Canepari P (2010) Drinking habits are associated with changes in the dental plaque microbial

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community. Journal of Clinical Microbiology 48:347-356 25. Jongsma MA, Pelser FD, Van der Mei HC, Atema-Smit J, Van de Belt-Gritter B, Busscher HJ, Ren Y (2013) Biofilm formation on stainless steel and gold wires for bonded retainers in vitro and in vivo and their susceptibility to oral antibacterials. Clin Oral Investig 17:1209-1218 26. Veeregowda DH, Van der Mei HC, De Vries J, Rutland MW, Valle-Delgado JJ, Sharma PK, Busscher HJ (2012) Boundary lubrication by brushed salivary conditioning films and their degree of glycosylation. Clin Oral Investig 16:1499-1506 27. Peterson BW, He Y, Ren Y, Zerdoum A, Libera MR, Sharma PK, Van Winkelhoff AJ, Neut D, Stoodley P, van der Mei HC, Busscher HJ (2014) Viscoelasticity of biofilms and their recalcitrance to mechanical and chemical challenges. FEMS Microbiol Rev Accepted 28. Rmaile A, Carugo D, Capretto L, Aspiras M, De Jager M, Ward M, Stoodley P (2014) Removal of interproximal dental biofilms by high-velocity water microdrops. J Dent Res 93:68-73 29. Cense AW, Peeters EA, Gottenbos B, Baaijens FP, Nuijs AM, Van Dongen ME (2006) Mechanical properties and failure of Streptococcus mutans biofilms, studied using a microindentation device. J Microbiol Methods 67:463-472 30. MacNeill S, Walters DM, Dey A, Glaros AG, Cobb CM (1998) Sonic and mechanical toothbrushes. An in vitro study showing altered microbial surface structures but lack of effect on viability. J Clin Periodontol 25:988-993 31. Riley P, Lamont T (2013) Triclosan/copolymer containing toothpastes for oral health. Cochrane Database Syst Rev 12:CD010514

32. He T, Barker ML, Biesbrock A, Miner M, Amini P, Goyal CR, Qaqish J (2013) Evaluation of anti-gingivitis benefits of stannous fluoride dentifrice among triclosan dentifrice users. Am J Dent 26:175-179 33. Cortelli SC, Cortelli JR, Shang H, McGuire JA, Charles CA (2013) Long-term management of plaque and gingivitis using an alcohol-free essential oil containing mouthrinse: a 6-month randomized clinical trial. Am J Dent 26:149-155 34. Stoeken JE, Paraskevas S, Van der Weijden GA (2007) The long-term effect of a mouthrinse containing essential oils on dental plaque and gingivitis: a systematic review. J Periodontol 78:1218-1228 35. Tufekci E, Casagrande ZA, Lindauer SJ, Fowler CE, Williams KT (2008) Effectiveness of an essential oil mouthrinse in improving oral health in orthodontic patients. Angle Orthod 78:294-298 36. Mei L, Busscher HJ, Van der Mei HC, Chen Y, De Vries J, Ren Y (2009) Oral bacterial adhesion forces to biomaterial surfaces constituting the bracket-adhesive-enamel junction in orthodontic treatment. Eur J Oral Sci 117:419-426 37. Wessel SW, Chen Y, Maitra A, Van den Heuvel ER, Slomp AM, Busscher HJ, van der Mei HC (2014) Adhesion forces and composition of planktonic and adhering oral microbiomes. Journal of Dental Research 93:84-88 38. Marsh PD (2004) Dental plaque as a microbial biofilm. Caries Res 38:204-211

Chapter 7

General discussion

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General discussionChapter 7

In this thesis we have shown that the amount of biofilm formed on orthodontic retention wires depends on the wire type, i.e. single-strand or multi-strand. More importantly, we established that manual brushing and chemical control of oral biofilms are less effective on multi-strand wires as compared to on single-strand wires. Powered toothbrushing not only removed more biofilms but also provided sufficient energy for disrupting the structure of biofilm left-behind after brushing in a way that it facilitated better penetration of oral antimicrobials into the brushed biofilm.1 This observation is of great clinical relevance for the control of biofilms on multi-strand wires, more difficult to keep clean than single-strand wires.

Several factors play a role in oral biofilm formation on orthodontic retention wires. Surface roughness is one of the factors we found to be important in the amount of biofilm formation. Due to the wire morphology, the surface roughness of multi-strand wires is higher than that of single-strand wires. In chapters 3 and 4 we found that significantly more biofilm is formed on multi-strand wires compared to single-strand wires. This coincides with literature stating that surface roughness is the dominant factor in biofilm formation and adhesion to surfaces.2 A larger surface roughness increases the surface area to which biofilm adheres and protects it against mechanical removal.3 Crevices and niches in the multi-strand wires provide areas out of reach for mechanical removal, thus creating a protected region for biofilms to grow in.4 In chapter 3, retention wires were placed in vivo, both bucally and palatally and biofilm formation was evaluated in absence of toothbrushing. A significantly smaller amount of biofilm was formed on single-strand wires placed palatally compared to buccally placed ones. This can be explained by the cleansing effect of the tongue acting on biofilms formed on palatally placed wires, that is virtually absent for the buccally placed wires. Interestingly this difference could not be observed for the multi-strand wires, due to the protected growth of the biofilm in the crevices and niches in the wire, out of reach for mechanical removal forces exerted by the tongue.2 A similar effect was observed in chapter 4, where we found that mechanical removal of the biofilm by brushing of orthodontic retention wires with a manual toothbrush is more effective for the single-strand wires than for multi-strand wires.

In chapters 3, 4 and 6 we found that the use of oral antimicrobials affects biofilm formation on the retention wires. Antimicrobials significantly reduced the amount of biofilm formed, but the reductions observed are probably too small to be of clinical relevance. More importantly, they significantly lowered the viability of biofilm organisms and affected the composition of the biofilm. Altering the composition of the biofilm is more and more considered as a clinically desired goal of oral hygiene measures rather than complete removal of oral biofilm, since oral biofilm is part of the resident microflora and the healthy oral microbiome5 with its health advantages such as prevention of fungal overgrowth.5 In chapters 4 and 6 we have shown that by using different regimens of oral antimicrobials, a clear change in composition

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of oral biofilm can be achieved. Particularly the use of triclosan combined with an essential oils containing mouthrinse led to a distinct decrease in the prevalence of cariogenic species, such as Streptococcus mutans and Lactobacilli,6 when compared to a NaF-containing toothpaste without antibacterial claims. This reduction in the presence of these cariogenic species might point to a shift in the composition of the adhering oral microbiome in a more healthy direction. Interestingly, this effect was already visible after only one week of using the antimicrobial regime and may become more strongly expressed after prolonged use of the oral antimicrobials.

In chapter 6 we demonstrate that antimicrobial penetration into oral biofilms can be improved by the use of a powered toothbrush based on a mechanism revealed in chapter 5: antimicrobial penetration in biofilms depends on the viscoelastic properties of the biofilm, reflecting both structure and composition of the biofilm. The energy output of powered toothbrushes is transferred to the biofilm resulting in an expansion of the biofilm and a more ‘fluffed-up’ structure.1 Due to the more ‘fluffed-up’ and open structure of the biofilm, as evidenced by changes in its viscoelastic properties, penetration of antimicrobials increases. This leads to a significant decrease in the amount of biofilm compared to manual brushing, as well as a significant decrease in the viability of the biofilm. Improved antimicrobial penetration also has another beneficial effect: once oral antimicrobials have penetrated the biofilm, the biofilm left-behind acts as a reservoir for the oral antimicrobial ensuring a prolonged action of the agent.7

Summarizing, this thesis forwards two possible new pathways for oral biofilm control on orthodontic retention wires that may have relevance for oral hygiene in general:

1. The use of regimens of antibacterial toothpastes and subsequent mouthrinses to alter the composition of oral bacteria in the biofilm and subsequently remove them from the oral cavity through use of an appropriate rinse,

2. The use of powered toothbrushes to enhance the action of oral antimicrobials. Although significant reductions and shifts on oral biofilm composition support these new pathways, the outcome measures used are not directly related to clinical outcome parameters, such as an actual reduction in caries and gingivitis prevalence in orthodontic patients or patients wearing orthodontic bonded retainers. Such an extended clinical demonstration of the benefits of the pathways outlined in this study remains to be done before actual clinical recommendations can be made.

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REFERENCES

1. Busscher HJ, Jager D, Finger G, Schaefer N, Van Der Mei HC (2010) Energy transfer, volumetric expansion, and removal of oral biofilms by non-contact brushing. Eur J Oral Sci 118:177-182 2. Quirynen M, Bollen CM (1995) The influence of surface roughness and surface-free energy on supra- and subgingival plaque formation in man. A review of the literature. J Clin Periodontol 22:1-14 3. Lang PL, Mombelli A, Attström R (1997) Dental plaque and calculus. In: Lindhe J, Karring T, Lang PL (eds) Clinical periodontology and implant dentistry, 3rd edn. Munksgaard, Copenhagen, pp 102-137 4. Al-Nimri K, Al Habashneh R, Obeidat M (2009) Gingival health and relapse tendency: a prospective study of two types of lower fixed retainers.

Aust Orthod J 25:142-146 5. Marsh PD (2012) Contemporary perspective on plaque control. Br Dent J 212:601-606 6. Marsh PD (2006) Dental plaque as a biofilm and a microbial community - implications for health and disease. BMC Oral Health 6 Suppl 1:S14 7. Otten MP, Busscher HJ, Abbas F, Van der Mei HC, Van Hoogmoed CG (2012) Plaque-left-behind after brushing: intra-oral reservoir for antibacterial toothpaste ingredients. Clin Oral Investig 16:1435-1442

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Summary

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During active orthodontic treatment as well as in the retention phase after an active treatment, biofilm can cause complications such as gingivitis and white spot lesions. Prevention of these complications can be achieved either by mechanical removal or chemical biofilm control. However, orthodontic appliances and retention wires provide many crevices and niches in which biofilm can grow out of reach of mechanical removal, while the structure of a biofilm hampers penetration of antimicrobials into the biofilm to offer protection to organisms in a biofilm mode of growth. In Chapter 1 we hypothesize:

1. that biofilm formation is dependent on the wire type, since the crevices and niches in the multi-strand wires provide a protected environment for biofilm growth that is absent on single-strand wires,

2. that the effect of manual brushing and chemical biofilm control is smaller on multi-strand wires compared to single-strand wires,

3. that mechanical disruption of the structure of oral biofilm by powered tooth brushing will en hance antimicrobial action from toothpastes or mouthrinses as compared with manual brushing. Verification of the above hypotheses constitutes the general aim of this thesis.

Orthodontic treatment is highly popular for restoring oral facial function and esthetics in juveniles and adults. As a downside, prevalence of biofilm related complications is high. Literature on biofilm formation in the oral cavity is reviewed in Chapter 2 to identify special features of biofilm formation in orthodontic patients. Estimates are made of juvenile and adult orthodontic patient population sizes and biofilm-related complication rates are used to indicate the costs and clinical workload resulting from biofilm-related complications.

Biofilm formation in orthodontic patients is governed by similar mechanisms as common in the oral cavity. However, orthodontic appliances hamper maintenance of oral hygiene and provide numerous additional surfaces, with properties alien to the oral cavity, to which bacteria can adhere and form a biofilm. Biofilm formation may lead to gingivitis and white spot lesions, compromising facial esthetics. Whereas gingivitis after orthodontic treatment is often transient, white spot lesions may turn into cavities requiring professional restoration. Complications requiring professional care develop in 15% of all orthodontic patients, implying an annual cost of over US$ 500,000,000 and a workload of 1000 fulltime dentists in the USA alone.

Improved preventive measures and antimicrobial materials are urgently required to prevent biofilm-related complications of orthodontic treatment from overshadowing its functional and esthetic advantages. High treatment demand and occurrence of biofilm-related complications requiring professional care make orthodontic treatment a potential public health threat.

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Bonded retainers are used in orthodontics to maintain treatment result. Retention wires are prone to biofilm formation yielding greater incidence of gingival recession, bleeding on probing and increased pocket depths near bonded retainers. In Chapter 3 we compare in vitro and in vivo biofilm formation on different wires used for bonded retainers and the susceptibility of in vitro biofilms to oral antimicrobials.

To this end orthodontic wires were exposed to saliva and in vitro biofilm formation was evaluated using plate counting and live-dead staining, together with effects of exposure to toothpaste slurry alone or followed by antimicrobial mouthrinse application. Wires were also placed intra-orally for 72 hours in human volunteers and undisturbed biofilm formation was compared by plate counting and live-dead staining as well as by Denaturing Gradient Gel Electrophoresis for compositional differences in biofilms. Single-strand wires attracted only slightly less biofilm in vitro than multi-strand wires. Biofilms on stainless steel single-strand wires however, were much more susceptible to antimicrobials from toothpaste slurries and mouthrinses than on single-strand gold wires and biofilms on multi-strand wires. Also in vivo significantly less biofilm was found on single-strand than on multi-strand wires. Microbial composition of biofilms was more dependent on the volunteer involved than on wire type. Biofilms on single-strand stainless steel wires attract less biofilm in vitro and are more susceptible to antimicrobials than on multi-strand wires. Also in vivo, single-strand wires attract less biofilm than multi-strand ones. Therefore, use of single-strand wires is preferred over multi-strand wires, not because they attract less biofilm, but because biofilms on single-strand wires are not protected against antimicrobials as in crevices and niches as on multi-strand wires.

In Chapter 4 we compare in vivo biofilm formation on single-strand and multi-strand retention wires during different regimens of oral healthcare. Two-cm wires were placed in brackets bonded to the buccal side of first molars and second premolars in the upper arches of 22 healthy volunteers. Volunteers used a selected toothpaste with or without additional use of an essential-oils containing mouthrinse. Brushing was performed manually. Regimens were maintained for one week, after which wires were removed and oral biofilm was collected for enumeration of the number of organisms and their viability, microbial composition and electron microscopic visualization. Six weeks washout was applied in between regimens. Less biofilm was formed on single-strand wires than on multi-strand wires, on which bacteria were observed adhering in between strands. Use of antibacterial toothpastes only marginally decreased the amount of biofilm on both wire types, but significantly reduced the viability of biofilm organisms. No significant effects were observed on amount or viability of biofilms upon additional use of the mouthrinse. However, major shifts in biofilm composition were induced by combining a stannous fluoride or triclosan containing toothpaste with the essential-oils

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containing rinse. Tentatively, these shifts are attributed to small changes in bacterial cell surface hydrophobicity after adsorption of toothpaste components, that stimulate bacterial adhesion to hydrophobic oil, as illustrated for a Streptococcus mutans strain.

Biofilms are often tolerant to antimicrobials, due to a combination of inherent properties of bacteria in their adhering, biofilm mode of growth and poor physical penetration of antimicrobials through biofilms. Current understanding of biofilm recalcitrance toward antimicrobial penetration is based on qualitative descriptions of biofilms. In Chapter 5 we hypothesize that stress relaxation of biofilms will relate with antimicrobial penetration. Stress relaxation analysis of single-species oral biofilms grown in vitro identified a fast, intermediate and slow response to an induced deformation, corresponding with outflow of water and extracellular polymeric substances, and bacterial re-arrangement, respectively. Penetration of chlorhexidine into these biofilms increased with increasing relative importance of the slow and decreasing importance of the fast relaxation element. Involvement of slow relaxation elements suggests that biofilm structures allowing extensive bacterial re-arrangement after deformation are more open, allowing better antimicrobial penetration. Involvement of fast relaxation elements suggests that water dilutes the antimicrobial upon penetration to an ineffective concentration in deeper layers of the biofilm. Ex situ chlorhexidine penetration into two weeks old in vivo formed biofilms followed a similar dependence on the importance of the fast and slow relaxation elements as observed for in vitro formed biofilms. Chapter 5 therewith demonstrates that biofilm properties can be derived that quantitatively explain antimicrobial penetration into a biofilm.

Mechanical removal of oral biofilm is important for prevention of dental pathologies, but complete biofilm removal can never be achieved, especially not around orthodontic appliances. Use of antimicrobials can contribute to removal or killing of biofilm bacteria, but biofilm structure hampers antimicrobial penetration. It is known that oral biofilm left-behind after powered brushing in vitro possessed a more open structure, enabling better penetration of antimicrobials. In Chapter 6 we investigate whether biofilm left-behind on orthodontic retention wires after powered toothbrushing in vivo also enabled better penetration of antimicrobials as compared with manual brushing. To this end, two-cm stainless steel retention wires were placed in brackets bonded bilaterally to the buccal side of first molars and second premolars in the upper arches of 10 volunteers. Volunteers used a NaF-sodium lauryl sulphate containing toothpaste and an antimicrobial, triclosan containing toothpaste supplemented or not with the use of an essential-oils containing mouthrinse. Opposite sides of the dentition including the retention wires, were brushed manually or with a powered toothbrush. Regimens were maintained for 1-week, after which wires were removed and oral biofilm was collected. When powered toothbrushing was applied, slightly less bacteria were

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collected than for manual brushing, regardless of whether using an antimicrobial regimen or not. Strikingly, powered toothbrushing combined with an antimicrobial regimen yielded lower biofilm viability than manual brushing, indicating better antimicrobial penetration into biofilm left-behind after powered brushing. Also, major shifts in biofilm composition, with a decrease in prevalence of both cariogenic species and periodontopathogens, were induced after powered brushing using an antimicrobial regimen. This study herewith is the first to show that a synergy between mode of brushing and antimicrobial regimen exists with clinically demonstrable effects.

Summarizing, this thesis forwards two possible new pathways for oral biofilm control on orthodontic retention wires that may have relevance for oral hygiene in general which are discussed in Chapter 7:

1. The use of regimens of antimicrobial toothpastes and subsequent mouthrinses to alter the composition of oral bacteria in the biofilm and subsequently remove them from the oral cavity through use of an appropriate rinse.

2. The synergistic use of powered toothbrushes to enhance the action of oral antimicrobials.

Although significant reductions and shifts on oral biofilm composition support these new pathways, an extended clinical demonstration of the benefits of the pathways outlined in this study remains to be done before actual clinical recommendations can be made.

Nederlandse samenvatting

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Voor het behoud van het behandelresultaat na een orthodontische behandeling worden vaak retentiedraden achter de voortanden geplaatst. Deze draden kunnen verschillende vormen hebben welke kunnen worden gecategoriseerd als enkelstrengs retentiedraden en meerstrengs retentiedraden, waarbij laatstgenoemden bestaan uit verschillende dunnere draden die om elkaar gedraaid zijn. Zowel tijdens een orthodontische behandeling als in de fase daarna als de retentiedraden in de mond aanwezig zijn, kan biofilm complicaties veroorzaken in de mond zoals tandvleesontsteking (gingivitis) en witte vlek laesies. Biofilms zijn gemeenschappen van veel verschillende soorten bacteriën die zich in de mond hechten aan bijvoorbeeld het tandoppervlak, het tandvlees en de tong. De bacteriën produceren stoffen die dienen om hen te beschermen tegen invloeden van buitenaf en als lijm om goed aan elkaar en aan de ondergrond te kunnen hechten. Door deze bescherming en het feit dat de bacteriën in een biofilm samenwerken, zijn deze beter beschermd tegen bijvoorbeeld antibacteriële middelen dan losse bacteriën. Preventie van de complicaties die door biofilm in de mond veroorzaakt worden, kan worden bereikt door het verwijderen van de biofilm of door chemische bestijding van de biofilm met antimicrobiële middelen. Echter, orthodontische apparatuur en retentiedraden bieden veel ruimtes en nissen waarin biofilm kan groeien buiten het bereik van mechanische verwijdering, terwijl de structuur van de biofilm voorkomt dat antimicrobiële stoffen in de biofilm door kunnen dringen.

In Hoofdstuk 1 veronderstellen we dat de hoeveelheid biofilm die zich vormt op retentiedraden afhankelijk is van het type draad, omdat de spleten en nissen in de meerstrengs draden een beschermende omgeving voor de biofilm vormen. Hierdoor wordt het effect van handmatig verwijderen van de biofilm en chemische bestrijding door orale antimicrobiële middelen verminderd in vergelijking met enkelstrengs draden. Verder veronderstellen we dat om penetratie van antimicrobiële middelen in de biofilm te verbeteren, het gunstig is om de biofilm mechanisch te verstoren door het gebruik van een elektrische tandenborstel. De energie die de elektrische tandenborstel levert, is in staat om de structuur van de biofilm te veranderen, waardoor deze beter doordringbaar wordt voor antimicrobiële middelen. De verificatie van de bovenstaande hypothesen is de algemene doelstelling van dit proefschrift.

Orthodontische behandeling is zeer populair voor het herstellen van zowel functie als esthetiek van het aangezicht en de tanden bij jongeren en volwassenen. Een nadeel van orthodontische behandelingen is dat er vaak complicaties optreden die samenhangen met biofilm die zich op en rond de orthodontische apparatuur vormt, zoals cariës en gingivitis. In Hoofdstuk 2 wordt literatuur over de vorming van biofilm in de mondholte beoordeeld en wordt specifiek gekeken naar eigenschappen van de biofilm die zich vormt bij orthodontische patiënten. Er worden schattingen gemaakt over de omvang van de jeugdige en volwassen orthodontische patiëntenpopulatie. Aan de hand van deze gegevens wordt een schatting gemaakt van de jaarlijkse kosten en klinische werkbelasting van tandartsen die ontstaan als

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gevolg van biofilm gerelateerde complicaties na een orthodontische behandeling. Biofilm vormt zich bij orthodontische patiënten op een soortgelijke manier als gebruikelijk is in de mondholte. Echter, orthodontische apparatuur belemmert het schoonhouden van het gebit en biedt tal van extra oppervlakken waaraan bacteriën zich kunnen hechten om een biofilm te vormen. De biofilm hecht zich bovendien gemakkelijker aan de materialen die gebruikt worden voor de orthodontische apparatuur dan aan tandweefsel. Vorming van biofilm op en rondom de orthodontische apparatuur kan leiden tot gingivitis en witte vlek laesies welke afbreuk doen aan de esthetiek van het orthodontische eindresultaat. Gingivitis na een orthodontische behandeling is vaak van voorbijgaande aard, echter witte vlek laesies vereisen vaak behandeling door de tandarts, zeker als deze laesies zich hebben ontwikkeld tot caviteiten. Complicaties die professionele zorg door de tandarts nodig hebben ontwikkelen zich bij 15% van alle orthodontische patiënten, wat neerkomt op een jaarlijkse kostenpost van meer dan $500.000.000,- en een werkbelasting van 1000 fulltime tandartsen alleen al in de Verenigde Staten van Amerika.

Verbeterde preventieve maatregelen en antimicrobiële materialen zijn dringend nodig om te voorkomen dat biofilm gerelateerde complicaties van een orthodontische behandeling de functionele en esthetische voordelen van de behandeling overschaduwen. De hoge vraag aan orthodontische zorg en de hoge prevalentie van biofilm gerelateerde complicaties die professionele behandeling door de tandarts nodig maken, zorgen ervoor dat orthodontische behandeling een potentiële bedreiging vormt voor de volksgezondheid.

Retentiedraden die achter de voortanden worden geplaatst, worden gebruikt om na een orthodontische behandeling het eindresultaat vast te houden. Retentiedraden zijn gevoelig voor de vorming van biofilm waardoor rondom deze draden vaak sprake is van ontstoken en teruggetrokken tandvlees. In Hoofdstuk 3 vergelijken we in vitro en in vivo de vorming van biofilm op verschillende typen retentiedraden en beoordelen we de gevoeligheid van in vitro biofilms voor orale antimicrobiële middelen.

Orthodontische retentiedraden werden blootgesteld aan speeksel en de vorming van in vitro biofilm werd geëvalueerd door het tellen van de hoeveelheid bacteriën die gekweekt konden worden en door de bacteriën te kleuren om hun levensvatbaarheid te bepalen. De in vitro biofilm werd ook blootgesteld aan een tandpasta en aan een tandpasta gevolgd door een antibacterieel mondspoelmiddel om de effecten van antimicrobiële middelen op de in vitro biofilm te kunnen bepalen. Daarnaast werden verschillende typen retentiedraden gedurende 72 uur bij menselijke vrijwilligers in de mond geplaatst. Deze draden mochten niet gepoetst worden om ongestoorde vorming van biofilm te kunnen vergelijken op de verschillende draden. Ook de in vivo gevormde biofilm werd geëvalueerd door het tellen van de hoeveelheid bacteriën die gekweekt konden worden en door de bacteriën te kleuren

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voor het bepalen van de levensvatbaarheid. Daarnaast werd een DNA profiel gemaakt van de verschillende bacteriën in de biofilm door middel van denaturerende gradiënt gelelektroforese om daarmee de samenstelling van de verschillende biofilms te kunnen vergelijken.

Op enkelstrengs draden hechtte zich in vitro slechts iets minder biofilm dan op meerstrengs draden. Biofilms op enkelstrengs roestvrijstalen draden waren echter veel gevoeliger voor de antimicrobiële werking van tandpasta en mondspoelmiddelen dan biofilms op enkelstrengs gouden draden en biofilms op meerstrengs draden. Ook in vivo werd significant minder biofilm gevonden op enkelstrengs retentiedraden dan op meerstrengs retentiedraden. De bacteriële samenstelling van de biofilms die in vivo waren gevormd was meer afhankelijk van de betrokken vrijwilliger dan van het type draad.

Het plaatsen van enkelstrengs retentiedraden na een orthodontische behandeling heeft de voorkeur boven het plaatsen van meerstrengs retentiedraden, niet omdat zich minder biofilm op deze draden hecht, maar omdat biofilms op enkelstrengs draden in vitro gevoeliger zijn voor antimicrobiële middelen doordat deze niet wordt beschermd door spleten en in nissen in de draad zoals bij meerstrengs draden.

In Hoofdstuk 4 wordt in vivo vorming van biofilm op enkelstrengs en meerstrengs retentiedraden vergeleken tijdens verschillende regimes van mondverzorgingsproducten. Retentiedraden van twee centimeter lang werden tussen orthodontische brackets geplaatst die vastzitten aan de buccale zijde van de eerste molaren en tweede premolaren in de boventandboog van 22 vrijwilligers. Vrijwilligers gebruikten een geselecteerde tandpasta met of zonder aanvullend gebruik van een etherische oliën bevattend mondspoelmiddel. De draden werden gepoetst met een handtandenborstel. De regimes werden gedurende 1 week volgehouden, waarna de retentiedraden werden verwijderd en de orale biofilm werd verzameld voor de telling van het aantal bacteriën, het bepalen van hun levensvatbaarheid, het vaststellen van de bacteriële samenstelling van de biofilm en voor visualisatie van de biofilm door middel van een elektronenmicroscoop. Tussen de verschillende regimes poetsten de vrijwilligers gedurende 6 weken met een niet antibacteriële tandpasta om het effect van de antibacteriële middelen volledig uit te laten werken.

Op enkelstrengs retentiedraden werd minder biofilm gevormd dan op meerstrengs retentiedraden. De aanwezigheid van biofilm op de meerstrengs draden werd vooral waargenomen in de spleten en nissen in de draad, terwijl de biofilm op de enkelstrengs draden als een dunne over de draad verspreide film aanwezig was. Het gebruik van antibacteriële tandpasta verminderde de hoeveelheid biofilm op beide draadtypen marginaal, maar de levensvatbaarheid van de bacteriën in de biofilm werd significant verminderd door het

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gebruik van antibacteriële tandpasta. Er werden geen significante effecten waargenomen met betrekking tot de hoeveelheid of levensvatbaarheid van de biofilms na aanvullend gebruik van een antibacterieel etherische oliën bevattend mondspoelmiddel. Echter, grote verschuivingen in de samenstelling van de biofilm werden geïnduceerd door het combineren van een tinfluoride of triclosan bevattende tandpasta gecombineerd met een etherische oliën bevattend mondspoelmiddel. Voorlopig zijn deze verschuivingen toegeschreven aan kleine veranderingen in de hydrofobiciteit van het bacteriële celoppervlak na adsorptie van tandpasta componenten. Hierdoor wordt bacteriële hechting gestimuleerd aan hydrofobe etherische oliën, zoals geïllustreerd voor een Streptococcus mutans stam.

Biofilms zijn vaak minder gevoelig voor antimicrobiële stoffen door een combinatie van factoren die inherent zijn aan de manier waarop biofilms groeien en de slechte doordringing van antimicrobiële stoffen naar de diepere lagen van biofilms. Het huidige begrip met betrekking tot de beperkte penetratie van antimicrobiële stoffen in biofilms is voornamelijk gebaseerd op kwalitatieve beschrijvingen van biofilms. In Hoofdstuk 5 poneren we de hypothese dat stress-relaxatie van biofilms samenhangt met de penetratie van antimicrobiële stoffen. Stress-relaxatie analyse van in vitro gegroeide orale biofilm bestaande uit één soort bacteriën, toonde een snelle, middel-langzame en langzame reactie op de geïnduceerde vervorming, overeenkomend met respectievelijk de uitstroom van water, extracellulaire polymere substanties en herverdeling van bacteriën. De penetratie van chloorhexidine in deze biofilms nam toe met een toenemende waarde van het langzame en een afnemende waarde van het snelle element. De betrokkenheid van het langzame relaxatie element suggereerde dat biofilm structuren waarin na deformatie uitgebreide herverdeling van bacteriën kan plaatsvinden meer open zijn, waardoor de penetratie van antimicrobiële stoffen beter wordt. De betrokkenheid van het snelle relaxatie element suggereerde vervolgens dat in de diepere lagen van de biofilm, water de concentratie van antimicrobiële stoffen verlaagt tot ineffectieve waarden. Ex situ penetratie van chloorhexidine in twee weken oude, in vivo gegroeide biofilms toonde een zelfde afhankelijkheid van de snelle en langzame relaxatie elementen als in vitro gegroeide biofilms. Hoofdstuk 5 toont hiermee aan dat visco-elastische eigenschappen van biofilms een kwantitatieve verklaring kunnen geven voor de penetratie van antimicrobiële middelen.

Mechanische verwijdering van orale biofilm is belangrijk voor de preventie van tandheelkundige pathologieën, maar complete verwijdering van de biofilm kan nooit worden bereikt, zeker niet rond orthodontische apparatuur. Het gebruik van antibacteriële middelen kan bijdragen aan het verwijderen of doden van bacteriën in een biofilm, maar de structuur van een biofilm belemmert antimicrobiële penetratie. Het is bekend dat orale biofilm die in vitro wordt achtergelaten na elektrisch poetsen een open structuur heeft, waardoor betere penetratie van antibacteriële stoffen kan plaatsvinden. In Hoofdstuk 6

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onderzoeken we of biofilm die achterblijft op orthodontische retentiedraden na elektrisch tandenpoetsen in vivo ook een betere penetratie van antibacteriële stoffen mogelijk maakt in vergelijking met handmatig poetsen. Hiertoe werden retentiedraden van 2 centimeter lang tussen orthodontische brackets geplaatst die vastzaten aan de buccale zijde van de eerste molaren en tweede premolaren aan beide zijden van de boventandboog van 10 vrijwilligers. De vrijwilligers gebruikten een niet antibacteriële, natriumfluoride-natriumlaurylsulfaat bevattende tandpasta en een antimicrobiële, triclosan bevattende tandpasta al dan niet aangevuld met het gebruik van een etherische oliën bevattend mondspoelmiddel. De beide zijden van het gebit met inbegrip van de retentiedraden werden handmatig of met een elektrische tandenborstel gepoetst. De regimes werden gedurende 1 week volgehouden, waarna de draden werden verwijderd en de orale biofilm werd verzameld en geëvalueerd.

Wanneer de retentiedraden elektrisch werden gepoetst werden iets minder bacteriën gevonden dan wanneer de retentiedraden handmatig waren gepoetst, ongeacht of er een antimicrobieel regime was toegepast of niet. Opvallend is dat elektrisch tandenpoetsen gecombineerd met een antimicrobieel regime leidde tot een lagere levensvatbaarheid van de biofilm dan na handmatige poetsen, wat aangeeft dat er een betere penetratie is van antimicrobiële middelen in biofilm die achterblijft na elektrisch poetsen. Ook werden grote verschuivingen in de samenstelling van de biofilm geïnduceerd, met een daling van de prevalentie van zowel cariogene soorten als paropathogenen na elektrisch poetsen gecombineerd met een antimicrobieel regime. Hoofdstuk 6 laat hiermee voor het eerst zien dat er een synergie bestaat tussen de manier van borstelen en het gebruik van antimicrobiële middelen met klinisch aantoonbare effecten.

Dit proefschrift brengt twee mogelijke nieuwe wegen naar voren voor de preventie van orale biofilm op orthodontische retentie draden die relevant zijn voor de mondhygiëne in het algemeen. Deze worden besproken in Hoofdstuk 7:

1. Het gebruik van regimes van antimicrobiële tandpasta en daaropvolgende mondspoelmiddelen om de samenstelling van orale bacteriën in de biofilm te wijzigen en deze vervolgens te verwijderen uit de mondholte door het gebruik van een geschikt spoelmiddel.

2. Het synergistische gebruik van een elektrische tandenborstel om de werking van orale antibacteriële middelen te verhogen.

Dankwoord (acknowledgements)

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Dit proefschrift is het resultaat van 3 ½ jaar onderzoek bij de afdeling Orthodontie van het UMCG en bij de afdeling BioMedical Engineering, onderdeel van het W.J. Kolff Instituut. De eerste 2 jaar heb ik het doen van onderzoek gecombineerd met mijn opleiding tot orthodontist en de laatste 1 ½ jaar met het werken als orthodontist in mijn eigen praktijk. Als gevolg hiervan kon ik me maar 2 dagen per week met het onderzoek bezighouden. Dat het toch gelukt is om dit proefschrift te voltooien heb ik te danken aan een groot aantal mensen die mij op allerlei manieren hebben bijgestaan. Het is natuurlijk onmogelijk om al deze mensen bij naam te noemen, maar ik wil toch graag een aantal van hen persoonlijk bedanken.

Mijn eerste promotor Prof. dr. Yijin Ren, ik wil je heel erg bedanken voor de mogelijkheid die je me hebt gegeven om promotieonderzoek te doen. Dankzij jouw begeleiding heb ik me kunnen ontwikkelen als onderzoeker. Jij hebt me zowel tijdens mijn opleiding tot orthodontist als tijdens dit promotietraject altijd gemotiveerd om het beste uit mezelf te halen en daar ben ik je erg dankbaar voor.

Prof. dr. Henny van der Mei, heel erg bedankt voor de fijne manier waarop je me tijdens dit promotietraject begeleid hebt. Door jouw enorme wetenschappelijke kennis zette jij altijd de puntjes op de i waar ik weleens wat slordig kon zijn. Met al mijn vragen kon ik altijd bij je terecht, waardoor je me op allerlei terreinen veel geleerd hebt, bedankt hiervoor!

Prof. dr. Henk Busscher, de kennis die jij hebt is echt indrukwekkend en ik heb in de afgelopen periode enorm veel van je geleerd. De manier waarop je me hebt begeleid tijdens dit promotietraject heb ik als heel fijn en inspirerend ervaren. Het is voor mij een enorme eer geweest om met je samen te hebben mogen werken.

De beoordelingscommissie, prof. dr. S.K. Bulstra, prof. dr. J.M. ten Cate en prof. dr. He wil ik bedanken voor de tijd en aandacht die ze hebben besteed aan het doornemen van dit proefschrift. I would like to thank the reading committee, prof. dr. S.K. Bulstra, prof. dr. J.M. ten Cate and prof. dr. H. He for the time and attention they have put into reviewing this thesis.

Betsy van de Belt-Gritter, ik wil je bedanken voor de hulp die je me hebt gegeven om wegwijs te worden op het lab.

Marja Stiemsma-Slomp, bedankt voor de gezelligheid die je meebracht naar het lab en voor je hulp bij de live/dead kleuring.

Jelly Atema-Smit en Gesinda Geertsma-Doornbusch, de hoeveelheid werk die jullie voor mij gedaan hebben is echt enorm! Dankzij jullie is het gelukt om bij honderden samples DGGE uit te voeren. Jullie waren daarnaast ook hele fijne collega’s om mee samen te werken, bedankt!

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Jeroen Kuipers, bedankt voor je hulp bij het maken van de elektronenmicroscoop foto’s, zonder jouw begeleiding was dit niet gelukt!

Mei Li, thank you very much for introducing me to the world of microbiology in orthodontics. Your kind guidance in the first phase of my PhD project was of great help and has really helped to get me started.

Yan He, thank you for the great cooperation! It have had a great time working with you, I hope to see you again soon.

All my colleagues at the Department of Biomedical Engineering, Helen, Arina, Bu, Victor, Joana, Ferdi, Nina, Jan, Stefan, Barbara, Edward, Jesse, Mark, Raquel, Deepak, Agnieszka, Philip, Hilde, Brandon, Brian, Prashant, Danielle, Wya, Ina, Joop, Ed, Chris, René, Nisa, Violet and everyone else I forget to mention, thank you for all your help and for all the fun and inspiring conversations during coffee breaks!

Alle medewerkers van de afdeling orthodontie UMCG die mee hebben geholpen aan het mogelijk maken van mijn promotieonderzoek.

Floris Pelser, jouw onderzoek is de basis geweest van mijn (ons) eerste artikel. Bedankt dat je jouw werk aan mij toevertrouwd hebt.

Zachie Fourie, tijdens de eerste 2 jaar van mijn opleiding tot orthodontist mocht ik met jou een kantoor delen. Hiervoor ben ik nog steeds erg dankbaar, jij hebt me in die jaren geïnspireerd om door te gaan met het doen van onderzoek en jij liet me zien wat je met hard werken en een goed hart kunt bereiken!

Krista Janssen, dank je wel voor het luisterd oor dat je me af en toe gaf, ook al had jij het zelf altijd enorm druk. Je bent echt een fijne collega om mee samen te werken! Heel veel succes met jouw eigen promotietraject.

Joerd van der Meer, dank je wel voor de gezelligheid op de afdeling Orthodontie en bij de BDI momentjes! Succes met het afronden van je proefschrift.

Gea van der Bijl, heel erg bedankt voor al je hulp rondom de organisatie van mijn promotie.

Arjen Grotenhuis en Marieke van de Lagemaat, jullie hebben beide met het onderzoek voor jullie masterscriptie een significante bijdrage geleverd aan dit proefschrift, bedankt hiervoor. Marieke, dankzij jouw vele werk als coauteur van hoofdstuk 6 is dit laatste hoofdstuk in recordtempo tot stand gekomen, zonder jou was dat niet gelukt, bedankt!

Natuurlijk wil ik alle proefpersonen bedanken die hebben meegedaan met mijn onderzoek.

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Zij zijn de pijlers waarop dit proefschrift is gebouwd en zonder hen was dit niet tot stand gekomen.

Ik wil de firma’s Ortholab B.V., Ortho=solutions B.V., Dentsply Lomberg en Orthotec B.V. bedanken voor het belangeloos ter beschikking stellen van diverse materialen die zijn gebruikt in dit onderzoek.

Mijn vriendinnetjes Myriam Heerschop-Karsemeijer, Jessica Heerikhuisen, Sonja Tulner, Catherine Volgenant en Ilse de Boer, bedankt voor jullie vriendschap tijdens deze periode waarin ik niet altijd even gezellig was omdat ik te druk was met mijn onderzoek. Ik heb binnenkort hopelijk weer meer tijd om met jullie af te spreken!

Ellis Buining, Betty Vedder en Francien Nijdam-Massier, jullie zijn mijn rotsen in de branding geweest tijdens de periode van mijn promotie! Jullie vriendschap betekent heel veel voor mij en ik weet niet of het zonder jullie had volgehouden in Groningen! Heel erg bedankt voor alles wat jullie voor mij hebben gedaan! Matties for life!

Mijn lieve vriendinnetje en paranimf Monique Vink-Vos, ook al werd de afstand in letterlijke zin tussen ons wat groter toen ik in Groningen ben gaan wonen, zijn we gelukkig altijd heel close gebleven! De afgelopen periode heb je altijd voor me klaargestaan, door dik en dun en dat heeft heel veel voor me betekend. Bedankt dat je mijn vriendinnetje bent!

Leontine, mijn lieve zus en paranimf. Heel erg bedankt dat jij op deze belangrijke dag naast mij wilt staan net zoals ik bij jouw promotie naast jou mocht staan. Voor mij voelt dat als een enorme steun. Je weet dat ik zielsveel van je hou!

Papa, 30 jaar geleden stond jij in de aula van het Academiegebouw om je proefschrift te verdedigen, nu sta ik er. En dan promoveren we ook nog in hetzelfde vakgebied, dat is toch best wel bijzonder! Ik vind het fantastisch om met je samen te mogen werken in onze orthodontistenpraktijk! Je bent mijn hele leven een enorme steunpilaar en inspirator voor mij geweest en ik hoop de komende jaren nog heel veel van je te leren!

Mama, ik wil je bedanken voor alle liefde, steun en aanmoediging die je me mijn hele leven hebt gegeven. Dankzij jou en papa ben ik geworden wie ik ben en daar ben ik jullie enorm dankbaar voor. Je hebt altijd voor mij klaargestaan en dat ik zo ver ben gekomen heb ik daarom voor een groot deel aan jou te danken!

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CURRICULUM VITAE

Marije Albertine Jongsma was born on Februari 3rd, 1986 in Den Helder, the Netherlands. She finished secondary school in 2004 at the “R.S.G. Wiringherlant” in Wieringerwerf. and continued her education by studying dentistry at the Academic Center for Dentistry in Amsterdam (ACTA), a collaboration of the VU University Amsterdam and the University of Amsterdam. She obtained her Bachelor of Science in Dentistry degree in 2007 and her Master of Science in Dentistry degree in 2009. After graduating as a dentist, she started a postgraduate program in orthodontics at the Department of Orthodontics at the University Medical Center Groningen (UMCG) in 2009. In 2011, during her postgraduate training in orthodontics, she became involved in the research of biofilm on orthodontic retention wires and was given an opportunity to start a PhD project on this subject. This project was a collaboration of the Department of Orthodontics at the UMCG and the Department of Biomedical Engineering, part of the W.J. Kolff Institute. In August 2013 she finished the postgraduate program in orthodontics and in September 2013 she started working as an orthodontist at Jongsma & Jongsma Orthodontisten, a private practice in Den Helder, the Netherlands.


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INTRODUCTION

In the 17th century the Dutch fabric merchant Antonie van Leeuwenhoek started to construct his own microscopes in order to be able to better examine the quality of the fabrics he bought and sold. He examined more than just his fabrics and after utilizing one of his own microscopes in 1684 to look at the accumulation of matter on his teeth, he remarked in a report to the Royal Society of London: “The number of these animalcules in the scurf of a man’s teeth are so many that I believe they exceed the number of men in a kingdom”. This was not enough however, to satisfy the curiosity of the fabric merchant, who would become one of the most famous microbiologists of all times, and he furthermore discovered “that the vinegar with which I washt my Teeth, kill’d only those Animals which were on the outside of the scurf, but did not pass thro the whole substance of it”.

Translated to one of the important topics in modern microbiology, Van Leeuwenhoek was referring to the biofilm mode of growth of bacteria adhering on a surface,1 embedding themselves in a matrix of extracellular polymeric substances (EPS)2 that not only offers physical protection against antimicrobial penetration but can also yield bacterial properties that are different from their planktonic counterparts. Bacteria in their adhering, biofilm mode of growth can become inherently resistant to antimicrobials through mutation,3 formation of antibiotic degrading enzymes,4 endogenous oxidative stress,5 phenotypic changes,6 and low metabolic activities.7 Despite extensive studies over many centuries, prevention of biofilm formation remains a prime challenge in many industrial and biomedical applications. In industrial applications, biofilms inflict major damage when formed on processing equipment or in pipes used to transport resources.8 In the biomedical field, biofilm-related infections can develop everywhere in the human body from head (oral biofilms9) to toe (infected diabetic foot ulcers10). Biofilm-related infections are rarely cleared by the host immune system and especially infections that arise after implantation of biomaterial implants (e.g. prosthetic hips and knees) or devices (e.g. pace makers) are known to be persistent and difficult to treat, since the antimicrobial tolerance of bacteria in their biofilm mode of growth extends to many antibiotics used in modern medicine.11 Moreover, dental caries and periodontal diseases, the most wide-spread infectious diseases in the world, are due to biofilms that Van Leeuwenhoek tried to eliminate by using vinegar as an antimicrobial mouthrinse.12

Although the microscopes used nowadays are more sophisticated than the ones Van Leeuwenhoek employed, our understanding of the recalcitrance of biofilms toward antimicrobial penetration is still based on qualitative description of biofilms,13 using expressions as “water channels”, “mushroom structures”, “whiskers” and “streamers”.14,15 This raises the question whether quantifiable properties of biofilms exist that would relate with antimicrobial penetration into a biofilm. As for polymeric materials, structural and

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compositional properties of biofilms, should be reflected in their viscoelastic properties. Viscoelastic properties of oral biofilms depend on the degree of compaction during formation, the absence or presence of flow during growth, their architecture and microbial composition.16,17 The viscoelastic properties of oral biofilms can be determined by evaluating their relaxation after deformation during external loading. Stress relaxation during external loading is a time-dependent process and can be separated into a number of responses, each with a characteristic time-constant.18 Although Maxwell analysis of stress-relaxation to derive the characteristic time-constants of the various relaxation processes that occur in a biofilm under external loading has been done before,19 results have been regarded mainly from a mathematical perspective and the details of the relaxation-structure-composition relation in biofilms and the physical processes associated with the different time-constants, are mostly neglected. Stress relaxation may involve a number of processes, like the outflow of water and EPS from the biofilm and re-arrangement of the bacteria in the biofilm.20 Since penetration of an antimicrobial into a biofilm depends on diffusion21 and therewith on its structural and compositional features, like the presence of water-filled channels in the biofilm or EPS-containing spaces, we here hypothesize that the penetration of an antimicrobial into a biofilm may relate with stress relaxation and its underlying processes.

The aim of this study is to gain evidence in support of this hypothesis. To this end, single-species biofilms of two oral bacterial strains, Streptococcus oralis and Actinomyces naeslundii were grown in a parallel plate flow chamber (PPFC)22 and in a constant depth film fermenter (CDFF).23 Subsequently, we measured their viscoelastic properties using a low load compression tester, as well as the penetration of chlorhexidine into the biofilms. Following Van Leeuwenhoek, we chose to collect support for our hypothesis based on oral biofilms, because the human oral cavity is highly accessible and also allows for sampling of in vivo formed biofilm. Therefore, in order to not only gain in vitro evidence in support of our hypothesis, an intra-oral biofilm collection device was developed to grow oral biofilms in situ, in absence of mechanical perturbation. In vivo formed biofilms in the devices worn by human volunteers were examined ex situ with respect to their viscoelastic properties and chlorhexidine penetration and results and conclusions compared with those obtained for in vitro formed oral biofilms. Chlorhexidine is known to be the most effective oral antimicrobial to date24 and surprisingly, despite its extensive use, inherent bacterial resistance against chlorhexidine has hardly or never been reported as compared to antibiotic resistance of many bacterial pathogens. This makes chlorhexidine an ideal antimicrobial to separate a possible inherent tolerance of biofilm bacteria for the antimicrobial from the physical protection offered by the biofilm mode of growth and study its penetration through a biofilm.

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surfaces in different parts of the dentition, and in vitro on hydroxyapatite. Caries Res 32:447-455 37. Mandel ID (1987) The functions of saliva. J Dent Res 66 Spec No:623-627 38. Aas JA, Paster BJ, Stokes LN, Olsen I, Dewhirst FE (2005) Defining the normal bacterial flora of the oral cavity. J Clin Microbiol 43:5721-5732 39. Reich E, Lussi A, Newbrun E (1999) Caries-risk assessment. Int Dent J 49:15-26

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Bacterial strains and growth conditionsS. oralis J22 and A. naeslundii T14V-J1 grown on blood agar plates, were used to inoculate 10 ml modified Brain Heart Infusion broth (BHI, Oxoid Ltd., Basingstoke, UK) (37.0 g/l BHI, 5.0 g/l yeast extract, 0.4 g/l NaOH, 1.0 g/l hemin, 0.04 g/l vitamin K1, 0.5 g/l L-cysteine, pH 7.3) and were cultured for 24 h at 37°C in ambient air for S. oralis J22 and anaerobically for A. naeslundii T14V-J1. These cultures were used to inoculate 200 ml modified BHI and grown for 16 h. Bacteria were harvested by centrifugation at 870 g, 10°C for 5 min and washed twice in sterile adhesion buffer (50 mM potassium chloride, 2 mM potassium phosphate, 1 mM calcium chloride, pH 6.8). The bacterial pellet was suspended in 10 ml adhesion buffer and sonicated intermittently in an ice-water bath for 3 × 10 s at 30 W (Vibra cell model 375, Sonics and Materials Inc., Newtown, CT, USA) to break bacterial chains and clusters, after which bacteria were resuspended in adhesion buffer. A concentration of 3 × 108 bacteria/ml was used for PPFC experiments, while a concentration of 9 × 108 bacteria/ml was used in CDFF experiments.

Biofilm formation in a PPFC and CDFFBiofilms were grown on glass slides (water contact angle 7 ± 3 degrees) and hydroxyapatite discs (water contact angle 34 ± 8 degrees) in a PPFC and a CDFF, respectively after adsorption of a salivary conditioning film from reconstituted human whole saliva for 14 h at 4°C under static conditions. Reconstituted human whole saliva was obtained from a stock of human whole saliva from at least 20 healthy volunteers of both genders, collected into ice-cooled beakers after stimulation by chewing Parafilm®, 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. Freeze-dried saliva was dissolved in adhesion buffer (1.5 g/l). All volunteers, gave their verbal informed consent to saliva donation according to a fixed written protocol and were registered in order to document the gender, age and health status of the volunteers, in agreement with the guidelines set out by the Medical Ethical Committee at the University Medical Center Groningen, Groningen, The Netherlands (letter 06-02-2009). Written consent was not required since saliva collection was entirely non-invasive, saliva’s were pooled prior to use and the study was not aimed towards measuring properties of the saliva. Rather saliva was used to lay down an adsorbed protein film prior to biofilm formation studies.

For biofilm formation in the PPFC, 200 ml bacterial suspension was circulated at a shear rate of 15 s-1 in a sterilized PPFC till a bacterial surface coverage of 2 × 106 cm-2 was achieved on a saliva-coated glass bottom plate (for details see16). Subsequently, adhesion

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buffer was flowed at the same shear rate of 15 s-1 for 30 min in order to remove non-adhering bacteria from the tubes and flow chamber. Next, growth medium (20% modified BHI and 80% adhesion buffer) was perfused through the system at 37°C for 48 h, also at a shear rate of 15 s-1

Biofilms were grown in a sterile CDFF (for details see23) on saliva coated hydroxyapatite discs by introducing 200 ml bacterial suspension in the fermenter during 1 h, while the table with the sample holders was rotating at 1 rpm. Then, rotation was stopped for 30 min to allow bacteria to adhere before growth medium was introduced and rotation resumed. The biofilm was grown for 96 h at 37°C under continuous supply of a mixture of adhesion buffer and modified BHI at a rate of 80 ml/h. The system was equipped with 15 sample holders and each sample holder contained 5 saliva coated hydroxyapatite discs, recessed to a depth of 100 µm.

Oral biofilm collection in vivoThe intra-oral biofilm collection device (Fig. 1) was made of medical grade stainless steel 316, and is composed of two parts: a base (5×3×2 mm) that is fixed to the center of the buccal surface of the upper first molars and a replaceable cover plate (4×3×0.2 mm). Biofilms formed on the inner side of the replaceable cover plate in the absence of mechanical perturbations, were considered for this study.

Five volunteers (aged 26 to 29 years) were included in this study. Volunteers all had a complete dentition with maximally one restoration, no bleeding upon probing and were not using any medication. Each volunteer was assigned a random number between 1 and 5 used for later data processing. The study was approved according to the guidelines of the Medical Ethics Committee of the University Medical Center Groningen, Groningen, The Netherlands (letter 28-9-2011), including the written informed consent by the volunteers and the tenets of the Declaration of Helsinki.

A base device was fixed to buccal surfaces of the upper first molars of the volunteers (see also Fig. 1) after mild etching of the tooth surface using light cure adhesive paste (Transbond™ XT, 3M Unitek, USA), a procedure similar to the one used for the bonding of orthodontic brackets. Prior to bonding, the base and cover plate of the device were brushed using a rubber cup and cleaner paste (Zircate® Prophy Paste, Densply, Caulk, USA) at low speed (less than 2,500 rpm/min) and autoclaved. Subsequently, the base surface was coated with a thin layer of primer and bonding agent (CLEARFIL SE BOND, Kurary Medical Inc., Japan). The stainless steel cover plate was inserted using a pair of tweezers and kept in place using Light Cure Adhesive Paste (Transbond™ XT, 3M Unitek, USA). Volunteers were asked to wear the device for a total of eight weeks during which

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they were requested to perform manual brushing with a standard fluoridated toothpaste (Prodent Softmint®, Sara Lee Household & Bodycare, Exton, USA) according to their habitual oral hygiene but to refrain from the use of an additional mouthrinse.

The cover plates could be removed with a dental explorer, after which cover plates with biofilm were placed in a moisturized petri dish for transport from the dental clinic to the laboratory. In a separate pilot study, it was established that two weeks of intra-oral biofilm formation in the device yielded biofilm thicknesses that were similar to the ones obtained in vitro. Therewith, in vivo biofilms could be collected four times from each volunteer. After each experiment, cover plates were sanded to remove biofilm and other residuals, prior to autoclaving. After the experiments, the base of the device was removed from the tooth surface with a debracketing plier and residual adhesive was grinded off the tooth surface with a low speed hand piece. A base device was only used once in each volunteer. The tooth surface was polished and cleaned with rubber cup and cleaner paste. No signs of gingival inflammation were observed in any volunteer after removal of the base device.

Low load compression testingThe thickness and stress relaxation of the biofilms were measured with a low load compression

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tester, described before.16 Stress relaxation was monitored after inducing 10, 20, and 50% deformation of the biofilms within 1 s and held constant for 100 s, while monitoring the stress relaxation (see Fig. 2A). Each deformation was induced three times at different locations on the same biofilm.

Stress relaxation as a function of time was analyzed using a generalized Maxwell model containing three elements (see Fig. 2B) according to

(1)

321321)( ttt

ttteEeEeEtE��

++=in which E(t) is the total stress exerted by the biofilm divided by the strain imposed, expressed as the sum of three Maxwell elements with a spring constant Ei, and characteristic decay time, i (see also Fig. 2B). For calculating E (t), deformation was expressed in terms of strain, , according to the large strain model using

(2)

)1ln(hh�

+=e

where h is the decrease in height and h is the un-deformed height of the biofilm. The model fitting for Ei and i values of the three elements was done by minimizing the chi-squared value using the Solver tool in Microsoft Excel 2010. Fitting to three Maxwell elements yielded the lowest chi-squared values and increasing the number of Maxwell elements only yielded minor decreases in chi-squared values of less than 3%. The elements derived were rather arbitrarily named fast, intermediate or slow based on their values, i.e. 1 < 5 s, 5 s < 2 < 100 s and 3 > 100 s, respectively (see also Fig. 2B). Relative importance of each element, based on the value of its spring constant Ei, was expressed as the percentage of its spring constant to the sum of all elements’ spring constants at t = 0.

Penetration of chlorhexidine into biofilmsIn vitro and in vivo formed biofilms were all exposed in vitro to a 0.2 wt% chlorhexidine-containing mouthrinse (Corsodyl®, SmithKline Beecham Consumer Brands B.V., Rijswijk, The Netherlands) for 30 s and subsequently immersed in adhesion buffer for 5 min. After exposure to chlorhexidine, biofilms were stained for 30 min with live/dead stain (BacLight™, Invitrogen, Breda, The Netherlands) and CLSM (Leica TCS-SP2, Leica Microsystems Heidelberg GmbH, Heidelberg, Germany) was used to record a stack of images of the biofilms with a 40× water objective lens. Images were analyzed with Leica confocal software to visualize live and dead bacteria in the biofilms. The ratio of the intensity of red (dead

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bacteria) to green (live bacteria), R/G, was plotted versus the biofilm thickness (see Fig. 3).

The biofilm thickness where the ratio R/G became less than 1.5 was taken as the thickness of the dead band. Next, a penetration ratio was calculated according to

(3) thicknessbiofilm total

thicknessband dead ration Penetratio =

Penetration ratios were calculated for three different, randomly chosen locations on the biofilms and presented as averaged over the different locations.

Figure 3. Chlorhexidine penetration into in vitro and in vivo formed oral biofilms and the calculation of the penetration ratio.I. Representative CLSM-images (cross sectional view) of the penetration of chlorhexidine (0.2 wt%) during 30 s into oral biofilms grown in vitro and in vivo (exposure to chlorhexidine was done in vitro). A: S. oralis J22 biofilm grown under flow in a PPFC. B: S. oralis J22 biofilm grown under compaction in a CDFF. C: A. naeslundii T14V-J1 biofilm grown under flow in a PPFC.D: A. naeslundii T14V-J1 biofilm grown under compaction in a CDFF.E and F: two weeks old, in vivo formed oral biofilm. Scale bar represents 75 μm. II. Red to green intensity ratio (R/G), denoting the ratio of dead to live organisms in a biofilm versus the thickness of the biofilm. a is the dead band thickness and b is the total biofilm thickness. R/G = 1.5 was taken as the cut-off

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Statistical analysisStatistical analysis was performed with SigmaPlot software (version 11.0, systat software, Inc., California, USA). Differences in biofilm thickness and viscoelasticity were evaluated after testing for normal distribution and equal variance of the data. If data failed one of these tests, a Mann-Whitney Rank Sum test was used to determine statistical significance, otherwise a Student t-test was applied. Pearson Product Moment Correlation test was used to disclose relations between the penetration of chlorhexidine into and the relaxation of biofilms.

RESULTS

Biofilms of coccal-shaped S. oralis J22 and rod-shaped A. naeslundii T14V-J1 grown in the PPFC reached a thickness of 131 ± 15 μm and 109 ± 26 μm, respectively (Table 1). The biofilm thickness in the CDFF for S. oralis J22 was 119 ± 6 μm and 125 ± 9 μm for A. naeslundii T14V-J1. There were no significant differences (p > 0.05, Student t-test) in thickness between biofilms grown under flow and in the CDFF. Also differences in biofilms

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thickness across strains were not statistically significant (p > 0.05, Student t-test).

The penetration of chlorhexidine in biofilms grown in the PPFC was significantly different (p < 0.05, Mann-Whitney Rank Sum test) for S. oralis J22 and A. naeslundii T14V-J1, and the penetration ratio amounted 0.33 ± 0.09 and 0.56 ± 0.08, respectively (see also Table 1 and Fig. 3). On the other hand, there were no significant strain-dependent differences in penetration of chlorhexidine into biofilms grown in the CDFF, showing penetration ratios of 0.48 ± 0.04 and 0.39 ± 0.06 in biofilms of S. oralis J22 and A. naeslundii T14V-J1, respectively (p > 0.05, Mann-Whitney Rank Sum test). Interestingly, whereas biofilms offered a clear physical

protection against chlorhexidine, bacteria dispersed from biofilms grown either in the PPFC or in the CDFF were highly susceptible to chlorhexidine (Fig. 4), confirming that the absence of bacterial killing in the deeper layers of the biofilms are not due to changes in inherent properties of the bacteria in their biofilm mode of growth, but solely to difficulties encountered by the antimicrobial in penetrating to the deeper layers. Note that a similar conclusion has been drawn for three days old in vivo grown

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oral biofilms, after dispersal and exposure to chlorhexidine.25

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University of Groningen Biofilm on orthodontic retention ... · appliances and retention wires,...

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