Biodegradation of Resin-Dentin Interfaces Increases Bacterial … · 2010. 11. 3. · ii Biodegradation of Resin-Dentin Interfaces Increases Bacterial Microleakage Masters of Applied

Biodegradation of Resin-Dentin Interfaces

Increases Bacterial Microleakage

By

Sanaz Kermanshahi

A thesis submitted in conformity with the requirements for the degree

of

Masters of Applied Science

Biomaterials Department

Faculty of Dentistry

University of Toronto

Sanaz Kermanshahi, 2009

ii

Biodegradation of Resin-Dentin Interfaces Increases

Bacterial Microleakage

Masters of Applied Science, 2009

Sanaz Kermanshahi

Biomaterials Department, Faculty of Dentistry, University of Toronto

ABSTRACT

Bis-GMA-containing resin-composites undergo biodegradation in human saliva, yielding

Bis-hydroxy-propoxy-phenyl-propane (Bis-HPPP). This may compromise the integrity of

the resin-tooth interfacial interface, contributing to bacterial microleakage. The objective

of this work was to determine whether the biodegradation of resin-dentin restorative

margins and bacterial microleakage are correlated with eachother. Resin-composites

(Scotchbond, Z250, 3M) bonded to human dentin were incubated in either buffer, or

dual-esterase media (pseudocholinesterase/cholesterol-esterase) with activity levels

matching that of human saliva, for up to 90 days. Incubation solutions were analyzed for

resin degradation by-products using high-performance liquid-chromatography. Post-

incubation, specimens were suspended in a chemostat-based biofilm fermentor

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cultivating Streptococcus mutans NG8 for 7 days. Bacterial microleakage was assessed

through confocal laser scanning microscopy. Bis-HPPP production, as well as depth and

volume of bacterial cell penetration within the interface were higher at 30 and 90 days

PCE-CE incubation vs. buffer incubation (p<0.05). A high correlation (R2=0.97) was

found between Bis-HPPP and cumulative interfacial bacterial count. An overall decline in

interfacial integrity was observed following exposure to human saliva-like esterases over

time.

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TABLE OF CONTENTS

CHAPTER 1: INTRODUCTION ................................................................................... 1 1.1 HYPOTHESIS .............................................................................................................. 3

1.2 OBJECTIVES ............................................................................................................... 5

1.3 REFERENCES ............................................................................................................. 6

CHAPTER 2: LITERATURE REVIEW .................................................................... 12 2.0 THE PROBLEM: DENTIN BONDING .................................................................... 12

2.1 ENZYME-INDUCED BIODEGRADATION OF THE RESIN-DENTIN

INTERFACE ............................................................................................................. 15

2.2 MICROLEAKAGE..................................................................................................... 18

2.3 ORAL PLAQUE ......................................................................................................... 19

2.3.1 Current Conceptual Biofilm Model ................................................................. 20

2.3.1.1 Adherence and Colonization ......................................................................... 21

2.3.2 Streptococcus Mutans ...................................................................................... 23

2.3.2.1 Interaction with Resin-based Materials ........................................................ 24

2.3.3 Artificial Models Systems Used to Culture Oral Plaque In Vitro ................... 25

2.4 MICROSCOPY TECHNIQUES USED TO IMAGE THE RESIN-DENTIN

MARGINAL INTERFACE .............................................................................................. 27

2.5 FIGURES ............................................................................................................. 29

2.6 REFERENCES ........................................................................................................... 30

CHAPTER 3: Biodegradation of Resin-Dentin Interfaces Increases Bacterial

Microleakge ............................................................................................................. 53 3.1 INTRODUCTION ...................................................................................................... 53

3.2 MATERIALS AND METHODS ................................................................................ 54

3.2.1 Preparation of Resin-Dentin Specimens .................................................................. 54

3.2.2 Degradation Media Incubation of Resin-Dentin Specimens ........................... 54

3.2.3 Bis-HPPP Byproduct Isolation ........................................................................ 55

3.2.4 Incubation of Resin-Dentin Specimens in Chemostat-Based Biofilm Fermentor

(CBBF) ...................................................................................................................... 55

3.2.5 Confocal Laser Scanning Microscopy (CLSM) Analysis ............................... 56

3.2.6 Statistical Analysis ........................................................................................... 56

3.3 RESULTS ............................................................................................................. 57

3.3.1 Biodegradation ................................................................................................. 57

3.3.2 Bacterial Microleakage .................................................................................... 57

3.4 DISCUSSION ............................................................................................................. 60

3.4.1 Biodegradation ........................................................................................................ 60

3.4.2 Bacterial Microleakage ........................................................................................... 61

3.5 CONCLUSION ........................................................................................................... 63

3.6 ACKNOWLEGEMENTS ........................................................................................... 64

3.7 FIGURES ............................................................................................................. 65

3.7 FIGURE CAPTIONS ................................................................................................. 69

3.8 REFERENCES ........................................................................................................... 70

CHAPTER 4: GENERAL DISCUSSION .................................................................... 74 4.1 DISCUSSION RE: HYPOTHESIS #1 ....................................................................... 74

4.2 DISCUSSION RE: HYPOTHESIS #2 ....................................................................... 75

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4.3 DISCUSSION RE: HYPOTHESIS #3 ....................................................................... 75

4.4 REFERENCES ........................................................................................................... 78

CHAPTER 5 – CONCLUSIONS ................................................................................... 83

CHAPTER 6 – RECOMMENDATIONS ..................................................................... 84 6.1 REFERENCES ........................................................................................................... 88

APPENDIX A – RECIPES ............................................................................................. 90

APPENDIX B – GAMMA IRRADIATION ................................................................. 91

APPENDIX C - RESIN-DENTIN SAMPLE PREPARATION.................................. 95

APPENDIX D – STERILITY ASSAYS OF SPECIMEN PREPARATION

PROCEDURE ........................................................................................................... 99

APPENDIX E – SALIVARY-LIKE ESTERASES .................................................... 102

APPENDIX F – PCE-CE SOLUTION ....................................................................... 109

APPENDIX G – HALF-LIFE EXPERIMENTS OF PCE-CE SOLUTION ......... 112

APPENDIX H – COMPARATIVE BIODEGRADATION OF HSDE AND PCE-

CE SOLUTION ON COMPOSITE RESIN SPECIMENS ................................. 113

REFERENCES ........................................................................................................ 115

APPENDIX I – HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY

(HPLC) ........................................................................................................... 116

REFERENCES ........................................................................................................ 118

APPENDIX J – INCUBATION MEDIA FREEZE DRYING PROCEDURE....... 119

APPENDIX K – MICROBIOLOGY TECHNIQUES ............................................... 121

APPENDIX L – CHEMOSTAT-BASED BIOFILM FERMENTOR SET-UP....... 123

APPENDIX M – LIVE/DEAD BACLIGHT BACTERIAL VIABILITY

FLOURESCENT STAINING ................................................................................ 126

APPENDIX N – CONFOCAL SCANNING LASER MICROSCOPY .................... 129

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ABBREVIATIONS

ATR Acid tolerance response

Bis-GMA 2.2-Bis[4-(2-hydroxy-3-methacryloyoxy-propoxy)phenyl]propane

Bis-HPPP Bis-hydroxypropoxyphenyl propane

CBBF Chemostat-based Biofilm Fermentor

CDFF Constant depth film fermentor

CE Cholesterol esterase

CLSM Confocal laser scanning microscopy

EPS Extra-cellular polymeric substances

GTF Glycosyltransferase

HPLC High performance liquid chromatography

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MA Methacrylic acid

MMP Matrix metalloproteinase

MS Mass Spectometry

PBS Phosphate Buffer Solution

PCE Psuedo-choline esterase

ROI Region of interest

SEM Scanning electron microscopy

TEG Tri-ethylene Glycol

TEGDMA Tri-ethylene glycol di-methacrylate

TEM Transmission electron microscopy

THYE Todd Hewitt Yeast Extract

UV Ultra Violet

1

CHAPTER 1: INTRODUCTION

The evaluation of dental composite resin restorations over the past 20 years has centered

primarily on issues of biomechanics (1-4), as these parameters are most immediate in

determining short-term clinical restoration success. However in the midst, other important

issues concerning the biocompatibility of composite resins - though at times briefly

acknowledged - have gone otherwise largely un-addressed (5,6).

As can be expected of any synthetic material placed within a biological system, composite

resins are not completely inert (7-9). They interact dynamically with the host, forces and

conditions present in the oral environment (1,8,10). By the early 1990‟s, investigations into

the durability of resin-based restorative materials in vivo were reporting material loss at a

faster rate than could attributed purely to mechanical forces (11,12). Others had found

material discoloration at the tooth-composite marginal interface (13,14), a clear marker of

degradation at the adhesive-dentin interface over time (14,15). Release of toxic products

from composite restorations, such as methacrylic acid (MA) and formaldehyde (9,11,16),

and other leachable byproducts (17,18) were also being detected. These early findings

attested to the fact that resin-based dental materials are indeed subject to chemical

degradation in the oral cavity (7,19).

Polymers of resin-based composites are bound by unprotected ester linkages inherently

prone to cleavage by water (7,19,20,21). Since the composition of human saliva is nearly

99% water, these materials are highly susceptible to hydrolytic degradation in vivo. More

importantly, saliva also contains esterase-like enzyme activities capable of accelerating the

hydrolytic process and thus, the rate of chemical degradation (6,9,20,22,23). Many

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investigations confirm the degradative effects of solvolysis on resin materials

(6,9,18,22,24), however little is still known about the full potential for such degradative

interactions, particularly at the tooth-resin interface (5,6).

The fact is that much of the current knowledge base gained from biodegradation studies

conducted on composite resin materials stems from observations made at bulk occlusal

surfaces interfacing the oral cavity (6,17,18). While determining the capacity for chemical

degradation at the composite restoration-storage media interface does corroborate the

susceptibility of resin materials to chemical degradation in vivo, this in itself has not be

shown to possess any direct clinical implications (25,26). Ultimately, it is the physical and

chemical integrity of a restoration‟s adhesive bond layer - the biological interface that

exists between the restoration and the underlying tooth substrate – that determines the

clinical success of any dental restoration (25-30). This is of particular concern in clinical

scenarios where composite restorations must come in contact with dentin, a wet porous

tooth substrate existing below the protective layer of dental enamel (31). Studies

consistently note the difficulty with which resin materials adhere to dentinal margins, as

compared to enamel (25-30).

The potential for chemical interactions with biologically active factors present in vivo -

such oral bacteria (27,28) – also play a vital role in the integrity of a resin-dentin restorative

margin. The degradation of susceptible resin components within the adhesive layer, in

addition to resin leaching (32,33) result in enlarged marginal voids (34,35) - often found on

the scale of several micrometers wide in vivo (31) – which allow for bacterial microleakage

to take place (28,36,37). The release of metabolic byproducts by oral bacteria, once inside

3

the compromised resin-dentin interface, is capable of further propagating interfacial

chemical degradation (20). Both saliva and the demineralized dentinal substrate harbour

intrinsic proteolytic factors (38-43) capable of being activated upon marginal infiltration by

oral bacteria.

No commercial dentin bonding system to date is capable of achieving a completely

hermetic seal, consistently across the restorative margin (31,44,45). Bacterial

microleakage is the most frequently identified postoperative complication (25), with the

resultant secondary caries being cited as the principle cause of failure in Class I and II

direct composite restorations (46,47). The aim of the current investigation is to utilize a

salivary esterase-like media solution, as well as an artificial oral cavity model system -

modified from Li et al (2001) – in order to mimic relevant solvolytic-like and microbial

events (48) occurring along the restoration-tooth interface of composite restorations placed

in vivo.

1.1 HYPOTHESIS

Salivary esterase-like biodegradation of resin materials– determined through the detected

release of a known resin matrix degradation byproduct – corresponds to the marginal

integrity of a composite resin restoration bonded to a dentinal substrate, shown through

bacterial penetration of its margin.

Specifically, it is hypothesized that:

1) Human saliva-like esterase activities will degrade the polymerized matrices of both

the composite resin and resin adhesive materials of the restoration in a reproducible

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manner, and at a rate comparable to that which occurs in vivo with human saliva.

Challenge: The main esterase-like activities of human saliva have been identified

and it is possible to reproduce them in vitro. However the enzyme stability of these

esterases decrease significantly in the presence of a substrate when unaccompanied

by other proteins contained within human saliva.

2) S. mutans biofilm will adhere to and penetrate all voids/ gaps occurring at the

interfacial margin of a resin-dentin restoration suspended within an in vitro oral

model system, in a reproducible fashion. Challenge: Dental plaque occurring in

vivo is by definition a biofilm, whereas bacterial cell cultures grown in vitro often

occur in the planktonic phase. The differential transcription of genes depending on

mode of growth, mean that phenotypic characteristics relating to cellular adherence

and mechanisms of survival are altered drastically between the biofilm and

planktonic phases.

3) As the length of time and exposure of a resin-dentin restorative margin to human

saliva esterase-like activity increases, the total bacterial cell count and overall depth

of Streptococcus mutans penetration within the restorative margin will also rise.

Challenge: Visualizing the intact biofilm and any potential migration it may have

within the resin-dentin restorative margin may require a number of drastic specimen

processing steps, post-incubation, that may result in morphological distortions

within the specimen.

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1.2 OBJECTIVES

The following objectives will be pursued to address the hypothesis:

1) To induce chemical degradation of the resin-dentin interfacial margin using an in

vitro human saliva system on human molar teeth restored using clinical practice

methods.

2) To generate and sustain monoclonal biofilm of Streptococcus mutans over the

degraded resin-dentin specimen, suspended within a model artificial saliva system.

3) To qualitatively describe and quantify the level of bacterial microleakage taking

place at regions of interest (ROI) along the resin-dentin interfacial margin through

vital fluorescent staining combined with confocal scanning laser microscopy

(CLSM) and ImageJ software analysis.

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1.3 REFERENCES

1. van Noort R. Introduction to Dental Materials. Papel, Spain: Times Mirror

International Publishers Limited; 1994. pgs. 3-145.

2. El-Mowafy OM, Lewis DW, Benmergui C, Levinton C. (1994) Meta-analysis on

long-term clinical performance of posterior composite restorations. J. Dent. 22:33-

43.

3. Taylor DF, Bayne SC, Leinfelder KF, Davis S, Koch GG. (1994) Pooling of long

term clinical wear data for posterior composites. Am J Dent 7:167-174.

4. Turssi CP, De Moraes Purquerio B, Serra MC. (2003) Wear of dental resin

composites: insights into underlying processes and assessment methods – a review.

J Biomed Mater Res B Appl Biomater 15:65(2):280-285.

5. Finer Y, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and

psuedocholinesterase on the biodegradation of dental composites. Biomat 25:1787-

1793.

6. Lin BA, Jaffer F, Duff MD, Tang YW, Santerre JP. (2005) Identifying enzyme

activities within human saliva which are relevant to dental resin composite

biodegradation. Biomat 26:4259-4264.

7. Gopferich A. (1996) Mechanisms of polymer degradation and erosion. Biomat

17:103-114.

8. Schedle A, Franz A, Rausch-Fan X, Spittler A, Lucas T, Samorapoompichit P,

Sperr W, Boltz-Nitulescu G. Cytotoxic effects of dental composites, adhesive

substances, compomers and cements. Dent Mater 14:429-440.

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9. Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental

composites by cholesterol esterase. J Dent Res 78(8):1459-1468.

10. Wataha JC. (2001) Principles of biocompatibility for dental practitioners. J Prosthet

Dent 86:203-209.

11. Freund M, Munksgarrd EC. (1990) Enzymatic degradation of

BISGMA/TEGDMA-polymers causing decreased microhardness and greater wear

in vitro. Scand J Dent 4: 351-5.

12. Munksgaard EC, Freund M. (1990) Enzymatic hydrolysis of (di)methacrylates and

their polymers. Scand J Dent Res 98:351-355.

13. Asmussen E, Munksgaard EC. (1988) Bonding of restorative resins to dentine:

Status of dentine adhesives and impact on cavity design and filling techniques. Int

Dent J 38: 97-104.

14. Prati C, Nucci C, Davidson CL, Montanari G. (1990) Early marginal leakage and

shear bond strength of adhesive restorative systems. Dent Mater 6:195-200.

15. Lee SY, Greener EH, Mueller HJ. (1995) Effect of food and oral simulating fluids

on structure of adhesive composite systems. J Dent 23:1:27-35.

16. Oysaed H, Sjovik-Kleven IJ. (1988) Release of formaldehyde from dental

composites J Dent Res 67:1289-1294.

17. Bean TA, Zhuang WC, Tong PY, Eick JD, Yourtee DM. (1994) Effect of esterase

on methacrylates and methacrylate polymers in an enzyme simulator for

biodurability and biocompatibility testing. J Biomed Mat Res 28:59-63.

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18. Shajii L, Santerre JP. (1999) Effect of filler content on the profile of released

biodegradation products in micro-filled bis-GMA/TEGDMA dental composite

resins. Biomat 20:1897-1908.

19. Coury AJ, Levy RJ, McMillin CR, Pathak Y, Ratner BD, Schoen FJ, Williams DF,

Williams RL. Degradation of Materials in the Biological Environment. In:

Biomaterials Science: An Introduction to Materials in Medicine. Edited by: Ratner

BD, Hoffman AS, Schoen FJ, Lemons JE. San diego, California: Academic Press

Inc; 1996. pg. 243-281

20. Santerre JP, Shajii L, Leung BW. (2001) Relation of dental composite formulations

to their degradation and the release of hydrolyzed polymeric-resin-derived

products. Crit Rev Oral Biol Med 12(2):136-151.

21. Finer Y, Santerre JP. (2004) The influence of resin chemistry on a dental

composite‟s biodegradation. J Biomed Mater Res 69A:233-246.

22. Jaffer F, Finer Y, Santerre JP. (2002) Interactions between resin monomers and

commercial composite resins with human saliva derived esterases. Biomat 23:1707-

1719.

23. Finer Y, Santerre JP. (2004) Salivary esterase activity and its association with the

biodegradation of dental composites. J Dent Res 83:1:22-26.

24. Yourtee DM, Smith RE, Russo KA, Burmaster S, Cannon JM, Eick JD, Kostoryz

EL. (2001) The stability of methacrylate biomaterials when enzyme challenged:

Kinetic and systematic evaluations. J Biomed Mater Res 57:522-531.

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25. Maupome G, Sheiham A. (1998) Criteria for restoration replacement and

restoration life-span estimates in an educational environment. J of Oral Rehab

25:896-901.

26. Van Meerbeek B, Perdigao J, Lambrechts P, Vanherle G. (1998) The clinical

performance of adhesives. J Dent 26:1-20.

27. Santini A, Mitchell S. (1998) Microleakage of composite restorations bonded with

three new dentin bonding agents. J of Esthet Dent 10:6:296-304.

28. Murray PE, Hafez AA, Smith AJ, Cox CF. (2002). Bacterial microleakage and pulp

inflammation associated with various restorative materials. Dent Mat 18:470-478.

29. Gerdolle DA, Mortier E, Loos-Ayav C, Jacquot B, Panighi MM. (2005) In vitro

evaluation of microleakage of indirect composite inlays cemented with four luting

agents. J Prosthet Dent 93:563-570.

30. Donmez N, Belli S, Pashley DH, Tay FR. (2005) Ultrastructural correlates of in

vivo/in vitro bond degradation in self-etch adhesives. J Dent Res 84:4:355-359.

31. Bouillaguet S. (2004) Biological risks of resin-based materials to the dentin-pulp

complex. Crit Rev Oral Biol Med 15(1):47-60.

32. Tay FR, Pashley DH. (2003) Water treeing – a potential mechanism for degradation

of dentin adhesives. Am J Dent 13:6-12.

33. Chersoni S, Suppa P, Breschi L, Ferrari M, Tay FR, Pahley DH, Prati C. (2004)

Water movement in the hybrid layer after different dentin treatments. Dent Mater

20:796-803

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34. Sano H, Yoshiyama M, Ebisu S, Burrow MF, Takatsu T, Ciucchi D, Carvalho R,

Pashley DH. (1995) Comparative SEM and TEM observations of nanoleakage

within the hybrid layer. Oper Dent 20:160-167.

35. Sano H, Yoshikawa T, Periera PNR, Kanemura N, Morigami M, Tagami J, Pashley

DH. (1999) Long-term durability of dentin bonds made with a self-etching primer,

in vivo. J Dent Res 78(4):906-911.

36. Matharu S, Spratt DA, Pratten J, Ng, YL, Mordan N, Wilson M, Gulabivala K.

(2001) A new in vitro model for the study of microbial microleakage around dental

restorations : a preliminary qualitative evaluation. Inter Endo J 34 :547-553.

37. Zivkovic S, Bojovic S, Pavlica D. (2001) Bacterial penetration of restored cavities.

Oral Surg Oral Med Oral Path Oral Radiol Endod 91:353-358.

38. Pashley DH, Tay FR, Yiu C, Hashimoto M, Breschi L, Carvalho RM. (2004)

Collagen degradation by host-derived enzymes during aging. J Dent Res 83: 216-

221.

39. Arola D, Reprogel RK. (2005) Effects of aging on the mechanical behavior of

human dentin. Biomaterials 26:4051-4061.

40. Mazzoni A, Pashley DH, Tay FR, Gobbi P, Orsini G, Ruggeri A, Carrilho M,

Tjaderhane L, Di Lenarda R, Breschi L. (2008) Immunohistochemical identification

of MMP-2 and MMP-9 in human dentin: Correlative FEI-SEM/TEM analysis. J

Biomed Mater Res, March 11, 2008.

41. Sulkala M, Tervahartiala T, Sorsa T, Larmas M, Salo T, Tjaderhane L. (2007)

Matrix metalloproteinase-8 (MMP-*) is the major collagenase in human dentin.

Arch Oral Biol 52:121-127.

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42. Boushell LW, Kaku M, Mochida Y, Bagnell R, Yamauchi M. (2008)

Immunohistochemical localization of matrixmetalloproteinase-2 in human coronal

dentin. Arch Oral Biol 53: 109-116.

43. Asmussen E, Hansen EK. (1993) Dentine bonding systems. In: State of the Art on

Direct Posterior Filling Materials and Dentine Bonding. Vanherle G, Degrage M,

Willems G, editors. Proceedings of the International Symposuim. Euro Disney,

Paris. pgs. 33-47.

44. Manhart J, Chen HY, Mehl A, Weber K, Hickel R. (2001) Marginal quality and

microleakage of adhesive Class V restorations. J Dent 29 :123-130.

45. Ateyah NZ, Elhejazi AA. (2004) Shear Bond Strengths and Microleakage of Four

types of Dentin Adhesive Materials. J Contemp Dent Pract 5:1:63-73.

46. Mjor IA, Toffenetti F. (2000) Secondary caries: a literature review with case

reports. Quintessence Int 31:165-179.

47. Hickel R, Manhart J. (2001) Longevity of restorations in posterior teeths and

reasons for failure. J Adhes Dent 3(1):45-64.

48. Li YH, Lau PCY, Lee JH, Ellen RP, Cvitkovich DG. (2001) Natural genetic

transformation of Streptococcus mutans growing in biofilms. J Bacteriol 183:897-

908

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CHAPTER 2: LITERATURE REVIEW

2.0 THE PROBLEM: DENTIN BONDING

The clinical longevity of any composite resin restoration has been shown to be primarily

dependent on the structural integrity of its restorative margins (1-6). The clinical

applicability of composite resin restorations is complicated by the fact that these

materials are inherently hydrophobic (7-12). In most clinical cases requiring dental

restoration, restorative materials must come in direct contact with dentin in addition to

enamel (5, 13).

Dentin is approximately 70% by weight mineralized tissue in the form of hydroxyapatite

crystals, 20% collagen, and 10% water (14). The surface of cut dentin is hydrophilic

because fluid is constantly being released through capillary action from the exposed ends

of dentinal tubules. Tubules transverse the tissue and range from about 1.0 to 2.5 m in

diameter, depending on where in the tissue they occur (15, 16). Dentin‟s collagen

component contains an inherent network of mostly type I collagen fibrils that form a

matrix around the tissue‟s mineralized hydroxyapapite component (17); other collagen

types (III, V, and VI) and non-collagenous proteins and proteoglycans are also present as

minor components.

During the dentin bonding process, an acid conditioner is first applied to cut dentin to

superficially dissolve away the hydroxyapapite‟s calcium-phosphorous component. In

doing so, intertubular and peritubular dentinal zones become demineralized, leaving the

cut tubules and collagen fibril network bare and un-reinforced, often to a depth of 5-10

m (13). Access to exposed collagen fibers and dentinal tubules is essential in providing

13

the sites of adhesion required for the formation of micro-mechanical attachments (18).

Hydrophobic resin monomers such as 2.2-Bis[4-(2-hydroxy-3-methacryloyloxy-

propoxy)phenyl]propane (Bis-GMA), and, to a lesser extent, triethylene glycol

dimethacrylate (TEGDMA), are incapable of competing with water for access to these

potential adhesion sites (13-14).

Instead, a priming agent containing an amphiphilic resin monomer - commonly 2-

hydroxyethyl methacrylate (HEMA) dissolved in a volatile solvent of either ethanol or

acetone - is applied (13-14). As a function of its polar hydroxyl group, HEMA is capable

of interacting with water molecules and competing for access to adhesion sites within the

micro-porosities of dentin‟s collagenous phase (13-14, 19-24). The hydrophobic moiety

of amphipillic monomers remains un-bound until another low-viscosity methacrylate-

based bonding or adhesive agent is applied on top (12-13, 20).

Monomers of the adhesive agent (commonly Bis-GMA, TEGDMA and HEMA) co-

polymerize along the methacrylate groups of priming agent monomers (12, 19, 25).

Resin entanglement of conditioned dentin creates a transitional zone between the two

distinctly separate substrates – resin and dentin; by definition, this is known as a hybrid

layer (14, 18-20, 22-23, and 26-28). The formation of a packed, resin-dense hybrid layer

is the basic mechanism by which all classical multi-step adhesive systems achieve micro-

mechanical adhesion between composite resin restorative materials and the dentinal tooth

substrate (29).

Under optimal bonding conditions, resin penetration of cut tubules and associated

branches results in the formation of „resin tags‟ (24, 26, 27) that provide mechanical

14

interlocks, fusing the hybrid layer to the un-etched dentinal substrate below (30).

Adhesive monomers must also infiltrate all micro-porosities created following the acid-

etch removal of dentin‟s mineral phase and fully encapsulate the entire length of

superficially exposed collagen fibrils - devoid of any gaps or voids - down to the surface

of the mineralized layer (22-23, 31). In reality though, an overwhelming body of

literature reveals that this often not the case; the majority of current dentin bonding

agents, including 3M‟s Scotch Bond Multipurpose (3M ESPE, London, Ontario), are

incapable of generating a well-packed, consistently gap-free interfacial margin with

composite resin restorations (12, 23, 32 -47).

This is particularly the case with multi-step adhesive systems, where the acid etchant and

priming resins are applied as two separate steps. The depth of dentin demineralization is

often found to exceed that of resin infiltration, leaving a poorly enforced zone of collagen

fibrils at the bottom of the hybrid layer (32, 35-38, 40, 43, 48). A suggested cause has

been the potential failure of the amphiphillic resin monomers in completely removing all

water molecules near the demineralized-mineralized dentin junction (43, 49-51).

Incompletely resin-infiltrated demineralized zones have also been identified within the

hybrid layer itself (35-36, 41, 50).

More significantly, no dentin adhesive system to date has been able to totally prevent fluid

induction from surrounding media into the hybrid layer (46-47). Hydrophilic interaction

through the HEMA monomer‟s polar moiety greatly enhances water uptake potential (6,

22, 52- 54; 56- 58). SEM analyses confirm the existence of networks of nanometer-sized

water-filled voids intrinsic to all resin-dentin bonded interfaces (43, 46-47; 49). This

15

porosity has been well-documented throughout numerous nanoleakage studies (22, 46-47,

49-50, 55, 59-60), demonstrating the susceptibility of resin-dentin restorative interfaces to

hydrolytic degradation.

Condensation-type bonds within all polymeric resin matrices become severed through a

single-step reaction with water in a process known as hydrolysis (61-62). Long-term

immersion studies show a significant reduction in interfacial bond strength of resin

bonded composite restorations incubated in water (6, 22, 37, 49, 51, 53-55, 63).

Interfibrilar resin loss and exposure of collagen fibrils have also been reported (37, 46,

49, 64). Water immersion studies are the most commonly used in vitro method of

simulating long-term degradation at the adhesive interface (65). However, assessing the

degradative effects of water alone on the resin-dentin interface provides, at best, a

baseline of the potential for degradation present in the oral cavity (49, 66).

2.1 ENZYME-INDUCED BIODEGRADATION OF THE RESIN-DENTIN

INTERFACE

It has been nearly 20 years since Munksgaard and Freund first reported the significant

increase in rate of di- and mono-methacrylate hydrolytic breakdown through the

enzymatic activity of esterase derivatives isolated from human saliva (67-68). Nowadays

this is a well-established fact; esterase-like activities at levels identified in human saliva

(69- 72) catalyze the hydrolytic cleavage of unprotected esterase moieties within the resin

matrices of methacrylate-based resin materials (58, 62, 69-70, 73-77). High performance

liquid chromatography (HPLC), combined with mass spectra (MS) analysis of

16

degradation byproducts collected from incubation media post-digestion, confirm

hydrolysis at the ester bonds of Bis-GMA, TEGDMA and HEMA monomers (72, 75).

The breakdown of the parental methacrylate unit releases specific degradation byproducts

(75) which have been identified in vivo at concentrations of approximately 50 M (69).

The shared end-product for nearly all hydrolyzed un-reacted and partially reacted di- and

mono-methacrylate monomers is methacrylic acid (MA) (71). The diluent resin

monomer TEGDMA is hydrolytically cleaved in a sequential manner to release MA as

well as triethylene glycol (TEG) (75-76). The most direct marker indicating chemical

degradation of a resin material‟s polymerized matrix though is the Bis-GMA derived

bishydroxypropoxyphenylpropane (BisHPPP) (75; 69; 71, 73, 78) – Figure 2.1. Unlike

byproducts generated from residual methacrylates capable of being leached (78), the

majority of detected Bis-HPPP originates from the hydrolysis of ester bonds within the

polymerized Bis-GMA components of the resin matrix itself (71, 76). Bis-HPPP contains

no other hydrolysable ester bonds and is therefore a stable, single-source end-product that

accumulates in incubation media without under-going any further hydrolytic reaction

over time (69).

The majority of studies assessing chemical breakdown of the resin matrix though have

only been able to quantify overall byproduct release from the bulk composite resin

restoration as a whole (58; 69; 72, 75-78). Since hydrolysis of HEMA generates the MA

byproduct common to all methacrylate monomers (58), it has not been possible to isolate

a degradation byproduct specifically generated from resin components of the marginal

interface. Bis-GMA is however present a major constituent of the hybrid layer‟s

17

adhesive resin component. Quantifying the cumulative amount of Bis-HPPP release from

a composite resin-restored tooth sample, overall, is currently the best available method of

attaining some measure of implied chemical degradation occurring at the resin-dentin

marginal interface.

The fact that the restorative interface is characterized by high regional porosity and

continual fluid sorption implies that salivary esterases could potentially gain access to a

greater number of susceptible polymeric moieties in the margin, more frequently, than the

corresponding composite restoration (62). The process of interfacial biodegradation is a

self-propagating cycle; continual solvolytic breakdown of interfacial nano-sized voids,

coupled with the leaching of released degradation byproducts (54, 22), result in the

progressive expansion of voids (50-51; 56). Marginal voids have the potential to span

several micrometers wide (12, 41), allowing for the elution of larger oligomers and

unbound residual monomers that then allow access of fluids to newly exposed sites of ester

linkages within the remaining polymerized resin matrix. Once enough of the interfacial

margin‟s resin component has been eluted, the exposed demineralized dentin beneath the

restoration is no longer protected from biologically active components present in the oral

cavity (37, 51).

Human saliva (79; 80), as well as the dentinal substrate itself (43, 81), harbor a family of

proteolytic enzymes known as matrix metallo-proteinases (MMPs) capable of degrading

exposed collagen fibrils within the hybrid layer (43, 79, 82-86). Once activated, these

peptidases are responsible for the intrinsic auto-degenerative process of dentinal

18

degradation (43, 83-86) and act in concert with host-derived enzymes in breaking down

components of the interfacial margin (65). Overall, incomplete resin infiltration of the

hybrid layer, nanoleakage, and fluid sorption carrying both host-derived salivary

esterases and MMPs are interrelated factors that compromise the integrity of the resin-

dentin restorative margin.

While stress parameters such as shear bond strength (13, 42, 87-88), microtensile bond

strength (6, 24, 29, 89-90), and fracture toughness (24, 40) have been most widely used to

evaluate the structural integrity of a restoration‟s interfacial margins, it should be noted

that these only assess a margin‟s mechanical properties and do not provide direct data on

interfacial porosity. The most immediate measure of interfacial porosity is marginal

leakage (3, 5, 6, 22, 39, 44, 50, 55-56). Several studies show that the degree of

microleakage at a resin-dentin interface (its relative porosity) and associated bond

strengths are poorly correlated (3, 42, 91). Transmission electron microscopy (TEM)

results correlate microleakage with zones of unprotected demineralized dentin within the

hybrid layer (92).

2.2 MICROLEAKAGE

Whereas nanoleakage represents an inherent penetration pathway within the hybrid layer

on a nano-scale (36), what is defined as „true‟ microleakage depends on the presence of a

pre-existing marginal gap (60). Interfacial microleakage takes place where a lack of resin

impregnation of dentin (37) is large enough to allow for the penetration of oral bacteria

(often with a cell span of 0.5-1.0 m in diameter) and/or similarly sized debris (4, 10, 40,

94). Commonly though, it has been traceable organic dyes - such as 0.5%-2% methylene

19

blue (5, 44, 59, 95) – are used to assess interfacial leakage in vitro (3, 5, 39, 44, 59, 87,

92-93). Dye particles are easily detectable, inexpensive and non-toxic; therefore easy to

use. However, study results are often inconsistent and problematic; cited issues range

from inter-study comparability (93) to the overall clinical applicability of such small

penetrating particles spanning a mere 0.12 m in diameter (96).

Microleakage causes decay beneath composite restorations only when existing marginal

gaps are large enough to allow for penetration by bacterial cells (1, 4, 30, 44, 45). As

such, a more clinically relevant method for assessing the microleakage potential of the

bonding interface is to trace for the presence or absence of bacterial cells within the

restorative interface (93-94, 97). Results from scanning electron microscopy (SEM) and

TEM studies have demonstrated bacterial microleakage in vitro (3, 50, 92-93).

Commercially available dentin bonding systems are incapable of providing a completely

hermetic seal of resin restorative margins to dentin (45), and given the predominance of

bacterial activity within the oral cavity, bacterial microleakage is still considered the

most common postoperative complication leading to the ultimate failure of the restoration

(98-99).

2.3 ORAL PLAQUE

Oral plaque collectively refers to the complex matrices of microbial aggregates (micro-

colonies) that naturally exist within the oral cavity (100). The oral cavity is a uniquely

ideal environment for the growth of large, diverse micro flora, owing to the presence of

organic nutrients (host-ingested food), moisture, warm temperature, and the availability

of a diverse range of substrata (tongue, cheek, teeth, and gingival tissues) (101).

20

However, local conditions within dental plaque are subject to considerable regional and

temporal fluctuation (102), which is why oral microbiota must be versatile in order to

proliferate. Under in vivo conditions, bacteria are found preferentially existing within

larger communities - known as biofilms (103). This is particularly true for the cariogenic

agent Streptococcus mutans, which has evolved to become almost completely dependent

on a biofilm-based lifestyle in the oral cavity (103).

Through molecular means of intra and inter-species communication, oral biofilms

function as highly coordinated communities (103-105) co-existing symbiotically (101;

104; 105-111). Most prominent are the S. mutans and sanguis bacterial strains associated

with caries, and actinomycetes associated with periodontal diseases such as gingivitis.

Lactobacilli, staphylococci, and corynebacteria, along with a number of other anaerobes

are also identified. Overall, current estimates predict that approximately 500 species of

microorganisms exists as plaque within the human oral cavity, many which have yet to be

cultivated in vitro (111-113).

2.3.1 Current Conceptual Biofilm Model

A biofilm is depicted as a spatially and temporally heterogeneous three-dimensional

microbial community irreversibly attached to a substratum at a solid-fluid interface (114).

Biofilms comprise of bacterial microcolonies imbedded - and more or less immobilized -

within a hydrated matrix of visco-elastic extracellular polymers known as extracellular

polymeric substances (EPS) (114-117). EPS consist of a mixture of alginates (linear

polysaccharides), proteins, and DNA secreted by the bacterial aggregates themselves

21

(118-119), and are directly responsible for the structural and functional integrity of a

biofilm.

Bacterial microcolonies imbedded within the EPS are interspersed by voids that connect

to form a network of channels within the matrix (120-121). These allow for the fluid

transport of nutrients, waste products, and cell-signaling molecules (103, 106, 112-113,

115). Confocal laser scanning microscopy shows the morphology of oral biofilms as

having discrete mushroom-shaped stacks with a densely compact sub-layer that is not

continuous - often exposing the substratum (120-123).

Phenotypic characteristics of biofilm cells are markedly different from their planktonic

counterparts (109, 114, 116,124-129). It is now widely accepted that cells within the

microbial species occur as two very separate physiological entities depending on culture

growth conditions (114; 109; 130). Altered cellular genetic expression results in

modified behavioral responses to environmental stimuli (131). In comparison to

planktonic growth, biofilms demonstrate greatly enhanced surface adherence properties

(132), genetic competence (103), lowered susceptibility to anti-microbial agents (133) as

well as increased resistance to acidic conditions (134) and host-immune defense

mechanisms (135). Virulence of the most dominant cariogenic microorganisms within

the oral cavity is thereby highly contingent on bacterial growth as a biofilm (103).

2.3.1.1 Adherence and Colonization

22

The process of plaque adhesion to resin and tooth substrates following initial placement,

eruption, or cleaning is a classic case scenario of any biofilm colonization. It occurs as a

step-wise process (116) instigated by the adhesion of early colonizing planktonic cells to

substratum at a solid-fluid interface. In the oral cavity however, all available substrates

are first conditioned by a thin layer of adsorbed host-derived proteins (136) that spread

over and penetrate substrate micro-porosities within minutes of exposure to the oral

environment (137-139). This acquired pellicle consists of salivary-derived proteins -

such as sialylated mucins, praline-rich proteins, agglutins, phosphate-rich proteins, and

enzymes, dietary components, as well as bacterial metabolic byproducts such as the

streptococcal glycosyltransferase (GTF) have also been identified (140). It plays a vital

role in plaque adhesion in vivo (136).

As microbes move randomly above the surface of the pellicle-coated substratum, cell-

surface adhesins come in close contact with ligands within the pellicle (116). Weak

molecular interactions - such as electrostatic and van der Waals forces - take place in a

specific manner that result in a „lock and key‟ mechanism, loosely adhering the bacterial

cell to the substratum. One example is the initial weak attachment achieved through the

specific molecular interaction of the S. mutans GTF and sialylated mucins of the pellicle

in the oral cavity (141). Once established however, the accumulation of extracellular

glucose polymers derived from metabolically released byproducts result in the formation

of much stronger cellular attachments later on in the colonization process (102).

23

Streptococci comprise approximately 60-90 % of the total population of early colonizers

accumulating on dental surfaces in the first 4 hours following initial eruption or cleaning

(142). Other early colonizing strains include Actinomyces spp., Capnocytophaga spp.,

Haemophilus spp., Prevotella spp., Veillonella spp., and Propionibacterium spp (105).

The proliferation of pioneering species change local physiological conditions that then

encourage the co-adherence and proliferation of late colonizing species (116). The

mature biofilm continues to increase in thickness to the point where it can become

unstable so that large sections slough off into the surrounding environment, colonizing

other clean surfaces in the same cyclical fashion.

2.3.2 Streptococcus Mutans

The dominance of streptococcal cells during early colonization in vivo is attributed to

their ability to bind a wide variety of host-derived molecules and other bacterial cells (16,

105, 143-145). As dental plaque matures, metabolically released lactic acids

accumulating within the EPS drastically reduce local pH conditions in the biofilm within

minutes of host carbohydrate ingestion (146-148). Depending on the age and

composition of the biofilm, as well as concentration of sugars ingested by the host, these

acidic conditions can persist for several minutes to hours (147-148). While the

glycolytic enzymes and cellular processes of less acid tolerant microbial species become

disrupted (149-150), certain microbial species are capable of continuing cellular

processes, thereby out-competing previously established pioneer species, such as

Streptoccus gordonii, Actinomyces spp., Capnocytophaga spp., Haemophilus spp.,

Prevotella spp., Veillonella spp., and Propionibacterium spp (Kolenbrander et al, 2002).

24

Frequent and prolonged cycles of plaque acidification then result in an ecological shift in

plaque microflora tending to favour aciduric species such as S. mutans, S. sobrinus, and

some lactobacilli (101-102, 148, 151-152). S. mutans has long since been identified as

the main cariogenic agent in matured plaque (16, 102, 151, 153) - particularly among

those implicated in secondary caries beneath composite restorations (154). Governed

through a dynamic acid tolerance response (ATR) mechanism, cellular uptake of

nutrients and glycolytic processes are maintained in S. mutans at extracellular pH values

as low as 5.0 (130, 150, 153, 155-158). Continued bacterial metabolic activity

propagates plaque acidification, driving pH even further down – to values well below 4.0

(102, 159- 160). Frequent and prolonged exposure of mineralized tooth surfaces to acidic

plaque conditions are what result in the development of dental caries.

2.3.2.1 Interaction with Resin-based Materials

Salivary proteins are known to have a particularly high affinity for polymeric materials

(139, 161), which could account for the increased growth of S. mutans biofilm found

along composite resin restorative margins (154). Yet it has also been shown that the

salivary pellicle is not necessarily a requirement for S. mutans adherence (162).

Particularly in the presence of sucrose (163), S. mutans does not require the substrate-

conditioning properties of the salivary pellicle to colonize either resin-based or

mineralized tooth substrates (164-167). Interactions taking place at the surface of

polymeric resin materials appear to modulate certain bacterial characteristics relating to

vitality at the cellular level (138,168).

25

Water insoluble glucans associated with a biofilm‟s EPS matrix forms a barrier against

solute diffusion. As a result, much of the leachable residual monomers and

biodegradation byproducts released from the surface of composite resin restorations

accumulate in overlying dental plaque (169). The readily leached TEGDMA and its

associated MA and TEG byproducts, for example, have been shown to penetrate cell

membranes and modulate growth in a concentration and pH-dependent manner (170,

171). In concentrations found in vivo, TEG exposure up-regulates the expression of. at

least two genes in the S. mutans NG8 strain implicated in plaque formation, while MA

appears to negatively influence growth (172-174). Most recently, it has been found that

the Bis-GMA derived BisHPPP byproduct also influences the growth of S. mutans in a

multifactor-dependent manner (129).

Overall, exposure to resin-based biodegradation byproducts impacts the gene expression

and growth of S. mutans plaque differentially (129, 172-174). However this interaction is

highly complex and contingent upon factors which are constantly in flux – temporally

and regionally – under conditions in vivo.

2.3.3 Artificial Models Systems Used to Culture Oral Plaque In Vitro

By definition, biofilm structures are heterogeneous. The distribution of any selected

component in any of the compartments of the biofilm system is non-uniform – this

includes biomass distribution, nutrients, metabolic byproducts, and species composition.

The inevitable variability between and within biofilms often impede consistency and

26

reproducibility of results among in situ plaque investigations (121-123, 175). Biological

variation within samples is also an issue among stationary batch culture grown dental

plaques (167, 176-177). To date, the best method of reproducibly obtaining steady-state

oral biofilm communities representative of phase interface conditions present in the oral

cavity in vitro are through continuous-flow biofilm fermentation systems (103, 178).

Laboratory biofilm fermentors can be any suitable vessel in which microorganisms grow

under continuous media influx while maintaining a constant culture volume (179).

Steady-state introduction of fresh media governs the rate at which cells divide. By

regulating the rate of media perfusion into the vessel, substrate availability in the oral

cavity can be mimicked to attain mean biofilm growth rates comparable to that in vivo

(180). In addition, the laminar flow of media within the vessel emulates the passing of

salivary fluids over growing plaque populations; a determinant of biofilm structural

properties in vivo. Yet while continuous culture systems mimic several of the key

parameters within the oral cavity, it is also critical for these systems to severely restrict

variations in other environmental parameters that may result in varied physiological

responses (179).

External stimuli such as local pH, temperature, carbohydrate source, and species

composition are tightly controlled for through regulatory and measurement devices

incorporated within fermentation systems. In doing so, variable factors that would

otherwise confound results become severely restricted (103). Oral plaque cultured within

these closed settings is highly reproducible, yet still considered to be representative of

27

interactions that take place between the material-biological interfaces under clinical

conditions (65, 120).

Of particular importance for microbiology investigations, bacterial populations under

constant growth conditions have been observed using many different in vitro model

systems – generating mono- or heterogeneous oral biofilm communities over various

substrata (103, 165; 180-182). However, the use of fermentation systems in examining

the influence of oral biofilm growth on the dentinal restorative margin is still fairly new.

In 2001, Matharu et al suspended amalgam-restored dentin samples in a constant depth

film fermentor (CDFF) to mimic the dynamics of microbial activity around amalgam

restored dentinal margins in vivo (93). Post-incubation, SEM analysis of CDFF-

suspended samples revealed the occurrence of material discoloration, as well as bacterial

microleakage within the exposed amalgam-dentin restorative margin (93). Attempts at

evaluating similar interactions between oral bacteria and the exposed restorative margin

of composite resin materials however, have yet to use continuous-flow biofilm

fermentation systems (94, 176).

2.4 MICROSCOPY TECHNIQUES USED TO IMAGE THE RESIN-DENTIN

MARGINAL INTERFACE

Micro- and nanoleakage investigations often base the integrity of the resin-dentin interface

on morphological assessments made from images captured through various SEM and TEM

imaging techniques (46-47, 183). While electron microscopy is a standard technique for

imaging surface morphology of polymerized resin materials (184), it is much less suitable

for imaging biological components within the resin-dentin margin. Several drastic

preparation steps required prior to imaging – including dehydration, fixation, and

28

imbedding of specimens - alter the morphology of naturally hydrated elements within the

specimen. Demineralized dentinal regions, un-reinforced and exposed in the hybrid layer,

for example, may collapse prior to image capture. The same issue is also often cited among

SEM and TEM investigations of biofilms (185). Desiccation of fluid-filled channels within

the biofilm is bound to cause some level of structural collapse and distortion among relative

positions of cells within the previously hydrated matrix (106). In the case of bacterial

microleakage assessments along the interface (93), confocal laser scanning microscopy

(CLSM) is a much more suitable imaging technique (185).

It is possible to view fully intact specimens of much greater thickness, with little to no

sample disruption. CLSM offers improved exclusion of out-of-focus noise and greater

resolution than conventional optical microscopy techniques (186-187) as well as the ability

to capture thin sequential sagital (xz) optical sections allowing for 3-dimensional

visualization of structures. Fully hydrated biofilm structures in their natural state (120, 123,

178, 188-191), as well as within the intact resin-dentin interface (60, 167, 185, 192-195)

have both been three-dimensionally reconstructed. When combined with fluorescent

agents (189), CLSM presents the best available technique for non-invasively imaging the

potential adherence of S. mutans biofilm on and within the resin-dentin interface, where

potential for marginal gaps may exist.

29

2.5 FIGURES

Bis-GMA

Bis-HPPP MA

Figure 2.1 The hydrolysis of two unprotected ester bonds within the 2.2-Bis[4-(2-

hydroxy-3-methacryloyloxy-propoxy)phenyl]propane (Bis-GMA) produces Bis-

hydroxy-propoxy-phenyl propane (Bis-HPPP) as a degradation byproduct, as well

as two methacrylate monomers (MA).

30

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53

CHAPTER 3: Biodegradation of Resin-Dentin Interfaces Increases Bacterial Microleakge 1 (Note: The following has been submitted to the Journal of Dental Research for

publication)

Kermanshahi S. a, Santerre J.P.

a, b, d, Cvitkovitch D.

c, d, Finer Y

a,d *.

a Biomaterials Discipline, Faculty of Dentistry, University of Toronto, 124 Edward Street, Toronto, Ont.,

Canada M5G 1G6 b Department of Chemical Engineering and Applied Chemistry, Faculty of Engineering, University of Toronto,

Toronto, Ont., Canada c Department of Oral Microbiology, Faculty of Dentistry, University of Toronto, Toronto, Ont., Canada

d Institute of Biomaterials and Biomedical Engineering, University of Toronto

3.1 INTRODUCTION

Resin-composite matrices based on 2.2-Bis[4-(2-hydroxy-3-methacryloyloxy-

propoxy)phenyl]propane (Bis-GMA) - undergo significant chemical biodegradation when

challenged by esterase activities (1, 2) contained within human saliva (3). Hydrolytic

cleavage of unhindered ester bonds at both ends of a Bis-GMA unit results in chemical

breakdown, releasing bis-hydroxy-propoxy-phenyl-propane (Bis-HPPP), a marker of

resin-matrix breakdown (4). However, much of the current knowledge of resin

biodegradation stems from observations of external surfaces of composite restorations

interfacing with fluids from the oral cavity or simulated aging solutions (5).

The physical and chemical integrity of a composite restoration‟s adhesive bond layer –

the existing interface between the restoration and the underlying tooth substrate- is the

most significant factor determining long-term clinical restoration success (6). Bacterial

microleakage is the most frequently cited post-operative complication among dentin-

* Based on a thesis to be submitted to the Graduate Department of the Faculty of Dentistry, University of

Toronto, in partial fulfillment of the requirements for the M.A.Sc. degree.

54

bonded composite restorations (7) and secondary caries is the principle cause of failure

(8). Very little research has focused on the impact of biodegradation along the tooth-resin

interface. Of particular concern are proximal and cervical restorations where the

cavosurface margin is formed against wet dentinal substrate (9).

The hypothesis is that exposure of resin-composite restorations to saliva-like esterase

activities accelerates marginal bacterial microleakage.

3.2 MATERIALS AND METHODS

3.2.1 Preparation of Resin-Dentin Specimens

Dentin blocks cut from fully intact sterilized human third molars were bonded

(Scotchbond MP, 3M) to resin-composite (Z250, 3M) under sterile conditions, according

to the manufacturer‟s instructions. Thin incremental layers of composite resin were

packed and photo-polymerized for 40 seconds using a hand-held light curing unit (The

Max, DENSPLY Caulk). A low-speed water-cooled rotary saw with a thin wafering

blade (Isomet, Buehler) was used to prepare standardized resin-dentin specimens with

cross-sectional areas of 3 mm2. All regions of exposed dentin directly adjacent to the

marginal interface were sealed with nail varnish to ensure no access to the resin-dentin

interface through cut dentinal tubules.

3.2.2 Degradation Media Incubation of Resin-Dentin Specimens

Specimens were randomly assigned to the following sterile incubation conditions; 0, 7,

30, or 90-days incubation in phosphate buffer solution (PBS) or pseudocholinesterase-

cholesterol esterase solution (PCE-CE) (n=9). PCE-CE was prepared by dissolving

55

cholesterol esterase (CE) (Item No. 70-1081-01, Lot No. 9750, Genzyme, Cambridge,

MA, USA) and pseudocholinesterase (PCE) (C-5386, Sigma, St. Louis, MO, USA) in

PBS (D-PBS, 21600-010, Gibco, Grand Island, NY, USA) based on methods previously

described (2) to match relevant esterase levels (16 units/ml) found in human saliva (3). A

media replenishment cycle of 48 hours was carried out for all incubated specimens.

Extracted media from defined incubation periods (0, 7, 30, 90 days) were pooled and

stored at 4oC until further analysis.

3.2.3 Bis-HPPP Byproduct Isolation

A WatersTM

high performance liquid chromatography (HPLC) system (2) was used to

isolate and quantify Bis-HPPP (3, 5) Product identification was confirmed by mass

spectrometry (MS/MS) (QStar-XL, Applied Biosystems/MDS Sciex), Sunnybrook

Research Institute, Toronto, Canada).

3.2.4 Incubation of Resin-Dentin Specimens in Chemostat-Based Biofilm Fermentor

(CBBF)

Following assigned incubation in either buffer or PCE-CE media, specimens were

suspended within a closed-system biofilm fermentor designed to cultivate steady-state

monoclonal biofilms of Streptococcus mutans NG8 over interfacial margins (10). Fresh

medium (Todd Hewitt yeast extract supplemented with 10 mM sucrose and 0.01% hog

gastric mucin, 4X diluted) was pumped into the vessel at a dilution rate of D=0.075 per

hour, mimicking the resting flow rate of saliva over human oral tissues (11). Daily

maintenance of the CBBF included optical density readings, viable cell count, and vessel

pH adjustments (7.0±0.5). Specimens were aseptically removed after seven days, rinsed

56

gently with sterile water, and stained using Live/Dead Baclight Bacterial Viability Kit

(Molecular Probes, Eugene, Oregon, USA).

3.2.5 Confocal Laser Scanning Microscopy (CLSM) Analysis

Stained specimens were assessed individually for bacterial penetration using CLSM

(Zeiss LSM 510 META NLO, Carl Zeiss MicroImaging Inc). Six equidistant (2m) Z-

stack series were captured along one side of each resin-dentin interface through a C-

Apochromat 63x/ 1.2 W (water-immersion) objective lens, zoom 2X. All six regions of

interest (ROI) were standardized for orientation; the marginal interface was defined as

that which occurred between the composite-resin and dentinal regions. CLSM Z-stack

images were processed (ImageJ software) (12) to remove background fluorescence and

allow for quantification of bacterial cells.

3.2.6 Statistical Analysis

Two-way ANOVA and Scheffe‟ post-hoc analysis (p<0.05) were conducted to determine

the effect of incubation time and media treatment on the amount of Bis-HPPP present and

the total levels and depth of bacterial cells identified within the resin-dentin interface.

All study groups were run in parallel, with three independent samples in each group.

Each experiment was conducted three separate times (n=9).

57

3.3 RESULTS

3.3.1 Biodegradation

Levels of cumulative Bis-HPPP released from specimens incubated in PCE-CE media

were significantly higher (p<0.0005) than PBS-incubated specimens for all incubation

times (Figure 1A). Total Bis-HPPP accumulation for the 90-day PCE-CE groups was

highest (297±62 g /cm2). In comparison, total Bis-HPPP accumulation for the 90-day

PBS-incubated specimens reached 9.68±0.55 g/cm2, with no Bis-HPPP detected prior to

30 days (Figure 1B). Among the PCE-CE incubation group, the highest rate of Bis-HPPP

daily production occurred within the first 7 days (Figure 1B).

3.3.2 Bacterial Microleakage

Specimens incubated for 90-days demonstrated significantly higher levels of interfacial

cellular penetration (P<0.001) than those incubated under the same culture conditions for

shorter periods of time (Figure 2A). After the 30-day incubation time point, the

cumulative number of bacterial cells found penetrating the marginal interface were found

to be significantly higher among PCE-CE incubated specimens (p<0.005) as compared to

PBS (Figure 2A).

Specimens in the 90-day PCE-CE group demonstrated nearly more than three times as

many (p< 0.005) bacteria than that of specimens incubated in PBS for 90-days (Figure 2B

compared to 2C). Maximum interfacial depths of penetration were also nearly four times

58

deeper among PCE-CE incubated specimens than their PBS counterpart at 90-days

incubation (p<0.05).

CLSM Z-stack images for the top10µm of specimens are shown in Figure 3. Among all

7-day incubated specimens incubated in PBS or PCE-CE media, resin-dentin interfacial

structural morphology resembled that of controls, which had been incubated in either

PBS or PCE-CE media but not suspended within the CBBF (Figures 3A and 3B, data for

controls not shown). Limited amounts of adherence and penetration of S. mutans biofilm

cells was observed among interfacial surface micro-porosities for 7-day incubated

specimens, up to a maximum depth of 4 m. All component layers of the resin-dentin

margin appeared linearly oriented to one another, and well-infiltrated by the adhesive

resin. Tubules of the hybrid layer were seen to be structurally intact; consistently spaced

apart, and oriented parallel to one another.

In 30-day specimens, morphological changes to the resin-dentin interfacial margin

emerged, albeit to varying extents, depending on incubation medium (Figures 3C and

3D). For PBS specimens, dentinal tubules of the hybrid layer commonly showed finite

structural degradation, particularly near the composite resin-resin adhesive region of the

marginal interface. For PCE-CE specimens, in addition to dentinal tubules fracture,

complete collapse of the tubules was frequently observed (Figure 3D). Among nearly all

interfacial ROIs examined from 30-day PCE-CE specimens, the adhesive interface

appears poorly delineated and more irregular; Figure 3Dii and 3Diii show an example of

a blister-like void (region labeled b) occurring in the adhesive zone.

59

In 90-day PBS specimens (Figures 3E, 3F), junctions between the resin restoration, the

adhesive layer and the hybrid layer were less distinguishable. Where junctions were

somewhat identifiable, they appeared blistered and non-linear with an undulating pattern

across the x-axis of each CLSM image (Fig. 3E). Dentinal tubules frequently showed

structural failures– primarily near the top and bottom of the hybrid layer. Periodicity

among tubules was interrupted, suggesting that some may have broken off or been

entirely displaced. Among the 90-day PCE-CE specimens, junctions between the

component layers of the interfacial margin were markedly less distinguishable and

dentinal tubules lacked periodicity all together (Figure 3F).

Two of the numerous examples of gross interface deformation found in the PCE-CE

samples are shown in Figure 4. Figure 4A contains a distinctive interfacial void at the

top of the hybrid layer, that was bordered top and bottom by S. mutans adherence. At

sample depths of 6 m and over, biofilm growth extended to span the width (Figure

4Aiv) and length of the voids (Figures 4Av to 4Aviii). Figure 4B shows an interfacial

gap spanning more than 20 m. The series of sagittal sections through this particular

ROI reveal a characteristically distinct 3-dimensional mushroom-shaped pattern of S.

mutans biofilm growth (12). The biofilm cellular penetration extended up to 34 m

beneath the surface of the interface (images not shown).

60

3.4 DISCUSSION

Relative to PBS controls PCE-CE incubated resin-dentin specimens generated

significantly higher amounts of Bis-HPPP (p<0.0005) at all incubation time points

(Figure 1) and resulted in greater bacterial surface adherence and penetration along the

resin-dentin marginal interface with time (Figure 2). Therefore, the hypothesis that

exposure of resin-composite restorations to saliva-like esterase activities accelerates

marginal bacterial microleakage was accepted. The data demonstrated that

biodegradation of resin-adhesives and composites were a time-dependent process that

progressively compromised the marginal integrity of dentin-bonded interfaces with

increased incubation time.

3.4.1 Biodegradation

A salivary-like esterase solution was developed using PCE and CE components

maintained at activity levels comparable to that of human saliva. A pilot study

determined the kinetic Bis-HPPP-release profile of PCE-CE incubated cured composite

resin samples to be comparable to that of cured composite resin samples incubated in

both human saliva-derived esterase activities and human saliva itself (p>0.05).

Periodic daily incremental release rates of Bis-HPPP within the 90-day period for PCE-

CE incubated varied over time (Figure 1B). The highest rate of degradation product

accumulation occurred within the first 7 days of PCE-CE incubation (13.8±1.49 m/cm2

per day). This decreased by a factor of 4 between 8 to 14 days of PCE-CE incubation,

and reached a constant release rate of 1.0±0.1 m/cm2 per day between days 30 and 90.

Theoretically, for the first seven days of incubation, hydrolytic reaction rates are defined

61

primarily by the catalytic activity of the esterases. By the 8th

day of incubation, the

number of readily accessible ester linkages within the Bis-GMA-based resin matrix

gradually declines. The rate-controlling factor becomes the physical access of esterases to

the un-reacted ester substrate, slowing down the hydrolytic process. After 30 days of

incubation, the most readily accessible ester linkages within the resin matrix have

become hydrolyzed. At this point, longer-term biochemical breakdown of the resin

matrix depends on rates at which previously inaccessible ester linkages become

unmasked by the elution of degraded oligomers (14, 15).

3.4.2 Bacterial Microleakage

Bacterial adherence and penetration among the superficial sub-layers (0–4 m) of

minimally compromised interfacial margins were mainly localized to the top and bottom

of the hybrid layer in specimens incubated for 7-days with either PBS or PCE-CE

(Figures 3A and 3B). Intrinsic interfacial porosities are often formed during bond

application; potentially generated by polymerization shrinkage at the top of the hybrid

layer, or incomplete resin impregnation of demineralization dentin occurs at the bottom

(16). Both phenomena may also account for the superficial bacterial microleakage

observed at the interfacial sub- layers of control resin-dentin specimens un-incubated

with degradation media (data not shown).

Discrepancy between the depth of the demineralization and resin infiltration is common

among commercial three-step “etch-and-rinse” adhesives such as Scotch Bond Multi-

Purpose (17). Oral streptococcus cells are approximately 0.5 to 0.7 m in diameter (18).

Given access, they are capable of penetrating expanding voids at the bottom of the hybrid

layer and directly bind to intra-tubular collagen type I components of the dentinal tubules

62

(18). Furthermore, given the widely-reported effects of water sorption at the top and

bottom of the restorative interface (19), as well as associated elution of adhesive resin

components over time (20), nano-meter sized interfacial voids can expand with prolonged

incubation in media (16). The results from this investigation corroborate such findings;

since localization of bacterial microleakage among resin-dentin specimens incubated for

7 and 30-day time periods were centralized near the top and base of the hybrid layer (Fig

3A, 3B, 3C, 3D). At 90-days, significant interfacial disruptions were found and bacterial

microleakage for all specimens at this time period was non-specific and occurred across

the entire interfacial span (Fig 3E, 3F).

Overall, it can be said that patterns of morphological change and bacterial microleakage

at the interface of the 90-day PBS-incubated specimens depicted the effects of base-line

hydrolytic processes (20). In comparison, an assessment of the 90-day PCE-CE

incubated specimens demonstrated the compounding degradative effects that salivary-like

esterase activities have on the structural integrity of the resin-dentin interface, beyond

that of the un-catalyzed hydrolytic process over time (PBS condition).

Qualitative assessments of interfacial structural integrity among resin-dentin specimens

led to the conclusion that while incubation in PBS altered the morphology of the marginal

interface over time (20), exposure to salivary-like esterase activities of PCE-CE media

greatly amplified the intrinsic effects of hydrolytic processes. The reduction of signal

scatter in the inter-tubular layer of the hybrid zone suggests a loss of resin and/or mineral

content; a morphological change consistent with that of carious dentin (21). While an

overall reduction in red fluorescence signal scatter was observed at inter-tubular regions

63

of the hybrid layer in 90-day PBS-incubated specimens (compare Figure 3A to 3E), it

was entirely absent among that of 90-day PCE-CE incubated specimens (Figure 3F).

The presence of blister-like voids (Figures 3Dii and 3Diii) and the observed undulating

pattern of the adhesive resin layer (Figures 3D, 3E and 3F) may be a consequence of

interfacial water sorption (22) causing swelling and plasticization of resin polymers (23).

Given that on-going hydrolytic processes can propagate marginal gap formation over

time (24), it is then not surprising that the largest marginal gaps were found exclusively

among 90-day PCE-CE incubated specimens (Figure 4), which generated the greatest

amounts of Bis-HPPP.

It was also within the expanded marginal gap region of 90-day PCE-CE incubated

specimens that the most extensive colonization of S. mutans biofilms were found (Figure

4B). Highly characteristic biofilm structures (13) were anchored to the composite resin

or dentinal axial walls of marginal gaps spanning 10 m or more. Recently, Totiam et al

(25) suggested that larger-sized marginal gaps provide the necessary space and access to

nutrients necessarily for successful colonization by larger numbers of microorganisms.

3.5 CONCLUSION

It is the current belief that secondary loss of marginal integrity is primarily attributed to

mechanical forces such as occlusal loading (9; 26) as well as thermal stress and

polymerization contraction (26). Yet, increasingly emerging data in the scientific

literature has suggested a potential for secondary loss of adhesion due to in vivo chemical

64

attack (6, 9, 24). The current findings make this case abundantly clear. Evidence of

increased bacterial penetration coupled with dentin demineralization suggests that the

biodegradation process can contribute to the formation of recurrent decay – the most

common cause of restoration failure.

Results from this study also demonstrated high reproducibility as well as clinical

relevancy - a factor which is imperative when evaluating biomaterials with an in vitro

experimental system. We believe that this model shows great potential for further

development into a standardized testing system of biochemical stability among various

commercial adhesive and composite materials prior to use in clinical settings. Where

hybrid layer interruption and marginal gaps do occur, the current system presents a

practical non-invasive imaging method for intact biofilms adhering to and proliferating

on and within the resin-dentin interface. To the best of our knowledge, the current

investigation provides the first physiologically relevant in vitro characterization of

bacterial microleakage within the resin-dentin interface.

3.6 ACKNOWLEGEMENTS

This study was supported by a Canadian Institute of Health Research Grant MOP 68947.

65

A

B

0

50

100

150

200

250

300

350

0 10 20 30 40 50 60 70 80 90

Time (days)

Am

ou

nt

(

g/c

m2

)

PCE/CE Solution

PBS

1.03 0.96

0.00 0.00 0.21 0.05

3.763.62

13.76

0.13

0

2

4

6

8

10

12

14

16

0 to 7 8 to 14 15 to 30 31 to 60 61 to 90

Incubation Time Period (Day)

Me

an

In

cre

me

nta

l B

isH

PP

P

Re

lea

se

(

g/c

m2 )

PCE/CE Solution

PBS

*

*

*

3.7 FIGURES

Figure 1

66

A

0

400

800

1200

1600

0 2 4 6 8 10

Depth of Penetration (m)

Inc

rem

en

tal

nu

mb

er

of

ce

lls

90 days

30 days

14 days

7 days

0 days

PBS ConditionB

0

400

800

1200

1600

0 4 8 12 16 20 24 28 32

Depth of Penetration (m)

Incre

men

tal

nu

mb

er

of

cells

90 days

30 days

14 days

7 days

0 days

Salivary-like Esterase (PCE-CE) ConditionC

135.1

373.4

1027.5

128.7294.8

1524.5

4091.5

190.6

0

1000

2000

3000

4000

5000

6000

7 14 30 90

Time (days)

To

tal

nu

mb

er

of

cells

pen

trati

ng

th

e m

arg

inal

inte

rface

PBS

PCE-CE

Figure 2

67

vi

vi

ii

D

E

A

F

C

(b

)

Depth: 0.0 m Depth: 2.0 m

A


Depth: 8.0 m Depth: 10.0 m B




Composite

Dentin

Composite

Dentin

Composite

Dentin

Composite

Dentin

Composite

Dentin

Composite

Dentin

i ii

iii iv

v vi

i

iii

v

i

iii

v

i

iii

v

ii

iv

vi

i

iii

ii

iv

i

v vi v vi

Hybrid Layer Hybrid Layer



b

b

ii

vi

iii

iv

vi

vi

iv

ii

Figure 3

68

A B

Composite i

Dentin

Marginal Gap/

interfacial void

Dentin

20

g

Marginal Gap/

interfacial void

i ii

iii iv

v vi

vii viii

v

vii viii

iii

Composite

iv

vi

ii

Figure 4

69

Figure 3. Selected Z-stack image series‟ captured from interfacial margins of resin-

dentin specimens assigned to either (A) 7- day PBS incubation, (B) 7- day PCE-CE

incubation, (C) 30- day PBS incubation, (D) 30- day PCE-CE incubation, (E) 90-

day PBS incubation, (F) 90- day PCE-CE incubation. All three interfacial zones

(composite, dentin, hybrid layer) are clearly distinguishable in A, B, C, D; however

in E and F the organization of these marginal components are disrupted. Resin

impregnation of dentinal tubules is absent (E and F) and no hybrid zone can be

distinguished. Specimens were stained using Live/Dead Baclight Viability Kit

(magnification X62, 2X zoom). Live cells indicated by green fluorescence through

interaction with Syto9; dead cells indicated by red fluorescence through interaction

with Propidium Iodide.

Figure 4. Z-stack image series captured at interfacial ROIs of two 90-day PCE-CE

incubated resin-dentin specimens. (A) Interfacial void spanning approximately 4-5

m in height (B) Interfacial void spanning over 20 m in height. Characteristic of

three-dimensional biofilm growth are interstitial voids that can be seen among

fluorescently stained S. mutans microcolonies. In (B), large mushroom-shaped

biofilm structures are found colonizing both the top and bottom axial walls.

Specimens were stained using Live/Dead Baclight Viability Kit (magnification X62,

2X zoom). Live cells indicated by green fluorescence through interaction with

Syto9; dead cells indicated by red fluorescence through interaction with Propidium

Iodide.

Figure 2. (A) Mean cumulative number of bacterial cells found penetrating the

marginal interface for PBS controls and PCE-CE incubated specimens over time.

(B) Mean number of cells at each depth of penetration at interfacial regions of

interest (ROI) of PBS incubated and (C) PCE-CE incubated specimens for 0, 7, 14,

30, and 90-days (pH 7, 37 oC). All data are reported with standard error of the mean

(n=3).

3.7 FIGURE CAPTIONS

Figure 1. (A) Mean cumulative amount of Bis-HPPP produced from resin-dentin

specimens incubated in PCE-CE solution or PBS for 7, 14, 30, and 90 days (pH 7,

37 oC). All data are reported with standard error of the mean (n=3). (B) Mean daily

incremental amount of Bis-HPPP produced from resin-dentin specimens incubated

in PCE-CE or PBS buffer during pre-set incubation time intervals. All data are

reported with standard error of the mean (n=3).

70

3.8 REFERENCES

1) Jaffer F, Finer Y, Santerre JP. (2002). Interactions between resin monomers and

commercial composite resins with human saliva derived esterases. Biomat

23:1707-1719.

2) Finer Y, Santerre JP (2004). Salivary esterase activity and its association with the


3) Lin BA, Jaffer F, Duff MD, Tang YW, Santerre JP (2005). Identifying enzyme



4) Finer Y, Santerre JP (2003). Biodegradation of a dental composite by esterases:

dependence on enzyme concentration and specificity. J Biomater Sci Polym Ed.

14:837-49.

5) Santerre JP, Shajii L, Leung BW. (2001). Relation of dental composite

formulations to their degradation and the release of hydrolyzed polymeric-resin-

derived products. Crit Rev Oral Biol Med 12:136-151.

6) Donmez N, Belli S, Pashley DH, Tay FR (2005). Ultrastructural correlates of in

vivo/in vitro bond degradation in self-etch adhesives. J Dent Res 84:355-359.

7) Murray PE, Hafez AA, Smith AJ, Cox CF (2002). Bacterial microleakage and

pulp inflammation associated with various restorative materials. Dent Mat

18:470-478.

8) Hickel R, Manhart J (2001). Longevity of restorations in posterior teeths and

reasons for failure. J Adhes Dent 3:45-64.

9) Bouillaguet S (2004). Biological risks of resin-based materials to the dentin-pulp

complex. Crit Rev Oral Biol Med 15:47-60.

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10) Cvitkovitch DG, Li YH, Ellen RO (2003). Quorum sensing and biofilm

formation in Streptococcal infections. J Clin Invest 112:1626-32.

11) Pratten J, Andrews CS, Duncan QMC, Wilson M (2000). Structural studies of


Micriobiol Lett 189: 215-218.

12) Hope CK, Clements D, Wilson M (2002). Determining the spatial distribution of


profiling. J Applied Microbiol 93:448-455.

13) Lewandowski Z, Beyenal H, Myers J, Stookey D (2007). The effect of

detachment on biofilm structure and activity: the oscillating pattern of biofilm

accumulation. Water Sci Technol. 55:429-36.

14) Finer Y, Santerre JP (2007). Influence of silanated filler content on the

biodegradation of bisGMA/TEGDMA dental composite resins. J Biomed Mater

Res A. 81:75-84.

15) Finer Y, Santerre JP (2004). The influence of resin chemistry on a dental

composite's biodegradation. J Biomed Mater Res A. 69:233-46.

16) Suppa P, Breschi L, Ruggeri A, Mazzotti G, Prati C, Chersoni S, et al. (2005).

Nanoleakage within the hybrid layer: a correlative FEISEM/TEM investigation. J

Biomed Mater Res B Appl Biomater. 73:7-14.

17) Spencer P, Wang Y (2002). Adhesive phase separation at the dentin interface

under wet bonding conditions. J Biomed Mater Res. 62:447-56.

18) Love RM, Jenkinson HF (2002). Invasion of dentinal tubules by oral bacteria.

Crit Rev Oral Biol Med 13:171-183.

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19) Sauro S, Watson TF, Mannocci F, Miyake K, Huffman BP, Tay FR, et al. (2008).

Two-photon laser confocal microscopy of micropermeability of resin-dentin

bonds made with water or ethanol wet bonding. J Biomed Mater Res B Appl

Biomater. 2008 Dec 17. [Electronically published ahead of print]

20) Pashley DH, Tay FR, Yiu C, Hashimoto M, Breschi L, Carvalho RM (2004).

Collagen degradation by host-derived enzymes during aging. J Dent Res 83:216-

221.

21) Zavgorodniy AV, Rohanizadeh R, Swain MV (2008). Ultrastructure of dentine


22) Sauro S, Pashley DH, Mannocci F, Tay FR, Pilecki P, Sheriff M, et al. (2008).

Micropermeability of current self-etching and etch-and-rinse adhesives bonded to

deep dentine: a comparison study using double-staining/confocal microscopy

technique. Eur J Oral Sci 116:184-193.

23) Ferracane JL. (2006) Hygroscopic and hydrolytic effects in dental polymer

networks. Dent Mater 22:221-222.

24) Hashimoto M, Tay FR, Ohno H, Sano H, Kaga M, Yiu C, Kumagai H, Kudou Y,

Kubota M, Oguchi H (2003). SEM and TEM analysis of water degradation of

human dentinal collagen. J Biomed Mater Res Part B: Appl Biomater 66:287-298.

25) Totiam P, Gonzalez-Cabezas C, Fontana MR, Zero DT (2007). A new in vitro


41:467-473.

26) van Noort R (1994). Introduction to Dental Materials. Papel, Spain: Times Mirror

International Publishers Limited; pp. 3-145.

73

74

CHAPTER 4: GENERAL DISCUSSION

The primary objective of this project was to utilize a salivary-like esterase solution in

simulating the biodegradation of composite resin-dentin marginal interfaces that takes

place in vivo, and to characterize the degree of loss of marginal integrity that results

through bacterial microleakage.

Similar to previous studies (6,7), Bis-HPPP went undetected from PBS-incubated resin-

dentin specimens within the first 30 days of incubation. Between 30 and 90 days of PBS

incubation however, minor accumulations of Bis-HPPP do occur, presumably as a result

of an un-catalyzed hydrolytic reaction with water. Analysis between media conditions

(PBS vs. PCE-CE) at 30- and 90-day incubation time points revealed cumulative levels of

Bis-HPPP production significantly higher among PCE-CE incubated specimens than their

PBS-incubated counterparts (p<0.05).

4.1 DISCUSSION RE: HYPOTHESIS #1

Historically, secondary loss of marginal integrity has primarily been attributed to

mechanical forces such as occlusal loading (8,9) as well as thermal expansion and

contraction (9). Increasingly, more recent scientific literature notes the potential for

secondary loss of adhesion due to in vivo chemical attack (2,3). Results from the current

investigation attest to this fact; chemical biodegradation is an on-going process which

progressively compromising the clinical value of the resin restorative interface with time.

75

As well, salivary-like esterase activities compound these effects and accelerate the

degradation process. With increased Bis-HPPP production, a marked decline in

interfacial integrity was found.


In the past CLSM has been used extensively to investigate the morphology of hydrated

biofilms (10-12); less common is the use of CLSM in imaging the resin-dentin interface

(13,14). Given the ability to non-invasively image undisturbed, intact biofilms (15)

colonizing both the surface and sub-surface of experimental substrates, CLSM combined

with fluorescent staining was shown to be an effective method for assessing bacterial

microleakage at the restorative interface. In the future, the combination of these

techniques will offer great potential for further analyses into psychio-chemical processes

taking place within the interfacial microenvironment.


The penetration and proliferation of S. mutans NG8 biofilm within the resin-dentin

interface was visualized through CLSM analysis. Significant differentiations in the

interfacial integrity of PBS and PCE-CE incubated resin-dentin specimens were made

following 30 and 90 day incubation time points (p<0.0001). Maximum depths of

interfacial biofilm penetration among those incubated in PCE-CE media over the course

of 90 days was found to be 3 times greater than that of 90-day PBS incubated specimen.

In addition, cumulative numbers of cells found penetrating resin-dentin interfaces were 3

76

to 4 times higher among 90-day PCE-CE incubated specimens than 90-day PBS

incubated specimens.

Intact interfacial morphology is characterized as having distinct, proportionally stacked

layers; comprised of mineralized dentin, resin-infiltrated demineralized dentin, and

adhesive resin (13,14). Similar to controls, observations of interfacial morphology made

among all 7-day incubated specimens– regardless of media type (ie. PBS or PCE-CE) –

fit this characterization. However some bacterial adherence and penetration among the

superficial sub-layers (0 – 4 m) of these more or less intact interfacial margins were

found; mainly localized to the top and bottom of the hybrid layer. It is know that intrinsic

interfacial porosities are often formed during bond application; the potential for

polymerization shrinkage exists at the top of the hybrid layer, while incomplete resin

impregnation of demineralization dentin occurs at the bottom (16,17).

It is at these vulnerable interfacial zones that the prolonged effects of water sorption and

associated elution of adhesive resin components over time (18,19) can cause nano-meter

sized interfacial voids to expand (17). It was shown in the present investigation that the

localization of initial bacterial microleakage found among resin-dentin specimens

incubated for 7 days begin at the restorative margin walls and progressively expand with

time. Among specimens incubated for 90 days, the localization of bacterial microleakage

was non-specific and ultimately spanned the entire height of the restorative margin.

77

The progressive reduction of red fluorescence signal scatter (compare Figure 3A to 3E) in

CLSM images taken of the hybrid layer‟s inter-tubular regions with increased

biodegradation suggest a loss of resin and/or mineral content; a morphological change

consistent with that of carious dentin (20). Among specimens incubated in PCE-CE

media for 90-days however, this red fluorescent signal from inter-tubular regions was

entirely absent (Figure 3F) – suggesting a complete lack of resin content remaining

within the hybrid layer.

Resin penetration of demineralized dentin serves an adhesive function in dentin bonding,

but it also acts to preserve the integrity of the bare collagen network (18, 21). When

absent, naked collagen within the marginal zone becomes highly susceptible to

proteolytic effects associated with esterases contained within human salivary enzymes

(19). Moreover, accumulating metabolic byproducts associated with biofilm proliferation

(10) create acidic microenvironments within the resin-dentin restoration margin.

Endogenous matrix metallo-proteins (MMPs) of mineralized dentin (22) become

activated under conditions of reduced pH and are associated with auto-degenerative

processes contributing to caries pathogenesis (23). It is highly likely that where the

greatest interfacial S. mutans biofilm accumulations were found, activated dentinal

MMPs contributed to the digestion of collagen components within the marginal interface

(23,24).

Because on-going hydrolytic processes propagate marginal gap formation overtime (18),

the largest marginal gaps were found to reside exclusively among 90-day PCE-CE

78

incubated specimens (Figure 4). It was also within these expanded marginal gaps that the

most extensive colonization of S. mutans biofilm was found. Highly characteristic

biofilm structures (10,11) were anchored to composite resin or dentinal axial walls of

marginal gaps spanning 10 m or more. Recently, Totiam et al (2007) suggest that

larger-sized marginal gaps provide the necessary space and access to nutrients necessarily

for successful colonization by larger amounts of microorganisms (25).

Bacterial microleakage occurs at the resin-dentin interface in vivo, contributing to

secondary caries and postoperative sensitivity (26). Matharu et al (2001) previously

characterized the penetration of bacterial cells at the amalgam-tooth restorative interface

in vitro (27), while Zivkovic et al (2001) attempted the same along the composite resin

restorative margin (28). However to the best of our knowledge, the current investigation

provides the first physiologically relevant in vitro characterization of bacterial

microleakage within the resin-dentin interface.

4.4 REFERENCES

1. Nakamura M, Slots J.(1999) Salivary enzymes origin and relationship to

periodontal disease. J Periodont Res;18:559–69.





composite‟s biodegradation. J Biomed Mater Res A. 69(2)69A:233-246.

79



23:1707-1719.


composite's biodegradation. J Biomed Mater Res A. 69(2):233-46.






8. Bouillaguet S. (2004) Biological risks of resin-based materials to the dentin-

pulp complex. Crit Rev Oral Biol Med 15(1):47-60.



10. Lewandowski Z, Beyenal H, Myers J, Stookey D (2007). The effect of

detachment on biofilm structure and activity: the oscillating pattern of biofilm

accumulation. Water Sci Technol. 55:429-36.

11. Hope CK, Clements D, Wilson M. (2002) Determining the spatial distribution

of viable and nonviable bacteria in hydrated microcosm dental plaques by

viability profiling. J Applied Microbiol 93:448-455

12. Sharma A, Inagaki S, Sigurdson W, Kuramitsu K (2005) Synergy between

Tannerella forsythia and Fusobacterium nucleatum in biofilm formation. Oral

Microbiol Immunol 20:39-42

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80

13. Pioch T, Jakob H, Garcia-Godoy F, Gotz H, Dorfer C, Staehle H. (2003)

Surface characteristics of dentin experimentally exposed to hydrofluoric acid.

Eur J Oral Sci 111:359-364

14. Pioch T, Jakob H, Garcia-Godoy F, Gotz H, Dorfer C, Staehle H. (2003)

Surface characteristics of dentin experimentally exposed to hydrofluoric acid.

Eur J Oral Sci 111:359-364



Micriobiol Lett 189(2000): 215-218

16. Breschi L, Gobbi P, Lopes M, Prati C, Falconi M, Teti G, Mazzotti G. (2003)

Immunocytochemical analysis of dentin: A double-labeling technique. J Biomed

Mater Res 67A:11-17.

17. Suppa P, Breschi L, Ruggeri A, Mazzotti G, Prati C, Chersoni S, Di Lenarda R,

Pashley DH, Tay FR.(2005) Nanoleakage within the hybrid layer: a correlative

FEISEM/TEM investigation. J Biomed Mater Res B Appl Biomater. 73(1):7-

14.

18. Hashimoto M, Tay FR, Ohno H, Sano H, Kaga M, Yiu C, Kumagai H, Kudou

Y, Kubota M, Oguchi H. (2003) SEM and TEM analysis of water degradation

of human dentinal collagen. J Biomed Mater Res Part B: Appl Biomater 66B:

287-298.


Collagen degradation by host-derived enzymes during aging. J Dent Res 83:

216-221.



81

20. Zavgorodniy AV, Rohanizadeh R, Swain MV. (2008) Ultrastructure of dentine


21. Yang B, Adelung R, Ludwig K. (2005) Effect of structural change of collagen

sibrils on the durability of dentin bonding. Biomaterials 26:5021-5031.





Tjaderhane L, Di Lenarda R, Breschi L. (2008) Immunohistochemical

identification of MMP-2 and MMP-9 in human dentin: Correlative FEI-

SEM/TEM analysis. J Biomed Mater Res, March 11, 2008.

24. Agematsu H, Abe S, Shiozaki K, Usami A, Ogata S, Suzuki K, Soejima M,

Ohnishi M, Nonami K, Ide Y. (2005) Relationship between large tubules and

dentin caries in human deciduous tooth. Bull Tokyo Dent Coll 46 (1-2): 7-15.

25. Totiam P, Gonzalez-Cabezas C, Fontana MR, Zero DT. (2007) A new in vitro


41:467-473.

26. Taylor DF, Bayne SC, Leinfelder KF, Davis S, Koch GG. (1994) Pooling of

long term clinical wear data for posterior composites. Am J Dent 7:167-174.


(2001) A new in vitro model for the study of microbial microleakage around

dental restorations : a preliminary qualitative evaluation. Inter Endo J 34 :547-

553.

82

28. Zivkovic S, Bojovic S, Pavlica D. (2001) Bacterial penetration of restored

cavities. Oral Surg Oral Med Oral Path Oral Radiol Endod 91:353-358.

83

CHAPTER 5 – CONCLUSIONS

Conclusion #1

Human salivary-like esterase activities can significantly degrade the integrity of the resin-

dentin interfacial margin. The release of Bis-HPPP degradation byproduct from

composite resin materials used to construct resin-dentin specimens was found to be

instigated through incubation with PCE-CE media at levels comparable to that of proteins

directly derived from human saliva.

Conclusion #2

The in vitro experimental model system developed in this study was capable of

investigating interfacial bacterial microleakage with high reproducibility as well as clinical

relevancy - a factor imperative in the evaluation of biomaterials. As a result, the present

model has shows great potential for further development into a standard methodized

assessment of the biochemical stability of various commercial adhesive and composite

materials. What is more, this study attests to the feasibility of non-invasive imaging

techniques in the evaluation of intact biofilms found adhering to and proliferating on and

within the resin-dentin interface where hybrid layer interruption and marginal gaps have

occurred.

84

CHAPTER 6 – RECOMMENDATIONS

1 - The success of any biomaterial depends on the nature interactions taking place at its

biological interface. As a result, future investigations needs to focus on the physico-

chemical processes taking place within the interfacial microenvironment. The family of

peptidases known as matrix metalloproteinases (MMPs) is capable of cleaving collagen

at sites of specific amino acids. Inactive MMPs intrinsically bound to mineralized dentin

(1) can become activated in low pH microenvironments that result from localized

metabolic activity of bacterial cells (2). In CLSM images, the interfacial zones with the

greatest structural deterioration were also the sites containing the largest volume and

deepest penetration of S. mutans biofilm. While this trend was noted, the specific

mechanisms of interaction between bacterial metabolic activity within the resin-dentin

interface and activation of host-derived MMPs were not targeted by the present study.

Future studies should narrow in on host-biofilm metabolic interactions taking place

within the compromised resin-dentin interfacial microenvironment and delineate

contributions of host-derived peptidases in the degradation of exposed collagen fibrils

(3,4).

2 –Pathogenic characteristics of biofilm penetrating the resin-dentin interface were not

addressed in the present study. The individual number of cells found penetrating the

resin-dentin interface was manually quantified, but the ratio and localization of live to

dead cells was not recorded. Assessing bacterial cell % viability of biofilms penetrating

the resin-dentin restorative interface could provide valuable information. Quantifying

85

fluorescence intensity of stained biofilms has been previously used to determine biofilm

volume and vitality (5). Fluorescence intensity of specifically marked live and dead

bacterial cells can be converted to cell count through computation of a calibration curve

of intensity to known cell counts. However to do so, highly specific non-overlapping

fluorescence markers targeting only live/dead bacterial cells are required. In addition,

auto-fluorescence of background substrate must be completely eliminated prior to such

analyses. Due to inherent interactions between both dentin and composite resin materials

with the fluorescent dyes used in this study, such methods could not be employed.

3 –The development and use of immuno-flourescent markers targeting bacterial cells or

alternatively, their metabolically released byproducts, present an ideal methodology for

assessing biofilm vitality within the resin-dentin interface. Immuno-fluorescence

combines the specificity of antibodies with the high sensitivity of fluorescence. The

antigen-antibody reaction is highly selective, and so it can be applied for identification,

localization, and visualization of cells within biofilm. Thus far in dental tissue research,

studies such as Breschi et al (2003) have used immunocytochemical analysis to assess for

morphological characteristics of dentin (6). The development of antibodies specific for

cell surface epitopes of oral bacterial and their subsequent use in resin-dentin bacterial

microleakage analysis needs to be encouraged. Immuno-flourescent probe specificity

can be tested in biofilm models in vitro.

4 – The green fluorescent protein (GFP) of the jellyfish Aequorea Victoria fluoresces

with fluorescein-like characteristics. It is widely used as a strongly visible fluorescent

86

reporter molecule which is species-independent and does not require co-factors of

substrates (7). Gene coding can be constructed for a fusion of GFP with almost any other

protein and the resulting fluorescent fusion should localize and behave similarly to the

original protein. The method allows protein localization to be visualized without having

to inject cells or purify and label proteins (8). Early attempts aimed at developing a GFP

infused NG8 strain for use during this investigation were made. However there were

several limiting factors that led us to believe that the expression of GFP would not be

strong enough to be detected within the resin-dentin specimen construct.

Variable pH and oxygen limitation within resin-dentin interfacial micro-environment

presented the most significant obstacles. Microelectrode studies show that oxygen

concentration and pH is not uniform within the biofilm 3D micro-structure; both pH and

oxygen levels fall progressively approaching the substratum (9,10). As well, because

GFP infusion is random, the gene itself may have ended up in a part of the genome not

expressed under biofilm conditions (as cultivated under CBBF conditions), discouraged

subsequent attempts at developed a GFP-infused S. mutans NG8 strain. Future studies

may focus on developing such a strain as an alternative to fluorescent dyes or immuno-

fluorescent markers in studying the penetration of bacterial biofilms within the resin-

dentin interface.

4 – Gwinnett and Yu (1995) demonstrated the highly degradative effects of long-term

water storage on resin-dentin bonds in vitro (11). Following 30 day incubation in

water/ethanol, results from Lee et al (1995) suggests that the bonding layer degrades

87

faster than the corresponding composite restoration (12). Though not many

investigations to date have been focused specifically on the degradation of the hybrid

layer, there are several reasons why it is expected that significant hydrolytic degradation

occurs at the restorative margin. It has long been know that bonding resins contribute

significantly to the eluted resin components detected (13). In comparison to more

hydrophobic methacrylate monomers, HEMA is highly water soluble (14) and easily

eluted from polymerized matrices. Future biodegradation studies should work to develop

methodology that can target the biodegradation of the marginal interface specific from

that of the bulk restoration surface.

In addition, leaching of unbound HEMA monomers (15) along with other low weight

oligomers within the resin-dentin interface is a time-dependent process (16). As

leachable components of the resin-dentin adhesive layer are eluted, the microstructure of

the marginal interface changes over time. Hashimoto et al (2003) found that interfibrillar

spaces between the collagen fibrils of fractured hybrid zones appear wider in specimens

that incubated (aged) in water for 1 year, as opposed to those incubated for only 24-hours

(16). Whereas the effects of long-term aging in water have been well-documented, the

same is not clear for salivary-like esterase activities. Based on results of the present

investigation, it is hypothesized that effects of long-term aging under the presence of

salivary-like esterase activities would far exceed that of incubation in water alone. Future

investigations utilizing the present biodegradation model should incorporate experiment

groups incubated for longer periods of time; up to a minimum of 1-year incubation in

either PBS or PCE-CE media.

88

6.1 REFERENCES





Tjaderhane L, Di Lenarda R, Breschi L. (2008) Immunohistochemical

identification of MMP-2 and MMP-9 in human dentin: Correlative FEI-

SEM/TEM analysis. J Biomed Mater Res, March 11, 2008.



4. Amaral FLB, Colucci V, Palma-Dibb RG, Corona SAM. (2007) Assessment of In

Vitro methods used to promote adhesive interface degradation: A critical review.

J Esthet Restor Dent 19: 340-354.

5. Takenaka S, Iwaku M, Hoshino E. (2001) Artificial Pseudomonas aeruginosa

biofilms and confocal laser scanning microscopic analysis. J Infect Chemother

7:87-93.

6. Breschi L, Gobbi P, Lopes M, Prati C, Falconi M, Teti G, Mazzotti G. (2003)

Immunocytochemical analysis of dentin: A double-labeling technique. J Biomed

Mater Res 67A:11-17.

7. Herman B. (1998). Fluorescence Microscopy. BIOS Scientific Publishers Ltd,

Oxford, UK, pp. 64-68.

89

8. Stretton, S., Techkarnjanaruk, S., McLennan, A. M. & Goodman, A. E. (1998).

Use of green fluorescent protein to tag and investigate gene expression in marine

bacteria. Appl Environ Microbiol 64: 2554–2559

9. Okabe, S., H. Satoh, and Y. Watanabe. 1999. In situ analysis of nitrifying

biofilms as determined by in situ hybridization and the use of microelectrodes.

Appl. Environ. Microbiol. 65:3182–3191

10. Shimkets, L. J. 1999. Intercellular signaling during fruiting-body development of

Myxococcus xanthus. Annu. Rev. Microbiol. 53:525–549

11. Gwinnett AJ, Yu S. (1995) Effect of long-term water storage on dentin bonding.

Am J Dent 8:109-111.

12. Lee SY, Greener EH, Mueller HJ. (1995) Effect of food and oral simulating fluids

on structure of adhesive composite systems. J Dent 23:1:27-35.

13. Gerzina TM, Hume WR ( 1995). Effect of hydrostatic pressure on the diffusion of

monomers through dentin in vitro. J Dent Res 74:369-373.




15. Sano H, Yoshikawa T, Periera PNR, Kanemura N, Morigami M, Tagami J,

Pashley DH. (1999) Long-term durability of dentin bonds made with a self-

etching primer, in vivo. J Dent Res 78(4):906-911.

16. Hashimoto M, Ohno H, Sano H, Kaga M, Oguchi H. (2003) In vitro degradation

of resin-dentin bonds analyzed by microtensile bond test, scanning and

transmission electron microscopy. Biomat 24:3795-3803.

90

APPENDIX A – RECIPES

Todd Hewitt Yeast Extract – THYE (1L distilled H2O)

Todd Hewitt Broth 30 g

Yeast Extract 3.0 g

Luria-Bertani – LB (1L distilled H2O)

NaCl 5.0 g

Tryptone 10 g

Yeast Extract 5.0 g

1N NaOH 1 ml

Supplemented 4X Diluted THYE (1L distilled H2O)

Todd Hewitt Broth 7.5 g

Yeast Extract 0.75 g

Hog gastric mucin (M-1778 Type III, Sigma) 0.1 g

CaCl2 1.0 mM

K2HPO4 10 mM

Sucrose 10 mM

91

APPENDIX B – GAMMA IRRADIATION

INTRODUCTION

The effectiveness and consistency of sterilization through the gamma irradiation process is

well-documented (1-3). What has been of concern though, particularly for researchers in

the field of dentistry is the potential for structural changes within tooth substrates as a result

of exposure to such high energy electromagnetic radiation (2, 4-7). In 1998, Titley et al

reported a significant decline in the shear-bond strengths of composite resin restored

samples of bovine dentine following gamma irradiation (5). In 2001, Sperandio and

colleagues repeated the shear bond strength testing of dentinal substrates following

exposure to gamma radiation. Contrary to Titley et al (1998), Sperandio et al (2001) found

shear bond strengths of dentin to be un-altered following gamma radiation. Sperandio et al

(2001) also assessed dentinal morphology (through SEM analysis) following the gamma

irradiation process (6). In support of findings by White et al (1994), as well as conclusions

made by DeWald (1997), their SEM results demonstrated a normal pattern of collagen

network organization for gamma-irradiated samples (6). Indeed, the majority of recent

studies now conclude that the exposure of mineralized tissue to moderate doses of gamma

radiation (25-30 kGy) does not significantly alter structural properties (2-3, 6-7).

Currently, a dosage level of 25 kGy (exposure time: 6 hours) is the standard for scientific

investigations using gamma irradiation for the sterilization of dental tissues (6-7). While

collagen fibrils are left unaffected at this dosage, 25 kGy is said to be capable of

inactivating most forms of microorganisms present (International Organization for

Standardization, 1995). The Gamma Irradiation services of the University of Toronto‟s

Department of Environmental Nuclear Science was employed to process collected teeth.

METHOD

In order to test the effectiveness of 25 kGy of gamma radiation sterilizing extracted teeth, a

total of 8 extracted human molars (collected in a glass container with distilled water) were

sent for processing at the University of Toronto‟s department of Environmental Nuclear

Science, Gamma Irradiation services from June 23 to August 8, 2006. For comparison

92

purposes, a total of 6 extracted molars were also stored in 1% chloramine-t solution (12%

active chlorine diluted in distilled water) for 24 hours. A total of 6 extracted molars

incubated only in double distilled water (dd H2O) were used as controls. Performed under

the sterile environment of a laminar flow hood, teeth from all experimental groups were

individually placed in 10 ml glass bottles filled (approximately 8 ml) with either THYE or

Luria-Bertani (LB) media (Recipes – see Appendix A). THYE media provides nutrients

for the optimal growth gram-positive bacteria, whereas LB media promotes the growth of

gram-negative strains. Glass bottles were then labeled and incubated at 37 degrees Celsius

for up to 6 days.

RESULTS

Results are shown in Table 1. All un-sterilized teeth stored in dd H2O demonstrated

bacterial contamination, as early as 24 hours incubation. Out of the three chloramine-T

sterilized specimens tested in THYE, one demonstrated bacterial growth following a period

of 6 days incubation. Gamma irradiated samples were the only experimental group to

demonstrate complete inactivation of all bacterial contaminants in every specimen tested.

93

Table 1. Individual glass bottles were labeled according to media type, followed by sample number.

The detection of bacterial growth was recorded as positive (+) or negative (-). Observations were made at

two separate time points for each sample; at 24 hours of incubation, and following 6 days of incubation.

(a) Teeth stored in dd H2O (control).

THYE Medium LB Medium

Tooth #1 Tooth #2 Tooth #3 Tooth #1 Tooth #2 Tooth #3

24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days

+ + + + + + + + + + + +

(b) Chloramine-T sterilization condition

THYE Medium LB Medium

Tooth #1 Tooth #2 Tooth #3 Tooth #1 Tooth #2 Tooth #3

24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days s 24 hrs r 6 days

- - - + - - - - - - - -

(c) Gamma Irradiation

THYE Medium

Tooth #1 Tooth #2 Tooth #3 Tooth #4

24 hrs r 6 days s 24 hrs s 6 days s 24 hrs r 6 days s 24 hrs r 6 days s

- - - - - - - -

LB Medium

Tooth #1 Tooth #2 Tooth #3 Tooth #4

24 hrs 6 days 24 hrs 6 days 24 hrs 6 days 24 hrs 6 days

- - - - - - - -

94

REFERENCES

1. Reid BD. (1995) Gamma processing technology: an alternative technology for

terminal sterilization of parenterals. PDA J Pharm Sci Technol 49(2):83-89.

2. DeWald JP. (1997) The use of extracted teeth for in vitro bonding studies: a

review of infection control considerations. Dent Mater 13(2):74-81.

3. Yaman A. (2001) Alternative methods of terminal sterilization for biologically

active macromolecules. Curr Opin Drug Discov Devel 4(6):760-763.

4. White JM, Goodis HE, Marshall SJ, Marshall GW. (1994) Sterilization of teeth by

gamma radiation. J Dent Res 73:9:1560-1567.

5. Titley KC, Chernecky R, Rossouw PE, Kulkarni GV. (1998) The effect of

various storage methods and media on shear-bond strengths of dental composite

resin to bovine dentine. Arch Oral Biol 43:305-311.

6. Sperandio M, Souza JB, Oliveira DT. (2001) Effect of gamma radiation on dentin

bond strength and morphology. Braz Dent J 12:3:205-208

7. Rodrigues LKA, Cury JA, dos Santos MN. (2004) The effect of gamma radiation

on enamel harness and its resistance to demineralization in vitro. J Oral Sci

46:4:215-220.

95

APPENDIX C - RESIN-DENTIN SAMPLE PREPARATION

METHOD

To ensure sterility, all sample preparations were conducted under the confines of a Laminar

Class II Biohazard Safety Cabinet (Esco Biotechnology Equipment, Singapore) located at

the Laboratory for Interfacial Research, faculty of Dentistry, University of Toronto. All

resin materials (both adhesive and composite) were kept refrigerated at 4oC until required

for use; approximately 30 minutes just prior to use, resin materials were removed and

allowed to reach room temperature.

The root of each gamma-irradiated tooth was amputated to a level 2-3 mm apical to the

cemento-enamel junction (CEJ); the pulp was extirpated from the apical opening into the

pulp chamber. Soft tissue debris was removed using of sterilized curettes/pumice. The

occlusal table was flattened by horizontal sectioning above the contour line using a water-

cooled Isomet Low-Speed Saw (Buehler, Lake Bluff, Illinois). The saw was operated

using a thin diamond wafering blade at less than 100 RPM under sterile water coolant. All

detachable components of the hand saw (including saw-arm, blade, water tray, and water

coolant) were autoclaved for 40 minutes prior to use. The flattened occlusal table was

lightly polished using a sterilized handheld sander.

Composite resin material was packed onto the occlusal surface in small increments, each

increment photo-polymerized for 40 seconds using a hand-held light curing unit (The Max,

Chaulk, Densply). The restored cavity preparation had surface area of approximately 30

mm2 in size. The Isomet Low-Speed Saw was used to section this restored tooth into 3

individual resin-dentin interfacial cross-sections each approximately 3mm x 3mm in size,

with a total surface area of approximately 54 mm2 (See Figure 1).

96

Figure 1. Illustration of the resin-dentin sample preparation procedure. Composite resin material

is bonded to the flattened occlusal surface of a tooth; this is then sectioned as shown by the dotted

lines to generate approximately 3 separate samples of the resin-dentin interfacial region, each

having an approximate total surface area of 54 mm2.

In the final step of the preparation, a small amount of composite resin material (Filtek

Z250, shade A1) was placed on top of the resin layer; a small piece of sterilized stainless

steel ligature wire (approximately 3 mm in length) was imbedded and the material cured

for 20 seconds. Once cured, the wire was secured to the top of the resin-dentin specimen;

this was required for the attachment of the specimen to glass rods for fermentation in the

chemostat-based biofilm fermentor (CBBF).

MATERIALS

The adhesive resin system used was Scotch Bond Multi-purpose (3M ESPE, London,

Ontario); applied according to manufacturer‟s instructions. Scotch Bond Multi-purpose is a

total etch 3-step adhesive system. Based on 3M‟s technical profile, the etchant is a gel

consisting of 35% phosphoric acid, the primer is composed of 40% 2-hydroxyethyl

methacrylate (HEMA) and14% polyalkenoic acid copolymer, with a pH of 3.3, and the

97

bonding resin contains 62.5% 2.2-Bis[4-(2-hydroxy-3-methacryloyloxy-

propoxy)phenyl]propane (Bis-GMA) and 37.5% HEMA. Numerous studies to date have

demonstrated the superior performance of Scotchbond Multipurpose over other

commercially available adhesive systems (1-4). Scotch bond Multi-purpose is widely used

clinically, as well as for both in vitro and in vivo investigations looking to assess various

aspects of performance at restorative margins (1-7).

The Filtek Z250 composite resin (3M ESPE, London, Ontario) shade A1 will be packed

onto the occlusal preparation in small increments using a sterilized condenser and a

standard dental plastic instrument. According to the manufacturer‟s product literature, the

resin monomers contained in Z250 consist of BisGMA and triethylene glycol

dimethacrylate (TEGDMA) as well as unspecified amounts of urethane dimethacrylate

(UDMA) and 2,2-Bis[4-(2-methacryloyloxyethoxy)-phenyl]propane (BisEMA). Its resin

matrix is filled with clusters of zirconia/silica particles ranging 0.01-3.5 microns in size;

filler loading is reported as 60% by volume. The significant degradative effect of human

saliva-derived esterase (HSDE) activity on the Z250 composite material has been

previously shown by Jaffer et al (2002).

98

REFERENCES

1. Leloup G, D‟Hoore W, Bouter D. (2001) Meta-analytical review of factors

involved in dentin adherence. J Dent Res 80:7:1605-1614.

2. De Munck J, Van Meerbeek B, Yoshida Y, Inoue S, Vargas M, Suzuki K,

Lambrechts P, Vanherle G. (2003) Four-year water degradation of total-etch

adhesives bonded to dentin. J of Dent Res 82:2:136-140.

3. Hewlett ER. (2003) Resin adhesion to enamel and dentin: a review. J Calif Dent

Assoc 31:6:469-476.



5. Gerzina TM, Hume WR. (1996) Diffusion of monomers from bonding resin-resin

composite combinations through dentine in vitro. J of Dent 24:125-128.

6. Wang Y, Spencer P. (2002) Quantifying adhesive penetration in adhesive/dentin

interface using confocal Raman microscopy. J Biomed Mater Res 59:46–55.

7. Wang Y, Spencer P. (2003) Hybridization efficiency of the adhesive/dentin

interface with wet bonding. J Dent Res 82:141–5.



23:1707-1719.

99

APPENDIX D – STERILITY ASSAYS OF SPECIMEN

PREPARATION PROCEDURE

INTRODUCTION

A series of small pilot studies were run to assess the sterility of the sectioning, bonding,

and restoration procedures outlined in Appendix C.

TOOTH SECTIONING

METHOD

A total of 4 extracted human molars were used; all were gamma irradiated prior to

sectioning. Each molar was sectioned three times. The sectioned pieces were placed in a

glass bottles containing either THYE or LB media. Two samples of the blade‟s water-

coolant (approximate volume: 3 ml) were also aseptically removed out of the water tray

(using an individually wrapped sterile Falcon transfer pipette, Becton Dickinson Labware,

NJ) before the first section and after every subsequent section made. Of the sample pairs

taken at each time point, one was added to THYE media, and the other to LB media, in

glass bottles. In total, the sectioning of one whole tooth generated 4 sectioned pieces (2 of

which were placed THYE media, and 2 in LB), and 8 water coolant samples (4 were placed

THYE media, and 4 in LB).

RESULT

Bacterial contamination was undetected in every water coolant samples and tooth sections

tested. The results of this preliminary assay suggest that the present measures being used

during the sawing procedure are sufficiently aseptic.

100

RESIN-DENTIN SPECIMEN PREPARATION

METHOD

Next, a trial run of the entire resin-dentin preparation procedure was performed under the

aseptic environment of the laminar flowhood. Resin-dentin specimens were prepared

according to the protocol outlined in Appendix C. These were then tested for sterility. In

total, 8 fully constructed resin-dentin specimens were included in the study; 6 were

incubated in THYE media, 2 in LB. More test specimens were incubated in THYE for this

experiment than LB; since the media to be used in the CBBF will be, the greatest

contamination risk concerns the growth of gram-positive bacterial strains than that of gram-

negative.

RESULTS

The results from this assay were as follows: specimens 1 through 5 demonstrated no

bacterial growth following incubation; specimen 6 was contaminated. We can therefore

conclude that the sterility measures used in the preparation of resin-dentin samples are

relative aseptic. However, the risk of possible contamination does exist. As a result,

individually prepared resin-dentin samples will be incubated in THYE for at least 6-7 days

(at 37 oC) prior to any further experimentation. After the 6-7 day period, if no bacterial

growth is observed in THYE media, the resin-dentin samples will be ascetically removed

and rinsed with sterile ddH20. These will then be used for biodegradation assays. Those in

vials demonstrating contamination will be discarded.

CHEMOSTAT-BASED BIOFILM FERMENTER

METHOD

The CBBF was run continuously for 14 days inside the laminar flowhood; during this time,

conditions remained constant and no bacterial contamination could be seen within the

vessel‟s media. After 14 days, the low-speed saw and restorative equipment were set up

underneath the flowhood along side the CBBF and the resin-dentin sample preparation

procedure described in Appedix C was conducted. After sample preparation was complete,

the extra equipment was removed and the area wiped with 70% ethanol.

101

RESULTS

On day 16 (24-hours after sample preparation), the CBBF had become contaminated. It is

likely that the contamination resulted from the interruption of laminar airflow, either by the

operation of the low-speed saw underneath the flowhood or the increased traffic in and out

of the work space. What can be concluded from this assay is that the preparation of resin-

dentin samples, conducted under the sterile confines of the laminar flowhood, should not be

carried out concurrently with the CBBF. Resin-dentin samples will therefore be prepared

prior to the set-up of the CBBF underneath the laminar flowhood.

102

APPENDIX E – SALIVARY-LIKE ESTERASES

Salivary esterases are generated from several different sources: salivary glands, human

gingival epithelia, inflammatory responses and the metabolic activities of oral

microorganisms (1-2). As a result, human saliva carries several different esterase species

(3-5). Lin et al (2005) isolated the distribution of these different esterase activities in

human saliva using gel filtration chromatography (5). Salivary esterase fractions were

separated according to molecular weight and two distinct peaks were found, one

exhibiting PCE-like activity and the other, CE-like activity (5).

Esterases are of a large family of catalytic proteins with active sites selective for

substrates containing esterified alkyl moieties (6-8). These enzymes bind to sites of ester

moieties on the polymer, and through regional electrostatic force rearrangements,

catalyze ester hydrolysis by reducing the energy required to cleave and reform bonds in

the carbonyl (9). All esterases catalyze the hydrolysis of esters, but while some act on a

more general range of substrate structures, other esterases are highly selective and bind

preferentially to certain structures more than others (3, 8). In agreement with Yourtee et

al (2001), Finer and Santerre (2003) found that psuedocholinesterase (PCE) preferentially

catalyzed the hydrolysis of TEGDMA over the Bis-GMA monomer. Cholesterol esterase

(CE) was shown to be 14 times more effective at catalyzing the hydrolysis of Bis-GMA

than PCE (3). Lin et al, (2005) confirmed these results; CE was significantly more

degradative towards Bis-GMA than PCE, while both CE and PCE degrade TEGDMA to

an equal extent (5). The observed difference in CE and PCE enzyme activity levels is

related to their different substrate reactivities (4).

CE is known to preferentially hydrolyze the breakdown of long-chain fatty acid esters of

cholesterol (10). It binds substrates containing large molecular side chains (8). In fact,

ester side-chain length is a positive variable for CE activity; substrates with longer side-

chain esters undergo higher rates of hydrolysis (4). On the other hand, PCE activity is

more selective towards low-molecular weight choline esters, such as butyrylcholine (4, 8,

11, 12). Acetylcholine esterase (ACHE), another choline ester related to PCE, also

103

preferentially hydrolyzes the TEGDMA substrate; 4.5 times more than so CE (8).

Overall, choline esters appear to be more selective towards smaller molecular substrates

(11).

Yourtee et al (2001) listed a number of physiologically relevant esterase fractions shown

to be capable of catalyzing the hydrolysis of resin monomers (8). These include

acetylcholinesterase, porcine liver esterase, cholesterol esterase, and pancreatic lipase (8).

Many of these esterases have been used as hydrolases for in vitro biodegradation assays

in the past (13-16). However in an attempt to make such in vitro investigations more

relevant to the oral cavity, subsequent attempts focused more specifically on esterase

species found in the mouth (2,4, 17-19).

Cholesterol Esterase (CE)

Cholesterol esterase (CE) is a common inflammatory cell-derived enzyme (20-22). The

detection of a foreign substance, such oral bacteria or a restorative biomaterial elicits an

inherent inflammatory immune response in the tissues of the oral cavity. As part of this

nonspecific immune reaction, gingival crevicular fluid flow increases and monocytic

cells adhere to the surface of the foreign substance (22). Given chronic stimulation of the

immune response sustained over the course of several weeks, these monocytes

differentiate into mature macrophages (22-23). These monocyte-derived macrophages

enlarge as they begin to synthesize a wide array of proteins and enzymes; CE is the

primary esterase produced (20, 22, 24-26). Results from Labow et al (1998) show a

significant increase in CE activity following the activation and differentiation of

monocytes into macrophages (26).

The presence such mononuclear phagocytic cells (monocytes and macrophages) have

been confirmed in both inflamed and healthy gingiva (27-28). It is not surprising then

that CE-like enzyme activity has been detected in human (4-5, 29); macrophage-derived

cholesterol esterases are thought to be the predominant source of this CE-like activity in

the mouth (24-26). As a result, CE is clinically relevant for biodegradation assays that

investigate the break-down of composite resin materials placed in vivo (4, 8, 15-16, 19).

104

This is particularly true in the case of resin materials used in Class V restorations that

come in close proximity to gingival margins (4, 27-28).

Psuedocholinesterase (PCE)

Cholinesterases (ChE) hydrolyze choline esters at a faster rate than they do other esters.

PCE is predominantly found in the liver, but has also been identified in human saliva (4,

11 30-31). There may be a possible diurnal pattern to salivary PCE levels (11); PCE

levels have been found to peak during early morning hours (4 am) and drop by almost

three-fold in late afternoon (4 pm) (11). This could be related to the fact that PCE has

multiple origins in human saliva (11, 32), each being regulated differently. While the

major contribution is made by the salivary glands, PCE can also be synthesized by oral

micro-organisms, leukocytes, and exists in gingival crevicular fluid (11, 32). Variations

in salivary PCE activity levels have been linked to periodontal disease (11, 30, 33) and

increased levels may result from the accumulation PCE-producing bacterial plaque in the

oral cavity (11, 32, 33). In fact, the level of salivary esterase activities (both CE and

PCE) can vary between different individuals depending factors such as on oral hygiene

and diet (11, 30). Nonetheless, the average in vivo esterase activity level has been shown

to be sufficient in hydrolyzing the synthetic matrix component of most Bis-GMA-based

resin systems (4-5, 19, 29).

Of the combined esterase fractions found in human saliva though, the most active fraction

is CE; results from Lin et al (2005) reveal that the highest amount of degradation

products were released from the Bis-GMA based Z250 composite resin (3M Inc, London,

Ontario, Canada) when incubated with the salivary CE-like fraction (5). The CE-like

fraction is significantly more degradative towards Z250 than the PCE-like fraction (4-5).

However, though highly effective at hydrolyzing resin matrices, CE enzyme activity is

also known to be very unstable, both on its own and even more so in the presence of

substrate (19). When incubated alone for 24 hours, CE‟s half-life diminishes by 55+/-

2.6% (19). In the presence of a BisGMA/TEGDMA-based resin substrate, the loss in

enzyme activity is even more drastic – by nearly 4-fold (19). PCE enzyme activity on the

105

other hand, though less sensitive to resin biodegradation than CE (4), does demonstrate

greater enzyme stability over time (19).

Human-derived Salivary Esterase (HSDE)

A crucial finding by Finer et al (2004) is that when CE and PCE coexistence together in

the presence of substrate, the activity levels of both enzymes increase (19). When CE

and PCE act in concert, the degradative effect on the resin substrate is found to be greater

than that of the sum of the individual effects by each enzyme measured on their own

(Finer et al, 2004). This may in part be due to the distinct enzymatic specificities of each

of these esterases (3, 5, 19); CE shows greater specificity towards Bis-GMA and Bis-

EMA monomers, and PCE towards TEGDMA and TEGMA components (3, 5). When

both enzymes are present, the two separate monomer components of the resin matrix

become hydrolyzed at the same time, creating access to more potential enzymatic sites

for both enzyme preferences and at a faster rate (19). There is however more to the story

since the enzyme activity of CE actually appears to become more stable in the presence

of PCE and vice versa (19). This implies that an additional synergistic effect is taking

place between the esterase species (19). Such observations have led Finer et al (2004) to

hypothesize that the co-existence of more than one enzyme species in experimental

incubation media will likely increase the overall efficiency of the resin matrix

biodegradation process (19).

What is more, multi-fraction esterase conditions are more physiologically relevant since

human saliva is composed of not one, but several different esterase species that coexist

together (4-5). There are also several other major groups of enzymes present besides the

salivary esterases that could potentially participate in syngertistic activities; they include

the carbohydrases, transferring enzymes (catalases and oxidases), proteolytic enzymes

(proteinase, peptidase), and other enzymes such as carbonic anhydrase and aldolase (2,

32). Though likely not direct participants in the resin degradation process themselves,

the co-existence of these other proteins may also potentially influence the activity levels

of salivary esterase fractions active in hydrolyzing the resin matrices (4, 29).

106

Given the likelihood of synergistic protein interactions, most recent biodegradation

studies are now using processed whole human saliva (3-5, 29), as opposed to pure CE or

PCE esterase fractions to evaluate enzymatic degradation in resin substrates (3, 13-14,

17). This is a major step towards advancing the clinical relevancy of in vitro trials.

Several investigators have identified the ability of processed whole human saliva to

significantly degrade the ester bonds of resin matrices in commercial Bis-GMA-based

composite materials, such as Z250 (3M Company, London ON) (3-4, 29). Whole saliva

samples are collected from human volunteers, homogenized, centrifuged, and then pooled

to form a dense concentrate of salivary proteins. The final product is known as human-

derived salivary esterase (HDSE). Aliquots of this concentrate can be diluted for use (as

needed) in biodegradation assays (4-5, 29). Results show that during the first 20 hours

of incubation, the stability of HSDE enzyme activity is considerably greater than that of

pure CE in solution, both in the presence and absence of Bis-GMA/TEGDMA composite

substrate (Jaffer et al, 2002). Within 25 hours of exposure to HDSE activity, both Bis-

GMA and TEGDMA monomers become completely hydrolyzed (29).

Due to time constraints in the present study though, HSDE was not used – instead, a

PCE-CE solution was made to mimic salivary levels of PCE and CE found in vivo.

Preliminary studies were conducted to determine whether this PCE-CE solution resulted

in comparable biodegradation as HSDE. The results are given in the graph below (Figure

E1).

REFERENCES

1) Lindqvist L, Nord CE, Soder PO. (1977) Origin of esterase in human whole saliva.

Enzyme 22:166-175.

2) Santerre JP, Shajii L, Leung BW. (2001) Relation of dental composite formulations



107

3) Finer Y, Santerre JP. (2003) Analysis of human saliva for esterase activity and its

association with the biodegradation of dental composite materials. J Dent Res

83:22-26.

4) Finer Y, Santerre JP. (2004) Salivary esterase activity and its association with the


5) Lin BA, Jaffer F, Duff MD, Tang YW, Santerre JP. (2005) Identifying enzyme



6) Coury AJ, Levy RJ, McMillin CR, Pathak Y, Ratner BD, Schoen FJ, Williams DF,




Inc; 1996. pg. 243-281

7) Gopferich A. (1996) Mechanisms of polymer degradation and erosion. Biomat

17:103-114.

8) Yourtee DM, Smith RE, Russo KA, Burmaster S, Cannon JM, Eick JD, Kostoryz



9) Soderholm, KJ, Mariotti A. (1999) Bis-GMA-based resins in dentistry: are they

safe? J Am Dent Assoc 130:201-209.

10) Feaster SR, Lee K, Baker N, Hui DY, Quinn DM (1996) Molecular recognition by

cholesterol esterase of active site ligands: structure-reactivity effects for inhibition

by aryl carbonates and carbamates and subsequent carbamylenzyme turnover.

Biochemistry 35:16723-16734.

11) Ryhanen R. (1983) Pseudocholinesterase activity in some human body fluids. Gen

Pharmacol 14:459-460.

12) Munksgaard EC, Freund M. (1990) Enzymatic hydrolysis of (di)methacrylates and


13) Bean TA, Zhuang WC, Tong PY, Eick JD, Yourtee DM. (1994) Effect of esterase

on methacrylates and methacrylate polymers in an enzyme simulator for

biodurability and biocompatibility testing. J Biomed Mat Res 28:59-63.

14) Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental


15) Shajii L, Santerre JP. (1999) Effect of filler content on the profile of released



16) Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental


17) Santerre JP, Shajii L, Leung BW. (2001) Relation of dental composite formulations


products. Crit Rev Oral Biol Med 12(2):136-151

18) Finer Y, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and


1793.

19) Cohn ZA, Benson B. (1965) The differentiation of mononuclear phagocytes:

Morphology, cytochemistry, and biochemistry. J Exp Med 121:153.

108

20) Anderson JM. (1993) Mechanisms of inflammation and infection with implanted

devices. Cardiovasc Pathol 2:233S-241S.

21) Labow RS, Meek E, Santerre JP. (1998) Differential synthesis of cholesterol

esterase by monocyte-derived macrophages cultured on poly(ether or ester)-based

poly(urethane)s. J Biomed Mater Res 39:469-477.

22) Anderson JM. (1993) Mechanisms of inflammation and infection with implanted

devices. Cardiovasc Pathol 2:233S-241S.

23) Labow RS, Meek E, Santerre JP. (2001) Hydrolytic degradation of poly(carbonate)-

urethances by monocyte-derived macrophages. Biomat 22:3025-3033.

24) Labow RS, Meek E, Matheson LA, Santerre JP. (2002) Human macrophage-

mediated biodegradation of polyurethanes: assessment of candidate enzyme

activities. Biomat 23:3969-3975.

25) Matheson LA, Labow RS, Santerre JP. (2002) Biodegradation of polycarbonate-

based polyurethanes by the human monocytes-derived macrophage and U937 cell

systems. Biomed Mater Res 61:505-513.

26) Kumar D, Klessig DF. (2003) High-affinity salicylic acid-binding protein 2 is

required for plant innate immunity and has salicylic acid-stimulated lipase activity.

Proceedings of the National Academy of Sciences of the United States of America.

10:26:16101-16106.

27) Lappin DF, Koulouri O, Tdver M, Hodge P, Kinane DF. (1999) Relative

proportions of mononuclear cell types in periodontal lesions analyzed by

immunohistochemistry. J Clin Periodontol 26:183-189.

28) Jaffer F, Finer Y, Santerre JP. (2002) Interactions between resin monomers and


1719.

29) Yamalik N, Ozer N, Caglayan F, Caglayan G. (1990) Determination of

pseudocholinesterase activity in the gingival crevicular fluid, saliva and serum from

patients with juvenile periodontitis and rapidly progressive periodontitis. J Dent Res

69:87-89.

30) Pershad N, Chakravarti N, Finer Y, Santerre JP. (1999) Effect of salica-like esterase

activities on micro-filled dental composites. IADR 77th

General Session, March 10-

13, Vancouver, Canada. J Dent Res (Special Issue) 78:314 (abstract #1667).

31) Chauncey H.H. (1961) Salivary enzymes. J Am Dent Assoc 63:360-368.

32) Yamalik N, Ozer N, Caglayan F, Caglayan G. (1990) Determination of

pseudocholinesterase activity in the gingival crevicular fluid, saliva and serum from

patients with juvenile periodontitis and rapidly progressive periodontitis. J Dent Res

69:87-89.

109

APPENDIX F – PCE-CE SOLUTION

A solution consisting of pseudo-choline esterase (PCE) and cholesterol esterase (CE) in

specific esterase activity levels was mixed in order to obtain a combined esterase activity

levels of 16 units/ml. Colorimetric activity assays specific to both CE and PCE were

performed to quantify CE and PCE activity levels within human saliva. Then the same

colorimetric activity assays were perform to determine the specific concentrations of PCE

and CE required to obtain a mixed PCE-CE solution having an activity level of 16

units/ml.

METHODS

CE-Like Colorimetric Activity Assay

Cholesterol esterase-like activity was measured using para-nitrophenylbutyrate (p-NPB)

(Sigma, St. Louis, MO), a reagent commonly used for the quantitative, kinetic

determination of cholesterol esterase activity in solutions (1-3). It has been shown to be a

more sensitive measure of CE-like activity (4) in comparison to the para-nitrophenol

acetate (P-NPA) substrate used in previous assays (5-6).

P-NPB solution was prepared in advance and stored at -78 oC until needed. For use, p-NPB

was defrosted in a desiccator and allowed to heat up to room temperature. In a 25 ml glass

tube, 17.75 ul of p-NPB was added to 5.5 ml acetonitrile. The solution was vortexed and

subsequently diluted in 19.5 ml of 50 mM distilled phosphate-buffer solution (D-PBS).

An Ultrospec II spectrophotometer unit (LKB Biochrom, Cambridge England) was used (1,

3) for colorimetric activity measurements.

Approximately 20 minutes prior to use, the spectrophotometer‟s Tungsten lamp WAS

turned on and the wavelength and temperature set to 410 nm and 25 oC, respectively.

PCE-SE solution (50ul) was diluted with D-PBS (950 ul) and mixed with 500 µl of P-

NPB in a 1.5 ml optical plastic cuvette. The optical density (OD) was recorded every 30

seconds for 300 seconds, with a blank cuvette containing 1000 µl of D-PBS and 500 µl of

p-NPB used as reference. The rate of absorbance/min was plotted and the resultant slope

110

was taken as the average optical density (OD)/min. One unit of PCE-CE activity was be

defined as the change in absorbance by 0.01 OD/ min (at 410 nm, pH 7.0 and 25 oC) (3),

or the release of 1 nmol of p-nitrophenol per minute according to the following chemical

reaction:

cholesterol esterase

p-nitrophenyl butyrate + H2O p-nitrophenol + butyric acid

PCE-Like Colorimetric Activity Assay

Based on the Ellman method, PCE activity was determined by measuring changes in OD

at a wavelength of 405 nm (similar to methods described above for CE), using

butyrylthiocholine iodide (BTC) as a substrate [cholinesterase (BTC) activity kit, Sigma,

Procedure No. 421] (7). Two chemical reactions are involved:

cholinesterase

Butrylthiocholine + H2O butyrate + thiocholine

Thiocholine + 5,5’-dithiobis-2-nitrobenzoic 5-thio-2-nitrobenzoic acid

BTC was prepared by mixing di-thiobisnitrobenzoic (DTNB) acid dissolved in PBS (0.25

mmol/L, pH7.2) with a 0.111 M solution of BTC [cholinesterase (BTC) activity kit,

Sigma, Procedure No. 421]. This BTC+DTNB solution (1000 µl) was added to 500 µl of

PCE-CE solution in a 1.5ml plastic cuvette and similar to colorimetric methods described

above, optical density (OD) was recorded every 30 seconds for 300 seconds.

One unit of PCE was defined as 1 mmol of 5-thio-2-nitrobenzoic acid released per

minute.

The equation used to determine PCE activity/ml was:

(change in absorbance)(1.5ml)(1000)

(13600 M-1

cm-1

)(1cm)(0.5ml)

111

REFERENCES



23:1707-1719.

2) Yang J, Koga Y, Nakano H, Yamane T. (2002) Modifying the chain-length

selectivity of the lipase from Burkholderia cepacia KWI-56 through in vitro

combinatorial mutagenesis in the substrate-binding site. Protein Eng. 15: 147–152.

3) Lin BA, Jaffer F, Duff MD, Tang YW, Santerre JP. (2005) Identifying enzyme



4) Pershad N, Chakravarti N, Finer Y, Santerre JP. (1999) Effect of salica-like esterase

activities on micro-filled dental composites. IADR 77th

General Session, March 10-

13, Vancouver, Canada. J Dent Res (Special Issue) 78:314 (abstract #1667).

5) Shajii L, Santerre JP. (1999) Effect of filler content on the profile of released





1793.

7) Ellman GL, Courtney KD, Andres V, Featherstone RM. (1961) A new and rapid

colorimetric determination of acetylcholinesterase activity. Biochem Pharmacol

7:88-95

112

APPENDIX G – HALF-LIFE EXPERIMENTS OF PCE-

CE SOLUTION

In the presence of substrate, continuous enzymatic activity results in the adsorption and

deactivation of individual enzyme molecules (1). As a result, progressive reductions in

enzyme activity levels take place over time. It was necessary to measure and maintain the

levels of enzyme activity within the PCE-CE solution relative to the surface area of

specimens being used in degradation assays at all times (1, 2).

METHODS

Cured composite resin specimens approximately 1.5 mm x 1.5 mm in size were prepared.

Placed in autoclaved glass vials, 9 composite resin specimens were incubated with HSDE

(prepared using previously published methods – Jaffer et al, 2002) and 9 specimens in

PCE-CE solution (activity level 16 units/ml). Esterase activity levels were tested through

colorimetric activity assays at 10 different incubation time points; 0, ½, 1, 2, 4, 6, 10, 24,

48, 96 hours (1). The activities of both HSDE and PCE-CE solution without the presence

of a composite resin specimen were also tested along the same time points for comparison.

RESULTS

It was found that the half-life of esterase activity in both HSDE and PCE-CE solution

decreased under the presence of a composite resin specimen (1). Based on results, a half-

life of approximately 60 hours was determined for HSDE incubated with a composite resin

specimen. PCE-CE solution demonstrated a half-life of 48 hours in the presence of a

composite resin specimen. Based on results of this pilot assay, the replenishment cycle for

resin-dentin specimens incubated in PCE-CE solution was set to 48 hours.

REFERENCES



1719.

113



1793.

APPENDIX H – COMPARATIVE BIODEGRADATION OF

HSDE AND PCE-CE SOLUTION ON COMPOSITE RESIN

SPECIMENS

INTRODUCTION

In order to determine whether the degradative effects of PCE-CE solution on composite

resin specimens was similar to that of HSDE used in previous studies (1-3), a pilot study

was performed to quantify and compare the release of Bis-HPPP from incubated

standardized composite resin specimens.

METHODS

In order to leach out a large fraction of unreacted monomers remaining within the resin

matrix following polymerization (4,5), experimental specimens were incubated in PBS

buffer solution (pH=7) for 48 hours at 37 oC (pH=7) prior to their biodegradation assays

(6). Several other studies have also employed this method (1-2, 6, 7-8) to ensure that the

bulk of degradation products released into the incubation media are indeed as a result of

chemical breakdown of the polymerized resin matrix, and not the passive elution of un-

polymerized components. After the 48 hour period, specimens were aseptically removed

and wiped with sterile gauze to remove excess moisture.

Experimental specimens (approximately 1.5 mm x 1.5 mm) were then randomly assigned

for incubated in one of three different experimental conditions:

1) incubation with HSDE

2) incubation with PCE-CE solution

3) incubation with PBS (buffer)

Experimental specimens were incubated separately in sterile glass vials for duration of 10

days. A media replenishing schedule of 48 hours was maintained in order to maintain

114

BisHPPP Peak Area Under Curve Over Time

0

10000

20000

30000

40000

50000

60000

70000

48 96 144 192 240

Time (hours)

Pe

ak

Are

a (

Vo

lt x

Se

c)

PBS

PCE+CE

HSDE

Figure 2. Graph of Bis-HPPP Peak Area under curve for HSDE, PCE-CE media, and PBS incubation

media at 48-hour time intervals over the course of a 10 day incubation period.

salivary-esterase like activity levels at all times during the experiment. At each

replenishment cycle, the total amount of incubation solution within each vial was extracted

aseptically and replenished with fresh incubation solution (as assigned). The extracted

incubation solutions were pooled individually for each specimen and stored at 4 oC until

the given experimental time point has been reached.

HPLC in combination with ultraviolet (UV) spectroscopy was used to isolate and obtain the

peak area under curve for the Bis-HPPP degradation product (1, 6, 9). Analysis of

incubation media was performed at every 48-hour time interval.

RESULTS

The Bis-HPPP peak area under curve recorded for specimens incubated in HSDE, PCE-CE

solution, and PBS for a duration of 10 days is shown in Figure 2.

115

Composite resin specimen incubation in both PCE-CE media and HSDE resulted in

significantly higher peak area under curve values than PBS at all time points tested

(p>0.05). It also shown that the kinetic Bis-HPPP-release profile of PCE-CE incubated

cured composite resin specimens was comparable to that of cured composite resin

specimens incubated in HSDE (p<0.05).

REFERENCES



23:1707-1719.


psuedocholinesterase on the biodegradation of dental composites. Biomat

25:1787-1793.




4. Sanders B, Baudach S, Davy KWM, Braden M, Clarke R. (1997) Synthesis of

Bis-GMA derivatives, properties of their polymers and composites. J. Mater Sci:

Mat Med 8:39-44.

5. Santerre JP, Shajii L, Leung BW. (2001) Relation of dental composite

formulations to their degradation and the release of hydrolyzed polymeric-resin-

derived products. Crit Rev Oral Biol Med 12(2):136-151.



7. Ferracane JL, Condon JR. (1990) Rate of elution of leachable components from

composites. Dent Mater 6:282-287.

8. Finer Y, Santerre JP. (2004b) Salivary esterase activity and its association with

the biodegradation of dental composites. J Dent Res 83:1:22-26.



116

APPENDIX I – HIGH-PERFORMANCE LIQUID

CHROMATOGRAPHY (HPLC)

Similar to previous biodegradation studies (1-3) HPLC in combination with ultraviolet

(UV) spectroscopy and mass spectrometry (HLPC/MS) was used to isolate and quantify

degradation products within the cumulative incubation media for each sample. Collected

media was centrifuged at 3000 RPM to separate high molecular weight proteins from resin

degradation products. These were then be filtered using a Millipore UF-15 filter device

(Ultrafree – CL, UFC4LCCOO 500 NMWL, Millipore, Bedford, MA) to remove proteins

having high molecular weights (greater than 10 KD) (2). A filtered incubation solution for

each sample group was kept refrigerated at 4 oC until ready for HPLC analysis (3).

A WatersTM

HPLC system (Waters, Mississauga, ON) was used for the chromatographic

separation of the degradation products. The unit consists of a 600E multi-solvent delivery

system and a 996 photodiode array (PDA) detector coupled with a Millennium

chromatography manager, version 2.15. A Phenomenex Luna 5um C18 4.6 X 250

(Phenomenex, Torrance, CA) column was used to for the isolation of the Bis-HPPP

byproduct.

ISOCRATIC METHOD

Separation of the biodegradation products was achieved through an isocratic method of

60% methanol and 40 % 2 mM solution of ammonium acetate (pH=7); run time of 30

minutes. Using this method, it was found that standard Bis-HPPP was released at

approximately the 11.02 minute mark.

A pilot study was conducted to determine whether the average peak area detected for a

standard Bis-HPPP solution was altered under an isocratic HPLC method – as compared to

the established gradient method used in previous studies (4). Results are shown in Figure 3

- no significant differences in average area under curve were found (p>0.05).

117

Gradient vs. Isocratic Method

30600

30800

31000

31200

31400

31600

31800

32000

32200

Gradient Method Isocratic Method

Method

Avera

ge A

rea u

nd

er

Peak (

mV

x S

ec.)

In order to quantify incremental amounts of Bis-HPPP released, the area under

chromatogram peaks were converted to mass (ug)/cm2 of composite surface area using a

standardized calibration curve (3, 5). HPLC chromatograms were reported at a UV

wavelength of 215nm. Figure 4 gives the standard curve for the Bis-HPPP byproduct.

Figure 3. A comparison between gradient and isocratic HPLC methods for isolating

and quantifying the Bis-HPPP byproduct in standard media. No significant

difference in average area under peak was found (p>0.05)

118

Standard Curve for BisHPPP Dissolved in Control Media

y = 79291x - 1769.2

R2 = 0.9953

-5000

0

5000

10000

15000

20000

25000

30000

35000

40000

0 0.1 0.2 0.3 0.4 0.5 0.6

Average Area under Peak (V x Sec)

Am

ou

nt

(m g

ram

)

Figure 4. Standard curve for the Bis-HPPP degradation product.

REFERENCES








23:1707-1719.

4. Shokati, B (2007). Effect of salivary esterase on integrity and fracture toughness

of resin-dentin interface. MSc. Thesis, University of Toronto, Faculty of

Dentistry

5. Finer Y, Santerre JP. (2004a) The influence of resin chemistry on a dental


119

Effect of Freeze Drying of Concentration of BisHPPP Standard

(10E-6) Dissolved in PBS

0.37

0.38

0.39

0.4

0.41

0.42

0.43

0.44

Control BisHPPP Standard Freeze dried BisHPPP Standard

Are

a u

nd

er

Peak (

V x

Sec.)

APPENDIX J – INCUBATION MEDIA FREEZE DRYING

PROCEDURE

INTRODUCTION

Prior to HPLC analysis collected incubation media were freeze dried and reconstituted to

obtain a solution of higher concentrations. A pilot study was run using Bi-HPPP standard

solution to determine whether any of the Bis-HPPP product was lost during the freeze-

drying procedure.

METHODS

Standard Bis-HPPP [10-6

] in volumes of 0.2ml was pipetted into centrifuged tubes and spun

in the mini-centrifuge for 2-3 seconds. Controls were refrigerated. Liquid nitrogen was

used to flash freeze all experimental media. Experimental media specimens these were

then connected to the freeze-drying apparatus. All were freeze dried for 3 hours and

subsequently reconsistuted in 0.2 ml of PBS. Controls were allowed to warm back up to

room temperature. HPLC was used to analyze peak area of Bis-HPPP product in both

control and experimental media specimens.

Figure 5. Comparison of the area under peak determined for the Bis-HPPP product

120

Effect of Freeze Drying of Concentration of BisHPPP Standard

(10E-6) Dissolved in PBS

0

0.005

0.01

0.015

0.02

0.025

0.03

0.035

0.04

0.045

Control 30 day post-incubation

media - 1:1 dilution

Freeze dried 30 day post-

incubation media - 2:1 dilution

Are

a u

nd

er

Peak (

V x

Sec.)

detected from control and experimentally freeze-dried media specimens. No significant

different was found (p>0.05).

RESULTS

The area under peak value for freeze dried media specimens was not significantly different

from that of control specimens (p>0.05) (Figure 5). Therefore it was concluded that no

Bis-HPPP was lost in the freeze-drying process.

A second experimental run was done to see if the peak area under curve for Bis-HPPP

would double in value when freeze-dried specimens were reconstituted in 0.1 ml of PBS

(2:1). Results of this experiment are shown in Figure 6 – it was found that the peak area

increased proportionally to the dilution factor.

Figure 6. Area under peak for the Bis-HPPP product detected from reconstituted solution

in a 1:1 dilution compared to a 2:1 dilution.

121

APPENDIX K – MICROBIOLOGY TECHNIQUES

STOCK CULTURE

The stock s. mutans NG8 culture was generously donated by Celine Levesque from the

Faculty of Dentistry, Department of Microbiology. The NG8 strain was stored in 30%

glycerol in cryogenic vials at minus 70 degrees Celsius. The strain was cultured on

THYE (Appendix 1) agar plates and incubated overnight at 37 degrees Celsius.

Following overnight incubation, it was refrigerated (4 degrees Celsius) until use.

CBBF INOCULATION

For inoculation of the CBBF, colonies of cultured NG8 were smeared off of the agar

plate and diluted in THYE – this was incubated overnight at 37 degrees Celsius. To

inoculate the CBBF, 10% of the working volume of the chemostat vessel was used;

therefore 40 ml of the overnight culture was hermetically introduced into the vessel

through ports. The CBBF was run for 2 full days to allow for biofilm formation, prior to

insertion of resin-dentin specimens. Growth was monitored through use of a

spectrophotometer to measure the optical density at 675 nm (1)

VIABLE CELL COUNT PROTOCOL

The number of colony-forming units (CFUs) per milliliter (ml) of culture is determined

using standard spreading techniques at various optical densities. Liquid culture (10 mL)

is centrifuged for 10 mins at 6000 rpm to pellet the microorganisms. The supernatant is

poured off and the pellet (containing the microorganisms) is re-suspended in 10 mL of

phosphate buffer (PBS). This liquid is then homogenized for 30 seconds at 20,500 rpm

using a probe homogenizer. This liquid is plated on THYE agar and incubated overnight

at 37 degrees Celsius.

Viable cell counts are based on the premise that the numbers of colonies which ensue on

each agar plate are equal to the number of viable bacteria in the sample that was spread

over the agar. Using this viable cell count protocol, liquid cultures can be diluted or

concentrated to achieve the desired CFU/ml.

122

For the purposes of this study, viable cell counts (CFU) were required not of the liquid

culture within the CBBF, but of the biofilm growing on the hard surfaces. The CFU

value needed to be compared to the surface area from which the biofilm was removed,

according to the following equation:

CFU/cm2 = (number of cells)(1/dilution)(1/surface area cm

2)(scraped volume)

Upon removal of resin-dentin specimens from the CBBF, biofilm was scraped off of the

top surface and then re-suspended in a centrifuge tube containing 5 mL PBS. The total

volume of the scraped biofilm and the 5ml PBS in the referred to as „scraped volume‟.

REFERENCES

1. Lobo MM, Goncalves RB, Ambrosano GMB, Pimenta LAF. (2005) Chemical or

microbial models of secondary caries development around different dental restorative

materials. J Biomed Mater Res Part B: Appl Biomater 74B:725-731.

123

APPENDIX L – CHEMOSTAT-BASED BIOFILM

FERMENTOR SET-UP

This laboratory model consisted of a sealed glass vessel containing multiple inlets

allowing for the flow of gas, media, as well as a sampling port. Inside the vessel, resin-

dentin specimens were suspended; liquid culture was stirred at a constant speed to

consistently smear incoming medium over surfaces; forming and maintaining a biofilm at

constant depth.

The CBBF model system was equipped with pH and temperature controllers set to pH 7

and 37 degrees Celsuis having sensitivities of +/-0.1 pH and +/-0.1 degrees Celsuis,

respectively. A solution of 1M NaOH was used to maintain neutral pH within the vessel.

Prior to use, the chemostat containing 400 ml of the THYE-based growth medium were

autoclaved for 30 min at 121 degrees Celsuis. A magnetic stirrer was used to

continuously stir media within the vessel and create low levels of physical stress within

the vessel environment.

In the interests of reproducibility, a mono-strain culture of streptococcus mutans NG8

was used in this study (1,2). Directly following inoculation, the CBBF was operated in

batch mode for the first 20-30 hours; no media was pumping in or out of the vessel

during this period. After this period, the incoming/outgoing pumps were turned on. In

order to establish a steady state, the reactor was run in continuous flow for 6-7

generations (pot volumes) (3). Steady state in the CBBF was determined based on cell

density. The microbial culture in the vessel was periodically tested for contamination by

plating on agar plates.

Flow rate within the vessel was calculated based on calibrated pumps operating at low

flow rates - by counting the drops per unit time and measuring the volume of the drops.

A rate of 0.72 L/day (2) was established to correspond to the mean resting flow rate of

saliva in humans (4-6).

124

Flow breakers in the tubing line of both incoming and outgoing media were used to

separate the liquid flow line in order to prevent the back contamination of growth media.

To minimize chances of microbial contamination, the reactor setup in assembled in a

laminar flow hood. Also, all components of the setup were sterilized using 70% ethanol

before and after connecting the components. Before autoclaving (121 degrees Celsius for

20 minutes), the ends of the tubing and the connectors were covered with aluminum foil.

Typically, growth media designed to culture microorganisms in planktonic form are

highly concentrated in nutrients. In order to maintain nutrient concentrations relevant to

in vivo conditions, THYE media was diluted four times (4X). For S. mutans biofilm

growth, the modulation of carbohydrate availability is the central psychological control

point (3). Li et al (2001) previously determined that 4X diluted THYE solution is optimal

for growth of S. mutans biofilm within the CBBF (7).

Mucin is a major glycoprotein found within human saliva and is commonly added in the

form of hog gastric music to supplement S. mutans culture media (8). It has been found

that the presence of mucin increases the growth rate of monoculture S. mutans biofilm

under sugar-limited conditions (9). The addition sucrose is also required for effective

biofiolm formation within monocultures of oral streptococci (3). Sucrose supplemented

biofilms appear to colonize the substratum more rapidly (8).

REFERENCES

1. Auschill TM, Arweiler NB, Netuschil L, Brecx M, Reich E, Sculean A, Artweiler

NB. (2001) Spatial distribution of vital and dead microorganisms in dental biofilms.


2. Hope CK, Clements D, Wilson M. (2002) Determining the spatial distribution of


profiling. J Applied Microbiol 93:448-455

3. Burne RA and Chen YM. (1998) The use of continuous flow bioreactors to explore

gene expression and physiology of suspended and adherent populations of oral

streptococci. Methods in Cell Science 20: 181-190.

125

4. Lamb JF, Ingram CG, Johnston IA, Pitman RM. (1991) Gastrointestinal system. In

Essentials of Physiology pp. 91-115. Oxford: Blackwell Scientific Publications.

5. Guyton AC, Hall JE. (1992) Secretary functions of the alimentary tract. In Human

Physiology and Mechanisms of Disease ed. Schmitt, W. pp. 524-536. Philadelphia:

Saunders.




7. Li YH, Lau PCY, Lee JH, Ellen RP, Cvitkovitch DG. (2001) Natural genetic

transformation of Streptococcus mutans growing in biofilms. J of Bacteriology

183:4:897-908.




9. Renye JA, Piggot PJ, Daneo-Moore L, Buttaro BA. (2004) Persistence of

Streptococcus mutans in stationary-phase batch cultures and biofilms. Appl Environ

Microbio 70(10):6181-6187.

126

APPENDIX M – LIVE/DEAD BACLIGHT BACTERIAL

VIABILITY FLOURESCENT STAINING

Live/Dead Baclight Bacterial Viability Kit (Invitrogen - Molecular Probes, Eugene

Oregon, USA L7012/ Lot: 41803A) allows bacterial cells to be distinguished according

to cytoplasmic membrane permeability (1-3). It contains two dyes; STYO9 (excitation

488 nm and emission 525 nm) penetrates both viable and nonviable bacteria, while

propidium iodide (excitation 488 nm and emission 560 nm) penetrates bacteria with

damaged plasma membranes only, quenching the green SYTO9 fluorescence. Following

excitation, dead cells interacting with Propidium Iodide are visualized in the red

wavelength, while live cells stained by Syto9 are visible in the green.

The Baclight stain kit was prepared according to the manufacturer‟s instructions; 1 ml of

Baclight stain solution was prepared by mixing1.5 l of Component A (SYTO9) and 1.5

l of Component B (Propidium Iodide) in distilled water. One drop was applied to the

surface of each resin-dentin specimen under investigation and these were allowed to

develop in the dark for 15 minutes, at room temperature.

Fluorescence intensity of CLSM images captured in the present study could not be

exclusively attributed to the presence of bacterial cells as both dentin and composite resin

materials were found to interact with the fluorescent dyes used. As a result, while it can

be assumed that live and dead bacterial cells were differentiated by SYTO9 and PI,

respectively, the matter of cell vitality along sections within the marginal interface could

127

not be addressed by this investigation. Other investigations though have demonstrated

the capacity of CLSM analysis combined with SYTO9/PI fluorescent staining to assess

biofilm vitality (4)

OTHER FLOURESCENT STAINS

Preliminary attempts at finding the most suitable fluorescent stain for this experimental

set-up were made. Stains displaying a signal in the ultra-violet (blue) spectral region,

such as Rhodamine B, are known to produces a strong signal. However such short

wavelength excited fluorochromes are often insoluble in water and are prone to photo-

bleaching (5). For this reason, fluorescent dyes excited by longer wavelength energy and

displaying emissions of longer wavelength light (specifically, green and red) were

targeted.

Commonly used long wavelength energy fluorescent stains such as flourescien diacetate

and ethidium bromide were explored. Ethidium bromide is a red fluorescent nucleic acid

stain that permeates only cells with damaged cell membranes. Fluorescein diacetate

penetrates all cells but is non-fluorescent until the green fluorescein moiety is freed by

intracellular esterases. A disadvantage of fluorescein is that it is not retained in the cells

for a prolonged amount of time; all analysis would have had to be performed within 15

min of staining.

128

REFERENCES


(2001) A new in vitro model for the study of microbial microleakage around

dental restorations : a preliminary qualitative evaluation. Inter Endo J 34 :547-

553.

2. Decker EM. (2001) The ability of direct fluorescence-based, two colour assays to

detect different physiological states of oral streptococci. Lett Appl Microbiol

33:188-192.

3. Sharma A, Inagaki S, Sigurdson W, Kuramitsu K (2005) Synergy between

Tannerella forsythia and Fusobacterium nucleatum in biofilm formation. Oral

Microbiol Immunol 20:39-42

4. 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

5. D‟Alpino PHP, Pereira JC, Svizero NR, Rueggeberg FA, Pashley DH. (2006) Use

of fluorescent compounds in assessing bonded resin-based restorations: A

literature review. J Dent 34:623-634.

129

APPENDIX N – CONFOCAL SCANNING LASER

MICROSCOPY

CLSM analysis allows for a 3-dimensional (3D) assessment of any potential microleakage

taking place at the resin-dentin interface. Following CBBF suspension, resin-dentin

specimens were individually characterized for bacterial microleakage through CLSM

analysis combined with a standard fluorescent staining technique. Located at the Advanced

Optical Microscopy Facility (AOMF) of Princess Margaret Hospoital/Ontario Cancer

Institute, a two-photon confocal microscope (Zeiss Axiovert 135M), equipped with a 63 X

1.4 NA water immersion lens (Zeiss, Carl Zeiss Ltd, Welwyn Garden City, Herts, UK)

was used.

An aqueous system is important for CLSM; therefore in the present study, a wet system

was employed. Resin-dentin specimens were not allowed to dry following staining prior to

CLSM analysis. Therefore we can be sure that no artificial shrinking of the plaque

architecture both on the surface as well as at penetrating depths within the interface took

place. Because auto-fluorescence of both the resin composite and dentin portions of the

resin-dentin specimen was found during preliminary analysis of resin-dentin specimens,

Detector Gain and Amplitude settings were adjusted prior to image capture. The minimum

threshold at which no further autoflourescence was detected from resin-dentin specimens

under the CLSM configuration used as a standardized setting for the remainder of the

investigation.

For each specimen, 6 sequential regions of interest (ROI) on one side were imaged through

a Z-stack series collected at 2 um intervals for a total depth of 50 um. Each ROI had X=1

mm, Y= 30 um, and Z= 50 um with a volume of 1.5x103 mm

3. An argon laser, at 488nm,

was used as the excitation source for the fluorescent probe. A 530/30 Band Pass (BP) filter

was utilized for SYTO9 and FITC-ConA and a 605 Long Pass (LP) filter was utilized for

PI. In order to facilitate comparisons between different specimens, the laser power and

pinhole settings were kept constant for all captured images.

130

IMAGEJ SOFTWARE

IMAGEJ is powerful image-analysis software available from the NIH with accessory

plug-ins contributed from many sources. Obtaining quantitative data from images is a

complex process, which can be subject to a high degree of subjectivity; computational

filtering/processing techniques can standardize variables. For this investigation,

background subtraction was applied to each captured image in order to remove low-

intensity noise while leaving higher intensity bacterial particles or clusters untouched.

Automatic thresholding using maximum entropy threshold was applied as a non-biased

thresholding method to convert the image to a binary image, permitting particle analysis.

Gamma multiplication was applied to highlight bright areas and to suppress the darkest

areas; gamma values were optimized for each colour channel (1.1 for green and 1.3 for

red). Overall, these steps facilitated maximum retrieval of stained bacteria and minimize

extraneous noise from background substrate-stain interactions.

No attempts were made to separate bacteria in close proximity, making evaluation of the

number of bacteria present unrealistic; however, quantitative data regarding the area

occupied by stained bacteria gives a valuable insight into the degree of bacterial

colonization.

IMAGEJ CELL COUNTER APPLICATION

This plug-in allows for the quantification of cell within a CLSM Z-stack image by

manual clicking of the mouse over each cell. A colored number corresponding to the

131

type of cell being counted is displayed on each image once „clicked‟. Results are given in

a table displaying cell counts per depth interval (slice) as well as total z-stack counts.

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