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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.
6
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.
8
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
2.6 REFERENCES
1. 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.
2. Van Meerbeek B, Perdigao J, Lambrechts P, Vanherle G. (1998) The clinical
performance of adhesives. J Dent 26:1-20.
3. Santini A, Mitchell S. (1998) Microleakage of composite restorations bonded with
three new dentin bonding agents. J of Esthet Dent 10:6:296-304.
4. Murray PE, Hafez AA, Smith AJ, Cox CF. (2002). Bacterial microleakage and pulp
inflammation associated with various restorative materials. Dent Mat 18:470-478.
5. 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.
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 84:4:355-359.
7. Phillips RW. Phillip‟s Science of Dental Materials: 10th
Ed. Philadelphia PA: W.B.
Saunders Co.; 1996. pg 243-280.
8. 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.
9. Mosner N, Salz U. (2001) New developments of polymeric dental composites. Prog
Polym Sci 26:535-576.
31
10. 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.
11. Hewlett ER. (2003) Resin adhesion to enamel and dentin: a review. J Calif Dent
Assoc 31:6:469-476.
12. Bouillaguet S. (2004) Biological risks of resin-based materials to the dentin-pulp
complex. Crit Rev Oral Biol Med 15(1):47-60.
13. Jacobsen T, Soderholm KJ. (1995) Some effects of water on dentin bonding. Dent
Mater 11:132-136.
14. van Noort R. Introduction to Dental Materials. Papel, Spain: Times Mirror
International Publishers Limited; 1994. pgs. 3-145.
15. Marshall GW, Marshall SJ, Kinney JH, Balooch M. (1997) The dentin substrate:
structure and properties related to bonding. J Dent 25(6): 441-458.
16. Love RM, Jenkinson HF. (2002) Invasion of dentinal tubules by oral bacteria. Crit
Rev Oral Biol Med 13(2): 171-183.
17. Hayakawa T, Kikutake K, Nemoto K. (1998) Efficacy of self-etching primers
containing carboxylic acid monomers on the adhesion between composite resin and
dentin. J Oral Sci 40:9-16.
18. Kugel G, Ferrari M. (2000) The science of bonding: from first to sixth generation. J
Am Dent Assoc 131:20S-25S
19. Nakabayashi N, Kojima K, Masuhara E. (1982) The promotion of adhesion by
infiltration of monomers into tooth substrates. J Biomed Mater Res 16:265-273.
32
20. Nakabayashi N, Takarada K. (1992) Effect of HEMA on bonding to dentin. Dent
Mater 8:125-130.
21. 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.
22. 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.
23. Spencer P, Wang Y, Walker MP, Wieliczka DM, Swafford JR. (2000) Interfacial
chemistry of the dentin/adhesive bond. J Dent Res 79:7:1458-1463.
24. Doi J, Itota T, Torii Y, Nakabo S, Yoshiyama M. (2004) Effect of 2-hydroxyethyl
methacrylate pre-treatment on micro-tensile bond strength of resin composite to
demineralized dentin. J Oral Rehab 31: 1061–1067.
25. Hayakawa T, Nemoto K, Horie K. (1995) Adhesion of composite to polished dentin
retaining its smear layer. Dent Mater 11:218-222.
26. Giachetti L, Bertini F, Russo DS. (2004) Investigation into the nature of dentin
resin tags: A scanning electron microscopic morphological analysis of
demineralized bonded dentin. J Prosthet Dent 92:233-238.
27. Titley K, Chernecky R, Chan A, Smith D. (1995) The composition and ultra-
structure of resin tags in etched dentin. Am J Dent 8:224-30.
33
28. Armstrong SR, Boyer DB, Keller JC, Park JB. (1998) Effect of hybrid layer on
fracture toughness of adhesively bonded dentin–resin composite joint. Dent Mater
14:91–98.
29. Yang B, Adelung R, Ludwig K. (2005) Effect of structural change of collagen
sibrils on the durability of dentin bonding. Biomaterials 26:5021-5031.
30. Buonocore MG, Matsui A, Gwinnett AJ. (1968) Penetration of resin dental
materials into enamel surfaces with reference to bonding. Arch Oral Biol 13(1):61-
70.
31. Eick JD, Robinson SJ, Byerley TJ, Chappell RP, Spencer P, Chappelow CC. (1995)
Scanning transmission electron microscopy/energy-dispersive spectroscopy analysis
of the dentin adhesive interface using a labeled 2-hydroxythelmethacrylate
analogue. J Dent Res 74:6:1246-1252.
32. Spencer P, Swafford JR. (1999) Unprotected protein at the dentin- adhesive
interface. Quintessence Int 30:501–7.
33. Spencer P, Swafford JR. (1999) Unprotected protein at the dentin- adhesive
interface. Quintessence Int 30:501–7.
34. Wang Y, Spencer P. (2004) Effect of acid etiching time and technique on interfacial
characteristics of the adhesive-bond using differential staining. Eur J Oral Sci
112:293-299.
35. Wang Y, Spencer P. (2002) Quantifying adhesive penetration in adhesive/dentin
interface using confocal Raman microscopy. J Biomed Mater Res 59:46–55.
34
36. Wang Y, Spencer P. (2003) Hybridization efficiency of the adhesive/dentin
interface with wet bonding. J Dent Res 82:141–5.
37. Hashimoto M, Ohno H, Kaga M, Endo K, Sano H, Oguchi H. (2000) In vivo
degradation of resin-dentin bonds in humans over 1 to 3 years. J Dent Res 79:1385-
1391.
38. Hashimoto M,Ohno H, Kaga M, Sano H, Endo K, Oguchi H. (2002) The extent to
which resin can infiltrate dentin by acetone-based adhesives. J Dent Res 81:74–8.
39. 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.
40. Armstrong SR, Keller JC, Boyer DB. (2001) Mode of failure in the dentin-adhesive
resin-resin composite bonded joint as determined by strength-based [mTBS] and
fracture-based [CNSB] mechanical testing. Dent Mater 17:201-210.
41. Kaaden C, Schmalz G, Powers JM. (2003) Morphological characterization of the
resin-dentin interface in primary teeth. Clin Oral Invest 7: 235-240.
42. 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.
43. 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.
44. Iwami Y, Yamamoto H, Ebisu S. (2000) A new electrical method for detecting
marginal leakage of in vitro resin restorations. J of Dent 28:241-247.
35
45. Piwowarczyk A, Lauer HC, Sorensen JA. (2005) Microleakage of various
cementing agents for full cast crowns. Dent Mater 21:445-453.
46. Reis AF, Giannini M, Pereira PNR. (2007) Long-term TEM analysis of the
nanoleakage patterns in resin-dentin interfaces produced by different bonding
strategies. Dent Mater 23: 1164-1172.
47. Yuan Y, Shimada Y, Ichinose S, Tagami J. (2007) Qualitative analysis of adhesive
interface nanoleakage using FE-SEM/EDS. Dent Mater 23:561-569.
48. Burrow MF, Satoh M, Tagami J. (1996) Dentin bond durability after three years
using a dentin bonding agent with and without priming. Dent Mater 12:302–7.
49. 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.
50. 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.
51. 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
52. Gwinnett AJ, Yu S. (1995) Effect of long-term water storage on dentin bonding. Am
J Dent 8:109-111.
53. Burrow MF, Inokoshi S, Tagami J. (1999) Water sorption of several bonding resins.
Am J Dent 12:295-298.
36
54. Tay FR, Pashley DH. (2003) Water treeing – a potential mechanism for degradation
of dentin adhesives. Am J Dent 13:6-12.
55. Tay FR, Pashley DH, Yoshiyama M. (2002) Two modes of nanoleakage expression
in single-step adhesives. J Dent Res 81:472-476.
56. Okuda M, Pereira PN, Nakajima M, Tagami J, Pashley DH. (2002). Long-term
durability of resin dentin interface: nanoleakage vs. microtensile bond strength.
Oper Dent 27:289-296
57. 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.
58. 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.
59. Ferrari M, Mannocci F, Cagidiaco MC, Kugel G. (1997) Short-term assessment of
leakage of class V composite restorations placed in vivo. Clin Oral Invest 1:61-64.
60. Pioch T, Stotz S, Staehle H, Duschner H. (1997) Applications of confocal laser
scanning microscopy to dental bonding. Adv Dent Res 11:453-461.
61. 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
37
62. Gopferich A. (1996) Mechanisms of polymer degradation and erosion. Biomat
17:103-114.
63. Hashimoto M, Ohno H, Kaga M. (2001) Resin-tooth adhesive interfaces after long-
term function. Am J Dent 14:211-214.
64. Tanaka J, Ishikawa K, Yatani H, Yamashita A, Suzuki K. (1999) Correlation of
dentin bond durability with water absorption of bonding layer. Dent Mater 18: 11-
18.
65. 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.
66. 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.
67. 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.
68. Munksgaard EC, Freund M. (1990) Enzymatic hydrolysis of (di)methacrylates and
their polymers. Scand J Dent Res 98:351-355.
69. Jaffer F, Finer Y, Santerre JP. (2002) Interactions between resin monomers and
commercial composite resins with human saliva derived esterases. Biomat 23:1707-
1719.
38
70. Finer Y, Santerre JP. (2003) Biodegradation of a dental composite by esterases –
dependence on enzyme concentration and specificity. J Biomater Sci Polym Ed
14:837-849.
71. Finer Y, Santerre JP. (2004a) The influence of resin chemistry on a dental
composite‟s biodegradation. J Biomed Mater Res 69A:233-246.
72. 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.
73. Finer Y, Santerre JP. (2004b) Salivary esterase activity and its association with the
biodegradation of dental composites. J Dent Res 83:1:22-26.
74. Leone CW, Oppenheim FG. (2001) Physical and Chemical Aspects of Saliva as
Indicators of Risk for Dental Caries in Humans. J Dent Educ 65:10:1054-1062.
75. Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
76. 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.
77. Finer Y, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and
psuedocholinesterase on the biodegradation of dental composites. Biomat 25:1787-
1793.
78. 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.
39
79. Ingman T, Sorsa T, Lindy V, Koski H, Konttainen YT. (1994) Multiple forms of
gelatinases/type IV collagenases in saliva and gingival circular fluid pf periodontitis
patients. J Clin Periodontol 21:26-31.
80. Tjaderhane L, Larjava H, Sorsa T, UItto VJ, Larmas M, Salo T. (1998) The
activation and function of host matrix metalloproteinases in dentin matrix
breakdown in caries lesions. J Dent Res 77(8): 1622-1629.
81. 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.
82. van Strijp AJ, Jansen DC, DeGroot J, Ten Cate JM, Everts V. (2003) Host-derived
proteinases and degradation of dentine collagen in situ. Caries Res 37:58-65.
83. Arola D, Reprogel RK. (2005) Effects of aging on the mechanical behavior of
human dentin. Biomaterials 26:4051-4061.
84. Nishitani Y, Yoshiyama M, Wadgaonkar B, Breschi L, Mannello N, Mazzoni A,
Carvalho R, Tjaderhane L, Tay FR, Pashley DH. (2006) Activation of
gelatinolytic/collagenolytic activity in dentin by self-etching adhesives. Eur J Oral
Sci 114: 160-166.
85. 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.
86. Mazzoni A, Pashley DH, Tay FR, Gobbi P, Orsini G, Ruggeri A, Carrilho M,
Tjaderhane L, Di Lenarda R, Breschi L. (2008) Immunohistochemical identification
40
of MMP-2 and MMP-9 in human dentin: Correlative FEI-SEM/TEM analysis. J
Biomed Mater Res, March 11, 2008.
87. 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.
88. Yoshida K, Kamada K, Atsuta M. (2001) Effects of two silane coupling agents, a
bonding agent, and thermal cycling on the bond strength of a CAD/CAM composite
material cemeneted with two resin luting agents. J Prosthet Dent 85:184-189.
89. Sano H, Takatsu T, Ciucchi B, Carvalho RM. (1994) Relationship between surface
area for adhesion and tensile bond strength – evaluation of a microtensile bond test.
Dent Mater 10:236-240.
90. Inoue S, Vargas MA, Abe Y, Yoshida Y, Lambrechts P, Vanherle G. (2001)
Microtenshile bond strength of eleven contemporary adhesives to dentin. J Biomed
Mater Res 3:237-245.
91. Neme AL, Evans DB, Mazson BB. (2000) Evaluation of dental adhesive systems
with amalgam and resin composite restorations: comparison of microleakage and
bond strength results. Oper Dent 25:6:512-519.
92. Piemjai M, Watanabe A, Iwasaki Y, Nakabayashi N. (2004) Effect of remaining
demineralized dentine on dental microleakage accessed by a dye penetration: how
to inhibit microleakage? J Dent 32: 495-501.
93. 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.
41
94. Zivkovic S, Bojovic S, Pavlica D. (2001) Bacterial penetration of restored cavities.
Oral Surg Oral Med Oral Path Oral Radiol Endod 91:353-358.
95. Deliperi S, Bardwell DN, Wegley C. (2007) Restoration interface microleakage
using one total-etch and three self-etch adhesives. Oper Dent 32(2): 179-184.
96. Taylor MJ, Lynch R. (1992) Microleakage. J of Dent 20:230-235.
97. Totiam P, Gonzalez-Cabezas C, Fontana MR, Zero DT. (2007) A new in vitro
model to study the relationship of gap size and secondary caries. Caries Res
41:467-473.
98. Alani AH, Toh CG. (1997) Detection of microleakage around dental restorations: a
review. Oper Dent 22:173-185.
99. Li YH, Lau PCY, Svensater G, Ellen RP, Cvitkovich DG (2002) A novel two-
component regulatory system involved in biofilms formation and acid resistance in
Streptoccus mutans. J Bacteriol 184:6333-6342.
100. Newman HN. (1974) Microbial films in nature. Microbios 9:247-257.
101. Marsh PD (1994). Microbial ecology of dental plaque and its significance in health
and disease. Adv Dent Res 8:263-271.
102. Colby SM and Russell RRB. (1997) Sugar metabolism by mutans streptococci. J
App Microbiol Symp Supp 83:80S-88S.
103. 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
104. Moller, S., C. Sternberg, J. B. Andersen, B. B. Christensen, J. L. Ramos, M.
Givskov, and S. Molin. (1998) In situ gene expression in mixed-culture biofilms:
42
evidence of metabolic interactions between community members. Appl Environ
Microbiol 64:721–732.
105. Kolenbrander PE, Andersen RN, Blehert DS, Egland PG, Foster JS, Palmer Jr RJ.
(2002) Communication among oral bacteria. Microbiol Molec Bio Rev 66:3:486-
505.
106. Costerton JW (1999). Introduction to biofilm. Int J Antimicrob Agents 11:217-221;
discussion 237-239.
107. Okabe, S, Satoh H, 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.
108. Davey ME, O‟Toole GA (2000). Microbial biofilms: from ecology to molecular
genetics. Microbiol Mol Biol Rev 64:847-867.
109. Stoodley P, Sauer K, Davies DG, Costerton JW (2002). Biofilms as complex
differentiated communities. Annu Rev Microbiol 56:187-209
110. Shu M, Browngardt CM, Chen YY, Burne RA. (2003) Role of urease enzymes in
stability of a 10-species oral biofilm consortium cultivated in a constant-depth film
fermenter. Infect Immun 12: 7188–7192.
111. Filoche SK, Soma KJ, Sissons CH. (2007) Caries-related plaque microcosm
biofilms developed in microplates. Oral Microbiol Immunol 22: 73–79.
112. Moore WEC, Moore LVH. (2000) The bacteria of periodontal diseases. Periodontol
1994(5): 60–77.
43
113. Paster BJ, Boches SK, Galvin JL, Ericson RE, Lau CN, Levanos VA, Sahasrabudhe
A, Dewhirst FE. (2001) Bacterial diversity in human subgingival plaque. J
Bacteriol 3770-3783
114. Donlan RM, Costerton JW. (2002) Biofilms: survival mechanisms of clinically
relevant microorganisms. Clin Microbiol Rev 15:167-193.
115. Watnick P, Kolter R. (2000) Minireview: Biofilm, City of Microbes. J Bacteriol
182(10): 2675-2679
116. Scheie AA, Petersen FC. (2004) The biofilm concept: consequences for future
prophylaxis of oral diseases? Crit Rev Oral Biol Med 15(1)4-12.
117. Ten Cate JM (2006). Biofilms, a new approach to the microbiology of dental
plaque. Odontology 94(1):1-9
118. Sutherland IW (2001). Biofilm exopolysaccharides: a strong and sticky framework.
Microbiology 147:3-9.
119. Whitchurch CB, Erova TE, Emery JA, Sargent JL, Harris JM, Semmler AB. (2002)
Phosphorylation of the Pseudomonas aeruginosa response regulator AlgR is
essential for type IV fimbria-mediated twitching motility. J Bacteriol 184:4544-
4554.
120. Pratten J, Andrews CS, Duncan QMC, Wilson M. (2000) Structural studies of
microcosm dental plaques grown under different nutritional conditions. FEMS
Micriobiol Lett 189(2000): 215-218.
121. Wood SR, Kirkham J, Marsh PD, Shore RC, Nattress B, Robinson C. (2000)
Architecture of intact natural human plaque biofilms studied by confocal laser
scanning microscopy. J Dent Res 79: 21-27
44
122. Zaura-Arite E, van Marle J, ten Cate JM. (2001) Confocal microscopy study of
undisturbed and chlorhexidine-treated dental biofilm. J Dent Res 80:1436-1440.
123. Wood SR, Kirkham J, Shore RC, Brookes SJ, Robinson C. (2002) Changes in the
structure and density of oral plaque biofilms with increasing plaque age. FEMS
Microbiol Ecol 39:239-244.
124. Davies, D. G., A. M. Chakrabarty, and G. G. Geesey. (1993) Exopolysaccharide
production in biofilms: substratum activation of alginate gene expression by
Pseudomonas aeruginosa. Appl. Environ. Microbiol. 59:1181–1186.
125. Costerton JW, Lewandowski Z, Caldwell DE, Korber DR, Lappin-Scott HM.
(1995) Microbial biofilms. Annu Rev Microbiol 49:711-745.
126. Garrett, E. S., D. Perlegas, and D. J. Wozniak. (1999) Negative control of flagellum
synthesis in Pseudomonas aeruginosa is modulated by the alternative sigma factor
AlgT (AlgU). J Bacteriol. 181:7401–7404.
127. Prigent-Combaret, C., O. Vidal, C. Dorel, and P. Lejeune. (1999) Abiotic surface
sensing and biofilm-dependent regulation of gene expression in Escherichia coli. J
Bacteriol. 181:5993–6002.
128. Svensator G, Welin J, Wilkins JC, Beighton D, Hamilton IR. (2001) Protein
expression by planktonic and biofilm cells of Streptococcus mutans. FEMS
Microbiol Lett 205:139-146.
129. Singh J, Khalichi P, Cvitkovitch DG, Santerre JP. (2008) Composite resin
degradation products from BisGMA monomer modulate the expression of genes
associated with biofilm formation and other virulence factors in Streptococcus
mutans. J Biomed Mater Res. In print.
45
130. McNeill K and Hamilton IR (2003) Acid tolerance response of biofilms cells of
Streptococcus mutans. FEMS Microbiol Lett 221:25-30
131. Costerton JW, CHeng KJ, Geesey GG, Lad TI, Nickel NC. (1987) Bacterial
biofilms in nature and disease. Annu Rev Microbiol 41:435-464
132. Whittaker, C. J., C. M. Klier, and P. E. Kolenbrander. (1996) Mechanisms of
adhesion by oral bacteria. Annu. Rev. Microbiol. 50:513–552.
133. Hope CK, Wilson M. (2004) Analysis of the effects of chlorhexidine on oral
biofilm vitality and structure based on viability profiling and an indicator of
membrane integrity. Antimicrob Agents Chemother 48:1461-1468.
134. Welin J, Wilkins JC, Beighton D, Wrzesinski K, Fey SJ, Mose-Larsen P, Hamilton
IR, Scensater G. (2003) FEMS Microbiol Lett 227:287-293
135. Svensater G, Welin J, Wilkins JC, Beighton D, Hamilton IR. (2001) Protein
expression by planktonic and biofilms cells of Streptococcus mutans. FEMS
Microbiol Lett 205:139-146
136. Lendenmann U, Grogan J, Oppenheim FG. (2000) Saliva and dental pellicle- a
review. Adv Dent Rev 14:22-28.
137. Ben-Amar A, Cardash HS. (1991) The fluid-filled gap under amalgam and resin
composite restorations. Am J Dent 4:226-230.
138. 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.
139. Steinberg D, Eyal S. (2002) Early formation of Streptococcus sobrinus biofilm on
various dental restorative materials J Dent 30:47-51.
46
140. Rykke M, Sonju T. (1991) Amino acid composition of acquired enamel pellicle
collected in vivo after 2 hours and after 24 hours.
141. Schilling KM, Bowen WH (1992) Glucans synthesized in situ in experimental
salivary pellicle function as specific binding sites for Streptococcus mutans. Infect
Immun 60:284-295.
142. Nyvad B, Kilian M. (1987) Microbiology of the early colonization of human
enamel and root surfaces in vivo. Scand J Dent Res 95:369-380.
143. Hasty DL, Ofek I, Courtney HS, Doyle RJ. (1992). Multiple adhesins of
streptococci. Infect Immun 60: 2147-2152.
144. Jenkinson HF, Lamont RJ (1997). Streptococcal adhesion and colonization. Crit
Rev Oral Biol Med 8:175-200.
145. Kolenbrander PE. (2000) Oral microbial communities: biofilms, interactions, ad
genetic systems. Annu Rev Microbiol 54:413-437.
146. Yamada T, Igarashi K, Mitsutomi M. (1980) Evaluation of cariogenicity of
glycosylsucrose by a new method of measuring pH under human dental plaque in
situ. J Dent Res 59:2157-2162
147. Igarashi K, Kamiyama K, Yamada T. (1981) Measurement of pH in human dental
plaque in vivo with an ion-sensitive transistor electrode. Arch Oral Biol 26:203-207
148. Stephan RM (1994) Intra-oral hydrogen-ion concentration associated with dental
caries activity. J Dent Res 23:257-266
149. van de Guchte M, Serror P, Chervaux C, Smokvina T, Ehrlich SD, Maguin E.
(2002) Stress responses in lactic acid bacteria. Antonie van Leeuwenhoek 82:187-216
47
150. Banas JA. (2004) Virulence properties of Streptococcus mutans. Front Biosci
9:1267-1277
151. Loesche WJ. (1986) Role of Streptococcus mutans in human dental decay.
Microbiol Rev 50:353-380
152. Bowden GHW. (1991) Which bacteria are cariogenic in humans? In: Risk Markers
for Oral Diseases. 1. Dental caries (Johnson, NW, Ed.) pp 2660286. Cambridge
University Press, Cambridge.
153. Hamada S, and HD Slade (1980) Biology, immunology and cariogenicity of
Streptococcus mutans. Microbiol Rev 44:331-384
154. Svanberg M, Mjor I, Orstavik D (1990) Mutans streptococci in plaque from
margins of amalgam, composite, and glass-ionomer restorations. J Dent Res 69: 861-
864.
155. Bender GR, Sutton SV, Marquis RE. (1986) Acid tolerance, proton permeabilities,
and membrane ATPases of oral streptococci. Infect Immun 53:331-338
156. deSoet JJ, Toors FA, deGraaff J (1989) Acidogenesis by oral streptococci at
different pH calues. Caries Res. 23:14-17.
157. Hamilton IR, and Buckley ND (1991) Adaptation by Streptoccus mutans to acid
tolerance. Oral Microbiol Immunol 6:65-71
158. Dashper SG and Reynolds EC (1992) pH regulation by Streptococcus mutans. J
Dent Res 71:1159-1165
159. Belli WA and Marquis RE (1991) Adaptation of Streptococcus mutans and
Enterococcus hirae to acid stress in continuous culture. Appl Environ Microbiol
57:1134-1138
48
160. Takahashi N, Yamada T (1999) Acid- induced acid tolerance and acidogenicity of
non-mutans streptocci. Oral Microbiol. Immunol. 12:43-48
161. Pedrini D, Gaetti-Jardim Junior EDE, Vasconcelos AC. (2001) Retention of oral
microorganisms on conventional and resin-modified glass ionomer cements. Pesqui
Odontol Bras 15:196-200.)
162. Marsh PD, Martin M (1992) Oral microbiology. 3rd
Ed. London: Chapman & Hall,
pp. 72-74.
163. Bowden GHW, Li YH. (1997) Nutritional influences on biofilm development. Adv
Dent Res 11:81-99
164. Pratten J, Wilson M. (1999) Antimicrobial susceptibility and composition of
microcosm dental plaques supplemented with sucrose. Antimicrob Agents Chemother
43:1595-1599
165. Guggenheim B, Giertsen E, Schupbach P, Shapiro. (2001) Validation of an in vitro
biofilm model of supragingival plaque. J of Dent Res 80(1):363-370.
166. Yabune T, Imazato S, Ebisu S. (2005) Inhibitory effect of PVDF tubes on biofilm
formation in dental unit waterlines. Dent Mater 21:780-786.
167. Rolland SL, McCabe JF, Robinson C, Walls AWG. (2006) In vitro biofilm
formation on the surface of resin-based dentine adhesives. Eur J Oral Sci 114:243-
249.
168. Takahashi Y, Imazato S, Russel RR, Noiri Y, Ebisu. (2004) Influence of resin
monomers on growth of oral streptococci. J Dent Res 83:302-306
169. Kawai K, Tsuchitani Y. (2000) Effects of resin composite components on
glucosyltransferase of cariogenic bacterium. J Biomed Mater Res 51: 123-127.
49
170. Hansel C, Leyhausen G, Mai U, Geurtsen W. (1998) Effects of various resin
composite (co)monomers and extracts on two caries-associated micro-organisms in
vitro. J Dent Res 77:60-67.
171. Khalichi P, Cvitkovitch DG, Santerre JP. (2004) Effect of composite resin
biodegradation products on oral streptococcal growth. Biomat 25:5467-5472.
172. Singh J, Khalichi P, Santerre JP, Cvitkovitch DG. (2005) Composite resin
biodegradation by-products modulate gene expression of Streptococcus mutans.
Presented at the International Association of Dental Research 83rd
Annual Meeting,
Baltimore MD, 2005. Abstract 1016.
173. Khalichi P, Singh J, Santerre JP, Cvitkovitch DG. (2006) Biodegradation by-
products from dental composite resins modulate the gene expression of oral pathogen
Streptococcus mutans. Presented at the Society for Biomaterials 2006 Annual
Meeting, Pittsburgh, PA. Abstract 198.
174. Khalichi P. (2006) Influence of composite resin biodegradation by-products on the
physiology and gene expression of oral bacteria. PhD Thesis, University of Toronto,
Toronto, ON.
175. Robinson C, Kirkham J, Percival R, Shore RC, Bonass WA, Brookes SJ, Kusa L,
Nakagaki H, Kato K, Battress B. (1997) A method for the quantitative site-specific
study of the biochemistry within dental plaque biofilms formed in vivo. Caries Res
31:194-200
176. 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.
50
177. Kramer N, Kunzelmann KH, Garcia-Godoy F, Haberlein I, Meier B, Frankenberger
R. (2007) Am J Dent 20(1):59-64
178. 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
179. 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.
180. Mcbain AJ, Sissons C, Ledder RG, Sreenivasan PK, De Vizio WD, Gilbert P.
(2005) Development and characterization of a simple perfused oral microcosm. J of
Applied MIcrobiol 98: 624-634.
181. 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.
182. Rathsam C, Eaton RE, Simpson CL, Browne GV, Berg T, Harty DWS, Jacques
NA. (2005) Up-regulation of competence- but not stress-responsive proteins
accompanies an altered metabolic phenotype in Streptococcus mutans biofilms.
Microbiol 151:1823-1937.
183. 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.
184. Shokati, B. (2007) Ph.D. Thesis, University of Toronto, Toronto, ON.
51
185. 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
186. White JG, Amos WB, Fordham M. (1987) An evaluation of confocal versus
conventional imaging of biological structures by fluorescence light microscopy. The
Journal of Cell Biology 105:41-48.
187. 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.
188. Skinner A, Hanson K, Caldwell D, Kurtz J. (1996) Comparison of microscope
techniques for the examination of biofilms. J Microbiol Methods 25:57-70
189. 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.
190. Hope CK, Wilson M. (2003) Measuring the thickness of an outer layer of viable
bacteria in an oral biofilm by viability mapping. J Microbiol Methods 54:403-410.
191. Vitkov L, Hannig M, Krautgartner WD, Herrmann M, Fuchs K, Klappacher M,
Hermann A. (2005) Ex vivo gingival-biofilm consortia. Lett Applied Microbiol
41:404-411.
192. Schupbach P, Krejci I, Lutz F. (1997) Dentin bonding: effect of tubule orientation
on hybrid-layer formation. Eur J Oral Sci 105(4): 344-52.
193. Banerjee A, Gilmour A, Kidd E, Watson T. (2004) Relationship between S. mutans
and the auto-fluorescence of carious dentin. Am J Dent 17(4):233-236.
52
194. Sauro S, Pashley DH, Mannocci F, Tay FR, Pilecki P, Sheriff M, Watson TF.
(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.
195. Zavgorodniy AV, Rohanizadeh R, Swain MV. (2008) Ultrastructure of dentine
carious lesions. Arch Oral Biol 53:124-132.
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: 4.0 m Depth: 6.0 m
Depth: 8.0 m Depth: 10.0 m B
Depth: 0.0 m Depth: 2.0 m
Depth: 4.0 m Depth: 6.0 m
Depth: 8.0 m Depth: 10.0 m
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
Hybrid Layer Hybrid Layer
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
biodegradation of dental composites. J Dent Res 83:1:22-26.
3) 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.
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.
71
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
microcosm dental plaques grown under different nutritional conditions. FEMS
Micriobiol Lett 189: 215-218.
12) 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.
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.
72
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
carious lesions. Arch Oral Biol 53:124-132.
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
model to study the relationship of gap size and secondary caries. Caries Res
41:467-473.
26) van Noort R (1994). Introduction to Dental Materials. Papel, Spain: Times Mirror
International Publishers Limited; pp. 3-145.
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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.
4.2 DISCUSSION RE: HYPOTHESIS #2
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.
4.3 DISCUSSION RE: HYPOTHESIS #3
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.
2. 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.
3. Finer Y, Santerre JP. (2004) The influence of resin chemistry on a dental
composite‟s biodegradation. J Biomed Mater Res A. 69(2)69A:233-246.
79
4. Jaffer F, Finer Y, Santerre JP. (2002) Interactions between resin monomers and
commercial composite resins with human saliva derived esterases. Biomat
23:1707-1719.
5. Finer Y, Santerre JP. (2004) The influence of resin chemistry on a dental
composite's biodegradation. J Biomed Mater Res A. 69(2):233-46.
6. 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.
7. Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
8. Bouillaguet S. (2004) Biological risks of resin-based materials to the dentin-
pulp complex. Crit Rev Oral Biol Med 15(1):47-60.
9. van Noort R. Introduction to Dental Materials. Papel, Spain: Times Mirror
International Publishers Limited; 1994. pgs. 3-145.
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
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
15. Pratten J, Andrews CS, Duncan QMC, Wilson M. (2000) Structural studies of
microcosm dental plaques grown under different nutritional conditions. FEMS
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.
19. 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.
81
20. Zavgorodniy AV, Rohanizadeh R, Swain MV. (2008) Ultrastructure of dentine
carious lesions. Arch Oral Biol 53:124-132.
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.
22. 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.
23. 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.
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
model to study the relationship of gap size and secondary caries. Caries Res
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.
27. 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.
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.
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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
1. 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.
2. 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.
3. Arola D, Reprogel RK. (2005) Effects of aging on the mechanical behavior of
human dentin. Biomaterials 26:4051-4061.
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.
14. 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.
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.
4. 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.
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.
8. Jaffer F, Finer Y, Santerre JP. (2002) Interactions between resin monomers and
commercial composite resins with human saliva derived esterases. Biomat
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
to their degradation and the release of hydrolyzed polymeric-resin-derived
products. Crit Rev Oral Biol Med 12(2):136-151.
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
biodegradation of dental composites. J Dent Res 83:1:22-26.
5) 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.
6) 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
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
EL. (2001) The stability of methacrylate biomaterials when enzyme challenged:
Kinetic and systematic evaluations. J Biomed Mater Res 57:522-531.
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
their polymers. Scand J Dent Res 98:351-355.
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
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
15) 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.
16) Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
17) 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
18) Finer Y, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and
psuedocholinesterase on the biodegradation of dental composites. Biomat 25:1787-
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
commercial composite resins with human saliva derived esterases. Biomat 23:1707-
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
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) 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
activities within human saliva which are relevant to dental resin composite
biodegradation. Biomat 26:4259-4264.
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
biodegradation products in micro-filled bis-GMA/TEGDMA dental composite
resins. Biomat 20:1897-1908.
6) Finer Y, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and
psuedocholinesterase on the biodegradation of dental composites. Biomat 25:1787-
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
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.
113
2) Finer Y, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and
psuedocholinesterase on the biodegradation of dental composites. Biomat 25:1787-
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
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, Jaffer F, Santerre JP. (2004) Mutual influence of cholesterol esterase and
psuedocholinesterase on the biodegradation of dental composites. Biomat
25:1787-1793.
3. 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.
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.
6. Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
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.
9. Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
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
1. Santerre JP, Shajii L, Tsang H. (1999) Biodegradation of commercial dental
composites by cholesterol esterase. J Dent Res 78(8):1459-1468.
2. 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.
3. Jaffer F, Finer Y, Santerre JP. (2002) Interactions between resin monomers and
commercial composite resins with human saliva derived esterases. Biomat
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
composite‟s biodegradation. J Biomed Mater Res 69A:233-246.
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.
Arch Oral Biol 46:471-476.
2. 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
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.
6. Pratten J, Andrews CS, Duncan QMC, Wilson M. (2000) Structural studies of
microcosm dental plaques grown under different nutritional conditions. FEMS
Micriobiol Lett 189(2000): 215-218.
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.
8. Pratten J, Andrews CS, Duncan QMC, Wilson M. (2000) Structural studies of
microcosm dental plaques grown under different nutritional conditions. FEMS
Micriobiol Lett 189(2000): 215-218.
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
1. 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.
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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