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Parvimonas micra adhesion on different implant surfaces: an in vitro pilot study

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Parvimonas micra adhesion on different implant surfaces: an in vitro pilot study

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Content Parvimonas micra
ADHESION ON DIFFERENT IMPLANT SURFACES:
AN in vitro PILOT STUDY

by

Ching Hsiu Ketty Chiu
_________________________________________________________

A Thesis Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
In Partial Fulfillment of the
Requirements for the Degree
MASTER OF SCIENCE
(CRANIOFACIAL BIOLOGY)

August 2010

Copyright 2010 Ching Hsiu Ketty Chiu
ii

Dedication
I would like to dedicate this manuscript to my father Dr. 邱文達, my mother 康文娟,
my brother Jason Chiu and Dr. Allan Leung. Special dedication to all my extended
family and friends who supported and trusted in me. Last but not least to Doby Chiu.

iii

Acknowledgements
I would like to thank Dr. Hessam Nowzari for providing me this wonderful
opportunity to pursue my master’s degree. His passion for research has inspired and led
me through this process of learning. Furthermore, I would like to thank Dr. Sandra Rich
who worked tirelessly and patiently with me on my project. Her passion for teaching is
truly inspiring.
I acknowledge my colleague Dr. Glenn Chang, who aided in designing this
project. I am grateful to Dr. Jorgen Slots for allowing me to be a part of his laboratory. I
also would like to thank Drs. Mahvash Navazesh and Kian Kar for being a part of this
process of pursuing my master’s degree and giving me valuable advice on my residency
and research project.
I also would like to thank Pauline Chang, Alicia Thompson and Dr. Cheryl Vigen,
for their support in providing their expertise in fields of microbiology, SEM processing
and operation, and statistical analysis.
I gratefully acknowledge my father and my mother for their love, inspiration and
support, my brother for his encouragement, and Doby for his loyalty and companionship.
Lastly, I would like to reserve a special word of gratitude to Allan Leung for his
patience, help, and encouragement.
iv

Table of Contents
Dedication ii
Acknowledgements iii
List of Figures v
List of Tables viii
Abstract ix
Chapter 1: Introduction and Background 1
Chapter 2: Materials and Methods 26
Chapter 3: Results 40
Chapter 4: Discussion 81
Chapter 5: Conclusion 111
References 112

v

List of Figures
Flowchart 1. Experiment design 28
Figure 1. Plastic vial containing different implants 34
Figure 2. US-12 utility 12 c.c. curved tip syringe 35
Figure 3. Example of x5000 magnification of image 37
with counting grids superimposed

Figure 4. 3i machined implant (x5000) 40
Figure 5. Branemark machine implant (x5000) 41
Figure 6. Osseotite implant (x5000) 41
Figure 7. Osseotite implant (x20,000) 42
Figure 8. Nanotite implant (x5000) 42
Figure 9. Nanotite implant (x20,000) 43
Figure 10. Implantium SLA+ implant(x5000) 43
Figure 11. Implantium SLA+ implant (x20,000) 44
Figure 12. Implantium anodized implant (x5000) 44
Figure 13. Implantium anodized implant (x20,000) 45
Figure 14. Branemark TiUnite implant (head x2000) 45
Figure 15. Branemark TiUnite implant (head x5000) 46
vi

Figure 16. Branemark TiUnite implant (middle x2000) 46
Figure 17. Branemark TiUnite implant (middle x5000) 47
Figure 18. Effect of serum in group A, B, and C 58
Figure 19. Effect of saline rinsing in group A 59
Figure 20. Effect of saline rinsing in group B 60
Figure 21. Group B2 3i machined implant 61
(head portion) without saline rinsing (x5000)

Figure 22. Group B3 3i machined surface implant 61
(head portion) with saline rinsing (x5000)

Figure 23. Group B2 Branemark machined implant 62
(tip portion) without saline rinsing (x5000)

Figure 24. Group B3 Branemark machined implant 62
(tip portion) with saline rinsing (x5000)

Figure 25. Group B2 Nanotite implant 63
(head portion) without saline rinsing. (x5000)

Figure 26. Group B2 Nanotite implant 63
(head potion) without saline rinsing (x20,000)

Figure 27. Group B3 Nanotite implant 64
(head portion) with saline rinsing (x5000)

Figure 28. Group B3 Nanotite implant 64
(head portion) with saline rinsing (x20,000)

Figure 29. Effect of saline rinsing in group C 65
Figure 30. Group C3 Nanotite implant 66
(middle section) after rinsing (x 5000)
Figure 31. Group A1 head portion of 3i machined implant (x5000) 67
Figure 32. Group A1 head portion of 3i machined implant (x20,000) 67
vii

Figure 33. Group A1 middle portion of Nanotite implant (x5000) 68
Figure 34. Group B2 head portion of Nanotite implant (x5000) 68
Figure 35. Group B2 head portion of Nanotite implant (x20,000) 69
Figure 36. Group B2 head portion of Osseotite implant (x5000). 70
Figure 37. Group B2 head portion of Osseotite implant (x20,000) 70
Figure 38. Group C1 head portion of Osseotite implant (x5000) 71
Figure 39. Group C1 head portion of Osseotite implant (x20,000) 72
Figure 40. Group A1 head portion of Implantium anodized implant (x5000) 73
Figure 41. Group A1 tip portion of Implantium anodized implant (x20,000) 74
Figure 42. Group A1 head portion of Implantium anodized implant (x5000) 75
Figure 43. Group C1 head portion of Implantium SLA+ implant (x5000) 75
Figure 44. Group C1 head portion of Implantium SLA+ implant (x20,000) 76
Figure 45. Group A1 head portion of Branemark machined implant (x5000) 77
Figure 46. Group A1 head portion of Branemark machined implant (x20,000) 77
Figure 47. Group B2 tip portion of Branemark machined implant (x5000) 78
Figure 48. Group A1 middle portion of TiUnite implant (x5000) 79
Figure 49. Group B1 tip portion of TiUnite implant (x5000) 80
viii

List of Tables
Table 1. Listing of implants, manufacturer and surface treatment description 26
Table 2. Specific implant information 29
Table 3. Implant groups and implants included 38
Table 4. Percentage of P. micra adhesion on different implants 48
among different treatment groups

Table 5. Percentage of P. micra adhesion in all groups 52
Table 6. Average adhesion percentage of P. micra in different groups 53
Table 7. Percentage of P. micra adhesion in all implants 55
Table 8. Percentage of P. micra adhesion in group A, B, and C 57
ix

Abstract
Objectives: P. micra is considered to be a putative peri-implantitis bacterium which is
often isolated from active peri-implantitis. The aim of this in vitro study is to evaluate the
ability of P. micra to adhere to different implant surface textures that are currently on the
market. In addition, same surfaced implants between different manufacturing companies
were also compared.
Materials and Methods: Seven types of implants were selected: 3i Nanotite implants, 3i
Osseotite implants, Implantium anodized implants, Implantium SLA+ implants (calcium
phosphate coated), 3i machined surface implants, Branemark TiUnite implants, and
Branemark machined implants. All implants were divided into three groups, A, B, and
C; each consisted of 21 implants (3 sets of seven different implants). Implants were
submerged in bacterial broth for 4 hours (group A), 24 hours (group B) in atmospheric
conditions and 4 days under anaerobic conditions in nutrient broth (group C). Groups A,
B, and C were further divided into 3 subgroups, 1, 2, and 3. Subgroups A1, B1, and C1
were incubated with bacteria only. For subgroup A2, B2, and C2, implants were pre-
treated with bovine serum to simulate gingival crevicular fluid. As for subgroup A3, B3,
and C3, implants were pre-treated with bovine serum and then irrigated with saline after
incubation. All implants were immediately fixed, processed and evaluated under
scanning electron microscopy. Images of SEM were photographed and used for
calculation of the percentage of P. micra adhesion. Adhesion percentages among
different surface characteristics and treatments were compared.
x

Results: P. micra adhesion was observed on all implant surfaces. Group B presented
with the highest percentage of adhesion. When implants were grouped according to their
surface characteristics, the average adhesion percentage of rough surface was 16.2% and
machined surface was 12.2%. The acid-etched group had a higher average bacterial
adhesion (23.2%) when compared to machined (12.6%) and anodized surfaces (5.7%).
The average bacterial adhesion percentage for implants with calcium phosphate was
23.5% which was higher than the non-calcium phosphate treated implants (11.9%).
Similar implant surfaces were compared among different companies and 3i machined
implants presented with an average adhesion rate of 8.8% while Branemark machined
implants averaged 16.5%. Implantium anodized implants presented with 6.7% which
was fairly similar to Branemark TiUnite with 4.7%. Effect of serum was inconsistent in
the experiment. Saline rinsing appeared to be effective in decreasing percentage of P.
micra, especially for the machined surface. Zero (0%) adhesion rate was observed on
machined surfaces (by both manufacturers) in group A and B after saline rinsing.
Conclusion: P. micra adhered to all implant surfaces in this investigation. Surface
treatments such as acid etching and deposition of calcium phosphate appeared to increase
percentages of P. micra adhesion. Even though the adhesion of P. micra was also
observed on the machined surfaces, it may be cautiously concluded by this study of
limited samples that smooth surfaces may be more easily cleansable by rinsing. In
addition, this study showed that same surface modification produced by different
manufacturers may present with different characteristics and possibly alter their
xi

interaction with bacteria. However, due to the in vitro nature of this study and the limited
number of samples examined, future experiments will be needed to confirm these results.

1

Chapter 1: Introduction and Background
The invention of dental implants several decades ago revolutionized dentistry. In
1950’s Dr. Per-Ingvar Brnemark discovered the stable fixation of titanium in the bone
tissue of rabbits and the term “osseointegration” was coined and described as a “direct
structural and functional contact between ordered, living bone and the surface of a load-
carrying implant” (Brånemark et al., 1977). Ever since then dental implants have
gradually become the best treatment option available to replace natural teeth in many
circumstances. Despite the high success rate of implant dentistry in recent years, implant
failures due to peri-implant diseases do exist.

Peri-implantitis
Two types of peri-implant diseases are commonly recognized, peri-implant
mucositis and peri-implantitis, both describing an inflammatory response around the peri-
implant tissue. Peri-implant mucositis is a term used to describe the inflammatory
reaction around the peri-implant tissue without any radiographic loss of bone. On the
other hand, peri-implantitis, one of the major causes of implant failure, is an
inflammatory response around osseointegrated implants, resulting in loss of bone around
an implant in function. (Albrektsson & Isidor, 1994 and Zitzman & Berglundh, 2008).
Several authors reported high rates of implant failures due to peri-implantitis.
Espositos et al. (1997) found implant removal rates due to peri-implantitis ranged from 8-
50%. Zitzman and Berglundh discussed prevalence of peri-implant diseases in a review
2

paper in 2008. They included only longitudinal and cross-sectional studies, excluding
any studies with less than fifty implants, less than 5 years of implant functioning time and
lack of information on bleeding upon probing. They found that 28% to more than 56%
of the subjects presented sites with peri-implantitis. In addition, between 12 – 43% of
implants showed bone loss when bleeding on probing was present.
Ample evidence exists demonstrating the role of oral bacteria in inducing peri-
implantitis. Many species of bacteria have been isolated from implants affected with
peri-implantitis. In 1987, Mombelli et al. compared microbiota associated with
successful and failing implants. Hollow cylinder ITI implants were used and they
classified failing implants as having 6mm or greater probing depths, presentation of
suppuration and radiographic bone loss. In the failing implant site, increased proportion
of gram negative anaerobic rods, black pigmented Bacteroides and Fusobacterium spp.,
spirochetes, fusiform bacteria, and motile and curved rods were found. As for control
sites (successful implants) in the same patient, coccoid cells were the predominant
morphotype. Rams et al. in 1991 sampled ailing hydroxyapatite-coated implants and
isolated high proportions of Fusobacterium species and Peptostreptotoccus prevotii.
Of all the common oral periodontal pathogens, Parvimonas micra was frequently
isolated from the peri-implantitis sites in a much higher level.
Alcoforado et al. isolated varieties of microbes from eighteen failing implants in
1991. Parvimonas micra and Wolinella recta were isolated from 6 out of 18 implants.
Enteric rods and pseudomonads were isolated in significant amounts. Actinobacillus
3

actinomycetemcomitans, Capnocytophage species, non-pigmented Bacteroides species
and staphylococci were also detected in several failing implants as were Candida
albicans and Bacteroides intermedius.
Salcetti et al. (1997) used gingival crevicular fluid and plaque samples with DNA
oligonucleotide probes and discovered increased levels of P. micra, P. nigrescens, F.
nucleatum, ss vincentii and F. nucleatum ss nucleatum at failing implant sites with more
than 2mm of bone loss. In addition, levels of microbes were found to be correlated to the
level of prostaglandin E2.
Listgarten and Lai (1999) studied failing implant sites in partially edentulous
patients. In 59% of these sites, B. forsythus was isolated, 54% had spirochetes, 41% had
Fusobacterium, 39% had P. micra and 27% had P. gingivalis. Furthermore, the study
compared the bacterial distribution above with those observed in adult periodontitis and
refractory periodontitis. It concluded that the same bacteria were present but in different
percentage distributions.
Hultin et al. (2002) retrieved samples from patients with peri-implantitis and
found that they harbored higher levels of periodontal pathogens including Actinobacillus
actinomycetemcomitans, Porphyromonas gingivalis, Prevotella intermedia, Bacteroies
forsythus and Treponema denticola. Of note, Parvimonas micra was present in all
samples collected.
4

Tabanella et al. (2009) examined clinically ailing implants, which was defined as
bone loss greater the level of the third implant thread. They found that peri-implant bone
sites appeared to have deeper pocket depths, were symptomatic, and contained T.
forsythia, Campylobacter species and Parvimonas micra. Implant thread exposure
without deeper peri-implant pocket depths were found with increased amounts of P.micra
and Campylobacter species. However, Fusobacterium species, T. forsythia,
Campylobacter species and P. micra were also common bacteria found in peri-implant
sites with bone loss. Interestingly, symptomatic implants had the tendency to isolate P.
micra, Fusobacterium species and Eubacterium species. Also of note, patients who were
premedicated with 2g amoxicillin 1 hour prior to implant surgery appeared to have more
P. micra and Campylobacter species in their ailing implant sites.
All of the previously mentioned studies suggest a role for P. micra as a putative
peri-implantitis bacterium.

Parvimonas micra microbiology
Parvimonas micra was originally named Peptostreptococcus micros. The genus
Peptostreptococcus was described in 1963 and considered the anaerobic equivalent of
Streptococcus. Peptostreptococcus micros was listed in the approved list in 1980.
However, it was re-classified in 1999 by Murdoch and Shah and its designation changed
to Micromonas micros. In 2006, due to the incorrect genus placement, P. micra was once
5

again proposed for a name change to its current name, Parvimonas micra by Tindall and
Euzeby. P. micra can be found outside of the oral cavity and detected in mixed anaerobic
infections from different body sites such as brain abscess, sinus infections, human bite
wounds, pleural empyema, otitis media, septicemia, anorectal abscess, prosthetic joint
infections, vertebral osteomyelitis, gynecological infections and intra-abdominal
infections (Song & Finegold, 2007).
P. micra is a gram-positive bacterium of 0.3 to 0.7μm in diameter, which usually
arranges in chains and pairs (Murdoch & Mitchelmore, 1991) but can also be found in
clumps when cultured on solid media (Moore et al. 1986). P. micra cells are relatively
aerotolerant. Features that enhance the pathogenesis of P. micra in periodontium include
proteolytic enzyme activity (Ezaki & Yabuuchi, 1985 and Murdoch et al., 1988),
inhibition of gingival fibroblast and epithelial growth (Tam et al. 1987) and elaboration
of hyaluronidase (Tam & Chan, 1985). Hyluronidase is known to destroy the ground
substance of connective tissue which leads to destruction of periodontium and facilitation
of spread of bacterial toxin. It is also known that certain levels of hyaluronidase cause
partial proteoglycan breakdown, exposing the cryptic antigenic determinants which in
turn induce autoimmune reactions (Tam & Chan, 1985). Collagenase production and
hemolytic activity had also been observed from P. micra (Ota-Tzusuki et al. 2010).
In addition, Persson et al. 1990 noted the ability of P. micra to produce volatile
sulfur compounds, such as hydrogen sulfide from L-cysteine. Hydrogen sulfide is a
cytotoxic agent and its presence can be detected in high amounts in the periodontal
6

pocket. Carlsson et al. (1993) further investigated this ability and found that L-cysteine is
only present in low concentrations in the periodontal pocket and that P. micra may form
hydrogen sulfide using other more abundant sources of sulfur, namely, glutathione, L-
gamma-glutamyl-L-cysteinylglycine. Furthermore, cell wall of P. micra was found to
have the ability to illicit pro-inflammatory response in macrophages, leading to increased
production of cytokines, chemokines and MMP-9 (Tanabe et al., 2007).
Morphologically, P. micra exists in a smooth (Sm) or rough (Rg) form, depending
upon the presence or absence of fibrillar structures (Van Dalen et al., 1993). Both types
can be isolated from subgingival specimens. Rg presents with fibril like structures while
Sm lacks the fibril structure. The smooth morphotype is hydrophobic, and the rough one
is hydrophilic (Kremer et al., 2000). In addition to the Sm and Rg forms, the Rg type can
be easily converted into a smooth variant type (Rg
sm
) which closely resembles Sm but
serologically and phylogenetcially is the same as Rg type (Kremer et al., 1997). The
Rg
sm
is similar to Sm in that it lacks fibril surface structures. A study by Kremer et al.
(1999) showed the fibril like structures of the rough morphotype do not facilitate the
adhesion of P. micra to epithelial cells. Instead this structure served as an obstruction for
adhesion. It was instead the extracellular polysaccharide present on all three
morphotypes that facilitated the adhesion of P. micra to epithelial cells. Kremer and van
Steenbergen’s study (2000) also reported the non-functional role of the proteinous fibril
like structure in the coaggregation of other bacteria with P. micra. The function of this
structure remains to be discovered.
7

Surface modification of titanium implants
Over the years, implant manufacture companies have developed numerous
designs of implant surface textures which claim to achieve a more efficient and superior
osseointegration. The concept of roughening the titanium surface is to create a more
developed area and increase bone anchorage to improve the interlocking mechanism of
bone and implant (Coelho, 2009a). Cooper (2000) in a systematic review concluded that
altering implant surface to increase surface topography was encouraged because of its
improved the bone-to-implant contact and mechanical properties. However, the author
noted the absence of controlled comparative clinical trials and studies on altered surface
in relation to peri-implantitis.
Le Guéhennec et al. (2007) in a review also agreed that surface roughening
enhanced osseointegration, although the exact mechanism and chemistry behind the early
bone healing events were still unclear. Further research in this area will be required.
Note: Our study exclusively used titanium implants, therefore this discussion will be
limited to implants of that material.
Roughness of implants can be measured quantitatively in two or three dimensions.
Average roughness in two dimensions, Ra, represents the average height deviation from
the profile. On the other hand, Sa is the average roughness over a surface and provides
information on a given surface area in three dimensions. Wennerberg and Albrektsson
(2010) advocated the use of Sa value over Ra in assessing implants because Sa is not
8

influenced by the direction of measurement and was more consistent and reliable.
Nonetheless, Ra values are more commonly found in the literature. Sdr, another
commonly used measurement represents the developed surface area ratio, a parameter
that describes the complexity of a surface. It provides information on height and number
of peaks in a given area. Thus, a surface with no height deviations (flat surface), will
have an Sdr of 0%.
Originally, from the original Branemark implants, machined surface or so called
turned surface implants were considered the “gold standard.” Albrektsson and
Wennerberg suggested in 2004 that machined surface implants have an Sa value of less
than 0.5µm. The same authors reported Branemark’s original machined surface
implants to have an Sa of 0.9µm and an Sdr of 34% while the 3i machined surface
(recorded from the coronal portion of an Osseotite since machined surfaced 3i implants
were no longer on the market) had an Sa of 0.40µ and Sdr of 17%. This inconsistency in
implant measurements complicates comparison between implants of similar surface type.
Alteration of the machined implants produces another implant category, the
roughened surfaced implants. Generally speaking, there are two different methods of
modifying implant surfaces; by means of adding or subtracting materials. Examples of
subtraction processing of implant surface include acid etching, different methods of
polishing, blasting, and oxidation (Wennerberg & Albrektsson, 2009).
Acid etching is a method to remove some amount of material to create a pitted
surface on titanium. Hydrochloric acid, sulfuric acid, and hydrofluoric acids are some of
9

the frequently used acids. Osseotite is a dual etched implant (HCl/H
2
SO
4
) manufactured
by Biomet 3i (Palm Beach Gardens, FL, USA). The Osseotite implants feature a
machined collar and acid etched body. The Sa is 0.68µm and Sdr of 27% at the acid
etched portion while the machined part demonstrates Sa of 0.40µm and Sdr of 17%
(Wennerberg & Albrektsson 2010).
Another surface modifying technique is blasting. Blasting procedure creates a
roughness which is believed to be optimal for mechanical fixation (Wennerberg &
Albrektsson, 2009). Grit blasting can be performed by different sized particles, such as
silica (sand), bioceramic, alumina, or titanium oxide. These blasted implants are then
acid etched to remove processing byproducts as well as to smooth out sharp peaks and
edges and increase total surface area which may potentially improve protein adhesion
(Wennerberg & Albrektsson, 2009 and Coelho et al., 2009a). Commonly used acids are
sulfuric, hydrofluoric, nitric acids or a combination of different acids (Coelho et al.
2009a). The usual Ra value for this type of implant surface is approximately 0.5 - 2µm.
One of the better known blasted and acid-etched surface implants is the sand-
blasted with large grit and acid-etched (SLA) implant manufactured by Straumann (Basel,
Switzerland). It generally has an Sa of 1.5µm and an Sdr of 34% (Wennerberg &
Albrektsson 2010). This surface is not exclusive to Straumann as SLA surfaces are also
manufactured by other companies such as Dentium. Straumann utilizes sulfuric acid and
hydrochloric acid for etching and blast with alumina particles of 250 – 500 µm while
Implantium SLA (Dentium, Seoul, Korea) use warm hydrochloric acid and alumina
10

particles of unknown size (Kim et al., 2008). The resultant Ra value reported by Kim et
al. (2008) for Implantium SLA was determined to be 1.19 µm.
Another commonly utilized modification method is anodizing the titanium surface.
Anodized surfaces not only present with an increase in surface microtexture, but also a
change in surface chemistry (Coelho et al. 2009a). TiUnite surfaced implants are
anodized implants produced by Nobel Biocare (Zurich, Switzerland). An anodic
oxidation process (electrochemical anodization) in a strong acid solution is used to
modify its machined surfaces. Phosphoric acid was suspected to be used for TiUnite
according to Wennerberg and Albretksson’s investigation in 2010. This method creates a
thick layer of titanium oxide, and the characteristic pore formation in the oxide layer
(Coelho 2009b). According to Sul et al. (2006) the surface of TiUnite displays a duplex
oxide structure with an outer and inner film. This film is mainly composed of TiO2,
phosphorus, calcium, sodium, nitrogen and traces of sulfur. The outer porous film has
micropores present and has a thickness of 0.9 and 5µm. The inner barrier film is about
5.7µm at the head of the implant, 5.9µm at the third and 9.3µm at the fifth thread. The
pore/pit size is less or equal to 4µm. Albrektsson and Wennerberg (2004) reported an
observation of unevenly distributed roughness and oxide layer. These TiUnite implants
had an Sa of 1.1 µm and an Sdr of 37% (Wennerberg & Albrektsson, 2010).
Examples of additive processing include hydroxylapatite or calcium phosphate
coatings (CaP), titanium plasma sprayed (TPS) and ion deposit (Wennerberg and
Albrektsson, 2010). The concept behind addition of CaP is to increase the saturation by
11

body fluids which results in deposit of biological apatite layer onto the implant surfaces
(De Groot et al. 1998). On the other hand, calcium phosphate is claimed to be capable
of absorbing proteins to their surfaces, which could potentially lead to increase platelet
adhesion, activation and accelerate healing (Davies 2003). Increase protein adsorption
could also improve fibrin binding to the implant surface and facilitate the establishment
of matrix for osteogenic cell migration to implants (Davies, 2003).
In the current market, creating more surface area by means of subtractive
technique combined with adding calcium phosphate appears to be popular. Nanotite
from Biomet 3i (Palm Beach Gardens, FL, USA) is a good example. The coating
nanocrystals of calcium phosphate has also been advocated in addition to dual acid
etching. Nanotite possesses an Sa of 0.5 µm and an Sdr of 40% (Wemmerberg &
Albrektsson, 2010). According to Mendes et al. (2007) dual etched implants were dipped
into an alcohol based solution containing nano-sized CaP particles about 20-100nm in
size. These titanium implants were then dried in an oven at 100 degree Celsius after the
dipping process. This process created discrete crystalline depositions (DCD) of nano
CaP.
A similar concept was adopted by Implantium implants manufactured by Dentium.
A surface called SLA+ was manufactured by sand-blasted with large-grit, acid etched and
coated with calcium phosphate. For implantium SLA + implants, CaP was suspected to
be deposited by an ion-beam assisted deposition (IBAD) technique (Yoon et al., 2009)
which is different than that used in the manufacture of Nanotite implants.
12

The advantages of improved bone healing (Yoon et al., 2009), higher tensile
forces (Mendes et al., 2007), higher bone to implant contact (Mendes et al., 2009 and
Orsini 2007), and improved implant stability (Goené et al., 2007) were often described
for these newly developed roughened surface implants. Nevertheless the negative effects
of these surfaces were rarely discussed.

Bacteria adhesion on titanium surfaces
The composition of the bacterial environment as well as the ability of bacteria to
adhere to the implant surface appears to be two risk factors of peri-implantitis (Quirynen
et al., 2002). The ability of bacteria to adhere titanium implant surfaces has been
confirmed in various studies. Bacteria ranging from Streptococcus sanguis (Pereira da
Silva et al., 2005, Drake et al., 1999), Actinomyces viscosus (Wu-Yuan et al., 1995),
Porphyromonas gingivalis (Amoroso et al., 2006) and Actinobacillus
actinomycetemcomitans (Yoshinari et al., 2000) have been reported to have the abilities
to adhere to titanium surfaces. Cimasoni and McBride (1987) also documented the
successful adherence of Treponema denticola on modified hydroxyapatite. Nevertheless,
studies on P. micra and its ability to adhere to titanium cannot be located.
In addition, all titanium surfaces used in experiments mentioned above are in vitro
studies involving titanium surfaces modified by the authors instead of the manufacturer’s
factory made implants. Each manufacturer has its own processing method, whose details
13

may not be disclosed to the public. And although the principles of manufacturing
implants may be the same, minor details may differ from company to company.
Therefore these studies may not reflect the true characteristics of implants.

Physicochemical mechanisms to bacterial adhesion
Teughels and colleagues (2006) summarized the following phases involved in
bacterial adhesion process.
Phase I: bacterial transportation to the surface
In order for bacterial cells to adhere to a given surface (solid surface or another
cell), multiple forces and conditions need to come into play (Quirynen & Bollen, 1995).
First of all, the bacterial cells need to be transported to the vicinity of the surface through
random movement of particles in the liquid environment (Brownian movement),
convective transport along with liquid flow, or active movement of bacteria.

Phase II: The initial adhesion of bacteria
This initial adhesion stage involves weak and reversible adhesion through short
and long range forces. Two physicochemical aspects are commonly used to describe
these interactions.
14

1. Surface free energy of the interacting surfaces
Prior to bacterial adhesion, the water film between the surface and bacteria has to
be removed. Thus the energy of interaction may be calculated from an assumption that
this interface between bacterium to liquid (bl) and solid to liquid (sl) has been replaced
with a bacteria to solid surface (sb) interface. Absolom et al. (1983) proposed the
formula describing this change in energy upon adhesion:
∆Gadh = γsb – γsl – γbl
This formula explained the bacterial interfacial free energy of adhesion (∆Gadh)
and its correlation with solid-bacterium interfacial free energy (γsb), the solid-liquid
interfacial free energy (γsl) and the bacterium-liquid interfacial free energy (γbl). When
the ∆Gadh is negative, adhesion of the bacterium is favored.
The following assumption can be made from the formula mentioned above. If the
SFE of liquid is greater than the SFE of bacteria, then the SFE of the substratum will
increase and become hydrophilic. As a result, ∆Gadh will drop and discourage bacterial
adhesion to the substratum. On the other hand, if the SFE of liquid is lesser than SFE of
bacteria, SFE of substratum is increased becoming more hydrophobic and promoting
bacterial adhesion.
In the oral cavity, oral bacteria generally present with high SFE (Weerkamp et al.
1985) and saliva with a relatively low SFE (Glantza 1970), increasing the SFE of the
15

substratum and creating forces favoring bacteria adhesion to any substratum present (ie.
enamel, implant surfaces, or restorative materials) present.
In addition, the above mentioned formula of free energy also suggest that bacteria
with low SFE will preferentially adhere to surfaces with low SFE and bacteria with high
SFE will prefer the high SFE surfaces (Quirynen and Bollen, 1995).

2. The interaction energies between surface and bacterium
The initial adhesion of bacteria involves multiple forces which depend upon the
distance of the bacteria to the surface. Different forces that affect adhesion come into
play depending upon the distance the bacterium is from a surface. When the distance is
more than 50 nm, Van der Waal’s forces, which are the attractive or repulsive force
between molecules other than covalent bonds or electrostatic interaction, is the only force.
When the distance between the bacterium and the surface is about 10-20 nm, both Van
der Waal’s force and electrostatic forces will exert their effects. At this time, repulsion is
maximized due to both the bacterium and surface being negatively charged. Bacteria that
possess fimbrae or other polymers can be very effective in overcoming this repulsion.
This is due to the small diameter of the fimbrae, which result in smaller areas of negative
charge on the bacterial surface, reducing its repulsion from the surface to be adhered to.
Lastly, between 1-5nm, short range forces such as hydrogen bonding, ion pair formation,
steric interaction, covalent bond and bridging interaction from the cellular filamentous
16

appendages take effect. Eventually when the forces are strong enough, adhesion will turn
into an irreversible adhesion (Ofek et al., 2003).

Phase IV: Colonization/plaque maturation
The bacteria can start growing and form biofilm after they have attached firmly
onto the surface.

Negative aspect of roughened surface
Roughened surfaces produce a sheltering effect for bacterial adhesion protecting
them from shearing forces, and allowing them the stability and time to easily transform
their adhesion from a reversible bond to an irreversible bond onto the substrate (Quirynen
& Bollen, 1995). In addition, rough surfaces are capable of accumulating increased
thickness/area of plaque and number of colony forming units of bacteria. As time
progresses with undisturbed matured plaque, rough surfaces tend to harbor more motile
organisms and spirochetes (Quirynen & Bollen, 1995). Many studies have shown this
effect of rough surfaces on bacterial adhesion in multiple surfaces including restorative
material, teeth and titanium.
Quirynen et al. (1990) demonstrated the effect of surface roughness by applying
two strips of material glued on human teeth, one rough (Ra = 2µm) and one smooth (Ra =
17

0.1µm). Only a quarter of the smooth region displayed plaque accumulation while the
rough region was completely covered by biofilm after three days cessation of oral
hygiene.
Rimondini et al. (1997) demonstrated higher bacterial count in rough surfaces in
an in vivo study involving titanium discs with various roughnesses after suspension of
oral hygiene of 24 hours. In addition, more long and short rods were noted in the
rougher surfaces when compared to the smooth ones.
Tanner and colleagues (2005) tested four different materials with Ra values
ranging from 0.05 to 0.51 µm bonded to the buccal surface of a molar. After 24 hours
intra-orally they found the roughest surface with highest colony forming units of total
facultative bacteria and plaque formation.
Subgingivally, Waerhaug (1956) demonstrated roughening of subgingival enamel
in both dogs and monkey induced more deposit of plaque and calculus, and also resulted
in more inflammation.
From a microbiological standpoint, rough surfaces appear to harbor more oral
bacteria. Rams et al. in 1991 reported a higher percentage of P. micros (P. micra) in
healthy hydroxyapatite-coated fixtures (17.4%) than in healthy pure titanium fixtures
(9.8%). Quirynen et al. (1993) also reported rough surface abutments harbor twenty five
times more bacteria subgingivally when compared to smoothed abutments.
18

The majority of studies indicate that a rough surface in general creates a friendlier
environment for microbial adhesion. Nevertheless, data on newly marketed implant
surfaces are surprisingly lacking with regard to the infectious aspect of peri-implantitis.
Currently, no studies exist on Parvimonas micra and its ability to adhere to dental
implants even thought P. micra appears to be a putative pathogenic bacterium and is
likely a factor in peri-implantitis.

Peri-implant crevicular fluid (PICF)
Similar to gingival crevicular fluid (GCF), peri-implant crevicular fluid (PICF)
can be collected from sulcus adjacent to implants. GCF/PICF is presented as serum
transudate or more commonly inflammatory exudates from the periodontium (Lamster &
Ahlo, 2007). Peri-implant crevicular fluid is closely resembles gingival crevicular fluid
(GCF) in term of its composition as well as its mechanism of production (Adongianaki et
al., 1995).
Similar to the relationship seen in GCF and periodontitis, numerous studies have
linked an increase of inflammatory markers and specific enzymes in PICF to adverse
peri-implant conditions. Examples include, interleukin 1β (Curtis et al., 1997), matrix
metalloproteinases-8 (Teronen et al., 1997), and prostaglandin E2 (Salcetti et al., 1997).
Niimi and Ueda (1995) measured the volume of PICF around osseointegrated implants
and found that PICF volume was larger in implants with high gingival and plaque indexes.
19

However, the search for a sensitive diagnostic marker to detect peri-implant disease still
continues.
Similar to GCF, PICF contains components of the host epithelial and connective
tissues, substances from subgingival or supragingival microbial plaque, inflammatory and
immune cells and inflammatory markers (Lamster & Ahlo, 2007 and Curtis et al., 1997).
The total protein content in crevicular fluid appears to be closely resemble that of serum
(Bang & Cimasoni, 1971) suggesting an inflammatory exudative nature to the fluid.
Higher serum derived proteins were also isolated from the gingival crevicular fluid from
inflamed sites as observed by Curtis et al. (1988). For the purpose of the present study,
bovine serum was used to simulate the effect of the serum-like peri-implant crevicular
fluid.

Removal of bacteria on Implant Surfaces
Elimination of bacteria from the implant surface is necessary in order to terminate
the source of infection and disrupt the formation of biofilm. This aspect is important
when a peri-implant pathology of an infectious origin is suspected.
Numberous methods of decontaminating implant surfaces have been suggested.
Jovanovic et al. suggested in 1993 a “decontamination protocol” which consists of
exposure of titanium surfaces to a high-pressure sodium bicarbonate device for one
20

minute under aseptic conditions. Pereira da Silva et al. in 2005 tested this method of
decontamination and found no viable bacteria in all three titanium groups.
Augthun’s study (1998) also reported an air abrasive system to be the most
effective decontamination method and resulted in no damage to the underlying titanium
surface. However, Chairay et al. (1997) demonstrated altered morphology of machined
implants after administration of air powder abrasion. This difference was possibly due to
differences in duration of application, as Augthun’s group only applied the air abrasive
for 60 seconds. Besides altering implant surfaces, additional concerns such as possible
retained particles after administration and application of compressed air intra-orally may
raise some concerns toward this treatment modality.
Laser therapy had been suggested for the decontamination of implant surface.
However, studies show that not all types of lasers are suitable for this purpose.
Subsequent damage to the implant surface has been implicated in many cases. For
instance, the Nd:YAG laser was reported to cause pitting on implant surfaces in certain
settings (Bida, 1991 ) and also resulted in melting, loss of porosity and surface alteration
of plasma coated implants (Block et al., 1992). In addition, Block et al. reported failure
of sterilization by the Nd:YAG laser after contamination by spores of Bacillus subtilis.
In Duarte et al. (2009), the Er:YAG laser was not effective in removing S.
sanguinis. Stübinger et al. 2010 demonstrated surface alterations of different degrees
based on various energy settings of Er:YAG laser on both surfaces. No visible surface
21

alterations were seen when irradiated by CO
2
and diode lasers. The Er:YAG treated SLA
surfaces appeared to decrease in roughness due to melting of surface peaks, while smooth
surfaces increased in roughness as a results of developing cracks after irradiation. CO
2
and diode laser treated surfaces were not tested for surface roughness due to lack of
visible alteration.
Deppe et al. (2001) reported thermal changes as well as surface melting and
alteration by using CO
2
lasers in different settings. No data was reported on the efficacy
of bacteria removal even though the in vivo part of the study appeared successful. The
application of laser and its negative effects remains uncertain and further research will be
needed in this area before any solid conclusions can be drawn.
Persson et al. (1999) conducted an in vivo study on beagle dogs utilizing a
decontamination method of a rotating brush with pumice and cotton soaked in saline.
The rotating brush was used to polish the surface until a “gray and frosty” appearance
was noted and cotton dipped in saline was used to clean the surface of implant until no
visible plaque was noted. The histological parameter of “re-osseointegration” did not
reflect any advantage of either treatment; microbial parameters were not tested in this
study. In addition, systemic antimicrobial therapy was administered to the dogs two days
prior to the decontamination therapy and continued for 3 weeks. From Persson’s study,
no information on the direct effect of either decontamination therapy on the bacteria can
be extracted. The effect of “gray frosted” surface on bacterial adhesion will require more
studies.
22

The most direct method of physically removing bacteria, plaque or calculus from
a surface in the oral cavity is the use of the curette. However the potential damaging
effect of a curette to the titanium surface is of great concern. Augthun et al. (1998) found
that the usage of a metal curette, diamond polishing device, and ultrasonic scalers all
resulted in damage to the implant surface.
Mengel et al. (1998) advised the use of a plastic curette, prophy tips and air
abrasive systems because the metal curettes, sonic and ultrasonic devices with universal
tips cause pronounced traces or instrumentation and remove substantial substance from
titanium.
Due to the potential negative effect of the above mentioned implant
decontamination methods, more conservative methods have been explored. In a dog
experiment by Persson et al. in 2001 with ligature induced peri-implantitis, implant
surfaces were meticulously cleaned with cotton soaked in saline along with flap elevation.
Decontamination appeared to be successful due to the high percentage of “re-
osseointegration.” Once again, systemic antibiotic therapy was administered to the dogs
for 17 days in this study. The authors therefore concluded that systemic antibiotics and
granulation tissue removal along with implant surface cleaning (with saline) result in
resolution of peri-implant lesion and bone fill. However, the fact that this is a dog model
with ligature induced peri-implantitis may raise some questions about its actual effect in
human. In addition, no direct proof of complete removal of bacteria was provided. It is
23

unclear whether the systemic antibiotics, degranulation during flap surgery or the actual
surface cleaning of saline decontaminated and promoted the implant site healing.
Schou et al. (2003) tested four methods of decontamination: 1) air-powder
abrasive, 2) air- powder abrasive follow by citric acid, 3) gauze socked in saline
following citric acid and 4) gauze soaked in 0.1% chlorhexidine and saline alternatively.
Following decontamination of implant surfaces, peri-implantitis defects were treated with
autogenous particulate bone fill and covered by e-PTFE membrane. Positive results were
obtained and the authors recommended the decontamination of soaked gauze with
chlorhexidine and saline. Once again the bacterial aspect of results was not discussed
and many variables were included in the study.
Rinsing of saline and chlorhexidine was described by Hammerle et al. (1995).
Sites of peri-implantitis were treated with flap debridement along with rinsing of sterile
and 0.2% chlorhexidine digluconate. Authors reported positive results with bone gain
with this method of decontamination along with guided tissue regeneration. The sole
effect of rinsing of saline was not tested in this study.
Dennison et al. (1994) conducted an experiment on the removal of bacterial
endotoxin by means of burnishing implants with cotton pellet prepared with water, citric
acid, or 0.12% CHX and air-powder abrasive. Three types of implants were used:
machined, plasma sprayed and hydroxyapatite coated implants. An air abrasion system
appeared to be effective in decontamination of P.gingivalis endotoxin in all implant types.
24

Both air abrasion and citric acid were effective in decontamination of hydroxyapatite
coated implants. For machined surface implants, all treatments including cotton pellet
soaked in water can be effective in removal of endotoxin. One can speculate from this
study that the application of citric acid or CHX did not provide additional benefit. In
addition, the use of acid in decontamination of titanium surface could be alarming
because acids are used for surface modification of titanium.
Water rinsing is a non-invasive and inexpensive way to remove biofilm on teeth
(Gorur et al., 2009). Studies on the efficacy of water rinsing on different titanium
surfaces have not been conducted. If proven effective, rinsing titanium implants with
sterile saline water with an easily accessible syringe device could be a simple and
inexpensive way of treatment if proven to be effective.

The purpose of this study is to test the ability of P. micra to adhere to implant
surfaces that are newly developed in an in vitro setting. To the best of our knowledge,
this is a pilot study on P. micra since no other studies of P. micra adhering on titanium
surfaces could be identified in the literatures. The following hypotheses are generated to
answer the questions of bacterial adhesion on implant surfaces.
1. P. micra is capable of adhering to all implant surfaces
2. Rough surfaced implants currently on the market have significant higher bacterial
adhesion rate when compared to smooth surfaced implants.
25

3. Anaerobic culturing condition and nutrient broth provide P. micra a favorable
situation for promoting adhesion and growth.
4. Serum, which simulates the peri-implant crevicular fluid in the oral cavity, can
alter P. micra behavior in adhesion and proliferation on titanium surface.
5. Saline irrigation with irrigation syringe is capable of removing P. micra from
titanium implants.
26

Chapter 2: Materials and Methods
Seven types of implants were selected for this study based on their surface
characteristics and manufacturers: Nanotite implants (Biomet 3i, Palm Beach Gardens,
Florida, USA), Osseotite implants (Biomet 3i, Palm Beach Gardens, Florida, USA), 3i
machined surface implants (Biomet 3i, Palm Beach Gardens, Florida, USA), Implantium
anodized surface implants (Dentium, Seoul, Korea), Implantium SLA+ implants
(Dentium, Seoul, Korea), Branemark TiUnite implants (Nobel Biocare, Zurich,
Switzerland), Branemark machined surface implants(Nobel Biocare, Zurich, Switzerland).
Surface treatments and manufacturer information were listed in table 1.

Table 1. Listing of implants, manufacturer and surface treatment description
Type of Implant Manufacturer Surface treatment
1 Nanotite Implant Biomet 3i Dual acid etched and coating of
calcium phosphate
2 Osseotite implant Biomet 3i Dual acid-etched surface
3 3i machined surfaced implant Biomet 3i Machined surface
4 Implantium anodized surface implant Dentium Anodized surface
5 Implantium SLA+ implant Dentium Sand-blasted acid etched surface
coated with calcium phosphate
6 Branemark TiUnite implant Nobel Biocare Anodized surface
7 Branemark machined surfaced
implant
Nobel Biocare Machined surface

27

Control Group
Seven implants of different types were used as controls and scanning electron
microscopic photographs were taken for comparison to the test groups.

Experiment Groups
Experimental implants were divided into three groups, A, B, and C; each
consisted of 21 implants (3 sets of seven different implants). Diluted P. micra culture in
phosphate buffered saline was prepared. Implants were submerged in the bacterial broth
4 hours (group A), 24 hours (group B) in atmospheric condition and 4 days under
anaerobic condition in nutrient broth (group C). Group A, B, and C were further divided
into 3 subgroups, 1, 2, and 3. Subgroups A1, B1, and C1 were incubated with bacteria
only. For subgroup A2, B2, and C2, implants were pre-treated with bovine serum before
submersion into the bacterial broth. As for subgroup A3, B3, and C3, implants were pre-
treated with bovine serum as in subgroup 2, and then irrigated with saline after incubation.
The experiment design is outlined in flowchart 1.

28

Flowchart 1. Experiment design

29

Implants Information
Implants used in this study are listed in table 2 with detailed information on code, length,
reference, lot number, manufacturing date and expiration dates if provided on the
packaging of implants.

Table 2. Specific implant information
Group Implant Code Length REF LOT Manufactured
date
Expiration
date
Control
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite OSS 4x13mm OSS413 656964-5 2012-03
3i
Machined
Custom
#7270
4x10mm ICE410 774247 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 07J02-R 2007-12-11 2012-12-
10
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
MkIV TiU 4x11.5mm
RP
28935 664013 2011-02
Branemark
machined
MkIII 4x15mm RP 28895 637944 2008-01
A1
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite OSS 4x13mm OSS413 653896 2012-03
3i
Machined
Custom
#7270
4x13mm ICE413 774249 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 07J02-R 2007-12-11 2012-12-
10
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
MkIII, TiU 4x11.5mm
RP
28921 669978 2011-08
Branemark
machined
MKIII 4x8.5mm RP 28891 645078 2009-01
30

Table 2. Specific implant information (continued)
A2
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite OSS 4x13mm OSS413 461593 2010-12
3i
Machined
Custom
#7270
4x10mm ICE410 774247 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 07J02-R 2007-12-11 2012-12-
10
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
MkIII, TiU 4x11.5mm
RP
28921 669978 2011-08
Branemark
machined
MkIII 4x8.5mm RP 28891 645078 2009-01
A3
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite OSS 4x13mm OSS413 656964-5 2012-03
3i
Machined
Custom
#7270
4x13mm ICE413 774249 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 07J02-R 2007-12-11 2012-12-
10
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
MK IV
TiUnite
4.0x15mm
RP
28937 654718 2010-03
Branemark
machined
MkIII 4x8.5mm RP 28891 645078 2009-01
B1
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite NT CERT 4x11.5mm INT511 231673 2004-2 2009-2
3i
Machined
Custom
#7270
4x10mm ICE410 774247 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 08B01-R 2008-02-20 2013-02-
19
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
Mk III TiU 3.75x15mm
RP
28937 651927 2009-12
31

Table 2. Specific implant information (continued)
Branemark
machined
MkIII 4x8.5mm RP 28891 645078 2009-01
B2
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite NT CERT 5x13mm INT13 231338 2004-2 2009-2
3i
Machined
Custom
#7270
4x10mm ICE410 774247 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 07J02-R 2007-12-11 2012-12-
10
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
Fixture
TiUnite
MkIII
5x8.5mm
WP
27396 625699 2001-01
Branemark
machined
MkIII RP 4x15mm RP 28895 650409 2009-09
B3
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite NT CERT 5x13mm INT513 231338 2004-2 2009-2
3i
Machined
Custom
#7270
4x10mm ICE410 774247 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 08B01-R 2008-02-20 2013-02-
19
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
Fixture
MkIII TiU
5x10mm
WP
27397 625701 2001-01
Branemark
machined
MkIII 4x8.5mm RP 28891 645078 2009-01
C1
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite NT CERT 5x11.5mm INT511 231673 2004-2 2009-2
3i
Machined
Custom
#7270
4x11.5mm ICE411 774248 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 08B01-R 2008-02-20 2013-02-
19
32

Table 2. Specific implant information (continued)

Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
Nobel
Speedy
Groovy RP
4x15mm 32151 703260 2013-11
Branemark
machined
Fixture Pcs 1 5x12mm SDCA
147-0
616106 1999-03
C2
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite NT CERT 5x11.5mm INT511 231673 2009-2
3i
Machined
Custom
#7270
4x10mm ICE411 774248 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 08B01-R 2008-02-20 2013-02-
19
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
Nobel
Speedy
Groovy RP
4x15mm 32151 703260 2013-11
Branemark
machined
RP, fixture
MkIII
4x13mm 26969 621608 2000-04
C3
Nanotite NANO EX
HX
4x11.5mm NOSS411 687179 2012-05
Osseotite NT CERT 5x11.5mm INT511 231673 2004-2 2009-2
3i
Machined
Custom
#7270
4x11.5mm ICE411 774248 2012-12
Implantium
anodized
FX4310A 4.3x10mm 05-1014 08B01-R 2008-02-20 2013-02-
19
Implantium
SLA+
FX4310 4.3x10mm 02-662 07J02-R 2007-12-12 2010-12-
11
Branemark
TiUnite
Nobel
Speedy
Groovy RP
4x15mm 32151 703260 2013-11
Branemark
machined
Mk III
(Pcs1)
4x13mm 26969 623789 2000-09
33

Culture of Parvimonas micra
Lypolized P. micra (ATCC: 33270, LOT: 958302, REF: 0958P) was purchased
from MicroBioLogics (St. Cloud, MN, USA). Instructions were followed to activate P.
micra. P. micra were streaked with sterile loop onto the BRU plates, which were
consisted of 1000 mg double distilled water, 2g yeast extract, 3g bacto agar, 45g brucella
agar. Microorganisms were plated and incubated with AnaeroGen (Basingstoke, UK)
sealed with incubation jar inside the anaerobic chamber (N
2
:H
2
:CO
2
= 85:10:5) at 36
degree Celsius for 4 days to obtain mature growth of bacteria.
Brucella agar: http://www.condalab.com/pdf/1012.pdf
Bacto agar: http://www.bd.com/ds/technicalCenter/inserts/Agars.pdf

Harvest of the bacteria
For group A and B, P. micra colonies were harvested by metal sterile loop and
transferred into phosphate buffered saline, attention was given not to remove any agar
material. Concentration of bacteria was diluted and calculated by using a bright line/dark
line cell counting chamber. The concentration of P. micra in the experiment was
determined to be 5.40x10
6
cells per milliliter (ML). 1.5 milliliter of bacterial solution
was transferred into each individual plastic vial and plastic cap was replaced after
designated implants were placed inside the vial. During the incubation time, samples
were agitated every hour to avoid precipitation of bacteria.
34

For group C, P. micra colonies were harvested by metal sterile loop into diluted
broth (1000 ml double distilled water, 2.5g thiotone E peptone, 2.5g tryptose, 5.0g
sodium chloride). Attention was given not to remove agar material while harvesting.
Concentration of P. micra was also determined by bright line/ dark line cell counting
chamber. The average concentration of P. micra in the broth was determined to be
5.40x10
6
cells per milliliter (ML). 1.5 milliliter of bacterial solution was transferred into
each individual plastic vial and plastic cap was replaced after designated implants were
placed inside the vial.

Figure 1. Plastic vial containing different implants

Pre-treatment with bovine serum
Liquid ImmunO sterile filtered bovine serum by MP Biomedicals, LLC (Solon,
Ohio, USA) was poured into a sterile container, Implants designated to serum treatment
35

were submerged into bovine serum for one minute to ensure saturation of serum on the
implant surface. According to the manufacturer’s information, ImmuO sterile bovine
serum consists of hemoglobin: 12.3 mg/dl, total protein: 7.1 g/dl, pH: 7.4. osmolality:
300 mOsm/kg.

Irrigation with saline
Irrigation of sterile saline was performed using a US-12 utility 12 c.c. curved tip
syringe (Vista Dental Product, Racine, WI, USA) with 1 inch distance from the implants
for one minute. Implants were held in place by sterile cotton pliers during irrigation and
were rotated slowly to ensure all surfaces had been irrigated.

Figure 2. US-12 utility 12 c.c. curved tip syringe

36

Fixation and Scanning Electron Microscopy
All implants were then immediately fixed in Kanovsky fixative (2 gm
paraformaldehyde and 25 mL distilled water) inside a individual clean plastic vial, and
after designated processing, rinsed in cacodylate buffer, and placed in a graded series of
ethanol for dehydration. Then the samples were further dehydrated in the
hexamethyldisilazane and left air dried. After the gold coating process of the samples,
they were ready to be observed with a Cambridge 360 Scanning Electron Microscope
(Cambridge Instruments, Cambridge, UK) and JOEL JSM-6610 scanning electron
microscope machine (JOEL Ltd, Tokyo, Japan). Corresponding computer programs
provided by the manufacturers were used to examine and capture images from the
microscope.
A set of 7 implants without any treatment were processed and examined under
SEM as control samples. Photographs were also taken for the comparison to the
experimental implants.
Implants were divided into three sections: head, middle and tip. Two photographs
at two different magnification (x5000 and x20,000) were taken at each section of
implants. The selections were random. A total of six photographs were taken of each
implant. For Osseotite implants, the machined surface collar was not included in the
SEM images.
37

Grids were superimposed onto each photograph of 5000 magnification to count
the bacteria and calculate the percentage of bacterial adhesion per photo area. An
example of a photograph with grid is shown in Figure 3. Grid was considered occupied
if part of a single bacterium was inside the grid. Total of grid occupied by bacteria is
then divided by the total number of grids presented in the photograph to calculate the
percentage of bacterial adhesion in each photograph.

Figure 3. Example of x5000 magnification of image with counting grids superimposed.

38

Analysis
Effect of incubation environment and duration, serum, and saline rinsing were
analyzed by comparing the average percentage of bacteria adhesion. Statistical analysis
was conducted. Average percentage of adhesion rates were compared among different
implants.

Grouping and Comparison
Implants were further grouped based upon their surface modifications and the
average percentages of bacteria adhesion were compared. Table 3 listed all the implant
groups:

Table 3. Implant groups and implants included
Group Implants included
Machined Surface 3i machined, Branemark machined
Rough Surface Osseotite, Nanotite, Implantium anodized,
Implantium SLA+, Branemark TiUnite
Acid etched Osseotite, Nanotite, Implantium SLA+
Anodized Implantium anodized, Branemark TiUnite
3i Brand 3i machined surface, Osseotite, Nanotite,
Implantium Brand Implantium anodized, Implantium SLA+
Branemark Brand Branemark machined, Branemark TiUnite
Calcium phosphate
coated
Nanotite, Implantium SLA+
Non-calcium phosphate
coated
3i machined, Osseotite, Implantium anodized,
Branemark machined, Branemark TiUnite

39

Statistical Analysis
Comparisons between different implants under different conditions were made.
When more than two groups were compared, all p-values were calculated by utilizing
Kruskal-Wallis test (non-parametric equivalent of ANOVA). When two groups were
compared, Wilcoxon rank sum tests (the non-parametric equivalent of independent
sample t-tests) was used.
40

Chapter 3: Results
Control
A set of 7 implants were examined under SEM without any treatment, they served
as controls. The control images are listed as follow:
Figure 4. 3i machined implant (x5000). No surface irregularities noted, only striation
observed.

41

Figure 5. Branemark machine implant (x5000). Similar appearance to 3i machine
implant surface, only striations observed.

Figure 6. Osseotite implant (x5000). Characterized by the sharp peak and valleys

42

Figure 7. Osseotite implant (x20,000)

Figure 8. Nanotite implant (x5000). Presented with similar peaks and valleys as in
Osseotite implants, however, a layers of coating observed mostly at the trough area.

43

Figure 9. Nanotite implant (x20,000)

Figure 10. Implantium SLA+ implant (x5000). Surfaces were characterized by rounded
and thickened peak and troughs of unknown depth. Thick layer of deposits located at the
peak region.

44

Figure 11. Implantium SLA+ implant (x20,000).

Figure 12. Implantium anodized implant (x5000). Porous structures evenly distributed
among implant surfaces. Smooth surfaces observed between porous structures.

45

Figure 13. Implantium anodized implant (x20,000).

Figure 14. Branemark TiUnite implant (head x2000). Pores presented as volcanoes
rising upward. Surface pores are consist of different sizes, and unevenly distributed.
Mixture of larger raised pores and some flatter pores observed in the same area.

46

Figure 15. Branemark TiUnite implant (head x5000).

Figure 16. Branemark TiUnite implant (middle x2000). Loosely distributed pores were
observed in the more apical portion of TiUnite implant.

47

Figure 17. Branemark TiUnite implant (middle x5,000).

Statistical analysis
All p-values were calculated using Kruskal-Wallis tests (non-parametric
equivalent of ANOVA) when more than 2 groups were compared, or Wilcoxon rank sum
tests (non-parametric equivalent of independent sample t-tests) when 2 groups were
compared. Post-hoc analyses of significant Kruskal-Wallis test results found no
significant differences between pairs of groups after a multiple comparison adjustment.

Bacterial adhesion on surfaces
The hypothesis that P. micra was capable of adhering to all implant surfaces was
supported by the results of current study. Even though percentage of P. micra adhesion
48

varied among different groups, all implant surfaces showed P. micra adhesion regardless.
Table 4 presents a record of P. micra adhesions in raw percentage data.

Table 4. Percentage of P. micra adhesion on different implants among different treatment
groups

% of Bacterial Adhesion
Group Type of Implants Head Middle Tip Average
A1
Nanotite 0.0 51.2 17.9 23.0
Osseotite 34.5 2.4 23.8 20.2
3i Machined 45.2 29.8 9.5 28.2
Implantium anodized 46.4 17.9 16.7 27.0
Implantium SLA+ 0.0 6.0 23.8 9.9
Branemark TiUite 11.9 20.2 21.4 17.9
Branemark Machined 33.3 27.4 45.2 35.3
A2
Nanotite 15.5 14.3 0.0 9.9
Osseotite 0.0 0.0 0.0 0.0
3i Machined 0.0 0.0 0.0 0.0
Implantium anodized 4.8 0.0 0.0 1.6
Implantium SLA+ 0.0 0.0 1.2 0.4
49

Table 4. Percentage of P. micra adhesion on different implants among different treatment
groups (continued)
Branemark TiUite 1.2 1.2 0.0 0.8
Branemark Machined 33.3 75.0 41.7 50.0
A3
Nanotite 2.4 1.2 3.6 2.4
Osseotite 0.0 0.0 0.0 0.0
3i Machined 0.0 0.0 0.0 0.0
Implantium anodized 0.0 0.0 0.0 0.0
Implantium SLA+ 0.0 0.0 0.0 0.0
Branemark TiUite 0.0 3.6 3.6 2.4
Branemark Machined 0.0 0.0 0.0 0.0
B1
Nanotite 100.0 100.0 100.0 100.0
Osseotite 0.0 100.0 100.0 66.7
3i Machined 6.0 0.0 10.7 5.6
Implantium anodized 0.0 27.4 2.4 9.9
Implantium SLA+ 0.0 0.0 0.0 0.0
Branemark TiUite 6.0 1.2 1.2 2.8
Branemark Machined 4.8 0.0 0.0 1.6
B2
Nanotite 100.0 100.0 100.0 100.0
Osseotite 100.0 100.0 100.0 100.0
50

Table 4. Percentage of P. micra adhesion on different implants among different treatment
groups (continued)
3i Machined 100.0 0.0 0.0 33.3
Implantium anodized 2.4 4.8 0.0 2.4
Implantium SLA+ 0.0 0.0 0.0 0.0
Branemark TiUite 2.4 4.8 1.2 2.8
Branemark Machined 0.0 0.0 100.0 33.3
B3
Nanotite 100.0 100.0 100.0 100.0
Osseotite 0.0 0.0 0.0 0.0
3i Machined 0.0 0.0 0.0 0.0
Implantium anodized 4.8 0.0 3.6 2.8
Implantium SLA+ 0.0 0.0 0.0 0.0
Branemark TiUite 2.4 0.0 3.6 2.0
Branemark Machined 0.0 0.0 0.0 0.0
C1
Nanotite 100.0 26.1 5.4 43.8
Osseotite 29.3 10.9 4.3 14.9
3i Machined 16.3 12.0 6.5 11.6
Implantium anodized 12.0 20.7 10.9 14.5
Implantium SLA+ 9.8 4.3 18.5 10.9
Branemark TiUite 6.5 12.0 14.1 10.9
Branemark Machined 9.8 7.6 30.4 15.9
51

Table 4. Percentage of P. micra adhesion on different implants among different treatment
groups (continued)
C2
Nanotite 0.0 0.0 0.0 0.0
Osseotite 2.2 3.3 1.1 2.2
3i Machined 1.1 0.0 0.0 0.4
Implantium anodized 0.0 0.0 3.3 1.1
Implantium SLA+ 8.7 16.3 7.6 10.9
Branemark TiUite 2.2 1.1 2.2 1.8
Branemark Machined 17.4 14.1 0.0 10.5
C3
Nanotite 0.0 0.0 0.0 0.0
Osseotite 0.0 2.2 1.1 1.1
3i Machined 0.0 1.1 0.0 0.4
Implantium anodized 2.2 1.1 0.0 1.1
Implantium SLA+ 6.5 26.1 0.0 10.9
Branemark TiUite 0.0 1.1 3.3 1.4
Branemark Machined 0.0 3.3 1.1 1.4
*Raw percentage calculated from the grid occupied by bacteria divided by total number of grids

52

Table 5. Percentage of P. micra adhesion in all groups.
Group Head Middle Tip Average
A1 24.5 (20.2) 22.1 (16.3) 22.6 (11.1) 23.1 ( 8.2)
A2 7.8 (12.6) 12.9 (27.9) 6.1 (15.7) 9.0 (18.4)
A3 0.3 ( 0.9) 0.7 ( 1.3) 1.0 ( 1.7) 0.7 ( 1.2)

B1 16.7 (36.9) 32.7 (47.1) 30.6 (47.5) 26.6 (40.0)
B2 43.5 (52.8) 29.9 (47.9) 43.0 (53.3) 38.8 (44.1)
B3 15.3 (37.4) 14.3 (37.8) 15.3 (37.4) 15.0 (37.5)

C1 26.2 (33.4) 13.4 ( 7.5) 12.9 ( 9.3) 17.5 (11.8)
C2 4.5 ( 6.4) 5.0 ( 7.1) 2.0 ( 2.8) 3.8 ( 4.7)
C3 1.2 ( 2.5) 5.0 ( 9.4) 0.8 ( 1.2) 2.3 ( 3.8)
*Average (standard deviation)

53

Effect of surface modifications
Table 6. Average adhesion percentage of P. micra in different groups. Average
adhesion percentage from all experimental groups when grouped into surface treatment,
brand name, presence of calcium phosphate, machined surfaces and rough surfaces.
Average % of P. micra adhesion
Surface treatment
Acid Etched 23.2 (35.9)
Machined 12.6 (16.1)
Anodized 5.7 (7.4)
p-value 0.96
Brand name
Biomet 3i 24.6 (35.8)
Branemark 10.6 (14.7)
Implantium 5.7 (7.3)
p-value 0.43
Presence of Calcium
Phosphate

Calcium Phosphate 23.5 (36.9)
Non-Calcium Phosphate 11.9 (19.9)
p-value 0.81
Machined vs. Rough
Machined 12.6 (16.1)
rough Surfaced 16.2 (29.3)
p-value 0.89
*Mean (standard deviation)
54

The overall p-values were insignificant between all groups. The percentage of
adhesion for P. micra was higher in rough surfaces than in machined surfaces, 16.2% and
12.2% respectively.
When different surface modification methods were compared, acid etched group
had higher average of bacterial adhesion (23.2%) when compared to machined (12.6%)
and anodized surface (5.7%).
When implants were grouped according to the brand name, Biomet 3i, Branemark,
and Implantium, Biomet 3i implants (24.5%) had much higher average of bacterial
adhesion compared to Branemark and Implantium (10.6% and 5.7%, respectively).
The effect of calcium phosphate was also analyzed. Implants were grouped into
calcium phosphate treated group and non-calcium phosphate treated group. Average
bacterial adhesion percentage for implants with calcium phosphate was 23.4% which is
almost twice as high as the non-calcium phosphate treated implants (11.9%).
Comparisons were made between individual implants in all groups and table 7
listed the percentage of P. micra adhesion.

55

Table 7. Percentage of P. micra adhesion in all implants
Effect of
implant
Head Middle Tip Average
In all groups Nanotite 46.4(51.0) 43.6(45.2) 36.3(48.1) 42.1(45.5)
Osseotite 18.4(33.5) 24.3(43.0) 25.6(42.9) 22.8(36.1)
Implantium
anodized
8.1(14.9) 8.0(10.9) 4.1(5.9) 6.7(9.0)
Implantium
SLA+
2.8(4.2) 5.9(9.3) 5.7(9.2) 4.8(5.6)
3i Machined 18.7(33.9) 4.8(10.2) 3.0(4.6) 8.8(13.1)
Branemark
TiUite
3.6(3.9) 5.0(6.8) 5.6(7.2) 4.7(5.8)
Branemark
Machined
11.0(14.0) 14.2(24.6) 24.3(34.3) 16.5(18.7)
p-value .64 .28 .53 .47

No statistical differences were found between different implant types, as shown
in Table 7. Nonetheless, Nanotite implants had the highest adhesion percentage (42.1%),
followed by Osseotite (22.8%). Anodized surface had the least amount of bacterial
adhesion. Implantium anodized implants presented with 6.7% and Branemark TiUnite
with 4.7% which was the lowest amongst all implants.
Similar implant surfaces were compared among different companies. 3i
machined implants had an average of 8.8% of adhesion rate while Branemark machined
implants were presented with 16.5% which was much higher.
56

When calcium phosphate deposits were taken into consideration, Nanotite
presented with an average of 42.1% and Implantium SLA+ with 4.8%.

Effect of aerobic and anaerobic condition
Analysis from post-hoc analyses of significant Kruskal-Wallis tests results
showed no significant difference between pairs of group after a multiple comparison
adjustment.
However, P. micra was capable of adhering to implant surfaces under aerobic
condition up to 24 hours as well as 4 days under anaerobic condition. Interestingly,
anaerobic condition with nutrient broth (group C) did not increase the adhesion
percentage of P. micra. Group B (with 24 hour aerobic environment) presented with the
highest average adhesion percentage of 26.8% when comparing to group A and C (10.9%
and 7.9%). The percentage of adhesion in all three groups is listed in Table 8.

57

Table 8. Percentage of P. micra adhesion in group A, B, and C
% of Adhesion Head Middle Tip Average
group A 10.9(16.6) 11.9(19.9) 9.9(14.2) 10.9(14.6)
group B 25.2(42.9) 25.6(43.0) 29.6(45.7) 26.8(39.8)
group C 10.7(21.9) 7.8(8.7) 5.2(7.7) 7.9(10.1)
p-value .20 .10 .015 .04
*Mean (Standard deviation)

Effect of Serum on P. micra adhesion
The effect of serum appeared inconsistent throughout the experiment. From the
data presented in Table 5, serum treatment prior to incubation decreased the percentage
of bacterial adhesion in both group A and C. However, serum was not effective in group
B.

58

Figure 18 . Effect of serum in group A, B, and C.

Effect of Saline Rinsing on Removal of P. micra Adhesion
Saline rinsing appeared to have the ability to decrease bacterial adhesion in all
groups, especially in group A and B, 9% to 0.7% and 38.8% to 15%, respectively (Table
5). However, complete elimination of bacteria was not observed in many cases.
Group A
When individual implants were taken into account, figure 19 demonstrated the
effect with and without saline rinsing (A2 and A3). Branemark machined implants
showed large reduction of bacterial adhesion percentage (50% to 0%). Reduction in
bacterial adhesion was also noted in Nanotite implants (9.9% to 2.4%).

59

Figure 19. Effect of saline rinsing in group A

Group B
Within group B, the percentage of P. micra adhesion before and after rinsing was
recorded in figure 20. Nanotite implant showed no reduction of bacterial adhesion after
saline rinsing. On the other hand, complete removal of P. micra was observed for
Osseotite, 3i machined and Branemark machined implants. Figure 21 through 28 showed
SEM images of various implants in group B before and after saline rinsing.

60

Figure 20. Effect of saline rinsing in group B.

61

Figure 21. Group B2 3i machined implant (head portion) without saline rinsing (x5000)

Figure 22. Group B3 3i machined surface implant (head portion) with saline rinsing
(x5000)

62

Figure 23. Group B2 Branemark machined implant (tip portion) without saline rinsing
(x5000)

Figure 24. Group B3 Branemark machined implant (tip portion) with saline rinsing
(x5000)

63

Figure 25. Group B2 Nanotite implant (head portion) without saline rinsing. (x5000)

Figure 26. Group B2 Nanotite implant (head potion) without saline rinsing (x20,000).

64

Figure 27. Group B3 Nanotite implant (head portion) with saline rinsing (x5000)

Figure 28. Group B3 Nanotite implant (head portion) with saline rinsing (x20,000)

65

Group C
Within group C, the effect of saline rinsing was demonstrated in figure 29. No
bacterial adhesion was observed on Nanotite implant after saline rinsing; however, large
amount of debris persisted after saline rinsing (Figure 30). A slight increase in bacteria
adhesion was noted on Implantium SLA+ implant after rinsing. Other implant types had
insignificant changes in percentage of adhesion.

Figure 29. Effect of saline rinsing in group C.

66

Figure 30. Group C3 Nanotite implant (middle section) after rinsing (x 5000)

67

Description of P. micra Adhesion on Implants
Figure 31. Group A1 head portion of 3i machined implant (x5000). P. micra presented
in a cluster or chain of 4-10 bacteria. Bacteria appeared to adhere only at the base.
Bacteria retained their shape as adhering to the surface.

Figure 32. Group A1 head portion of 3i machined implant (x20,000).

68

Figure 33. Group A1 middle portion of Nanotite implant (x5000). Adhesion of P. micra
appeared to be on the peak of the etched surface. Coating of calcium phosphate was still
visible at the trough area. P. micra presented in chains or clusters.

Figure 34. Group B2 head portion of Nanotite implant (x5000)

69

Figure 35. Group B2 head portion of Nanotite implant (x20,000). Thick bacteria coating
observed in the higher magnification, while peak and valley texture of the implant surface
can still be observed.

70

Figure 36. Group B2 head portion of Osseotite implant (x5000). Bacteria coating
masking the surface texture of implant, hint of original valleys and peaks still present.

Figure 37. Group B2 head portion of Osseotite implant (x20,000)
Thick coating of bacteria noted.

71

Figure 38. Group C1 head portion of Osseotite implant (x5000). Irregularities noted in
the surface topography and was very different from the control samples obtained. No
coating of bacteria observed, however P. micra appeared to adhere to the peak area where
the roughness was more prominent.

72

Figure 39. Group C1 head portion of Osseotite implant (x20,000). In a cluster, P. micra
may be evidently separated with defined borders or fused together. Noted the two fused
bacteria by the arrow, possibly in the process of dividing

73

Figure 40. Group A1 head portion of Implantium anodized implant (x5000). Most P.
micra adhere mostly on the top or inside portion surrounding the pore. Inside the depth
of pore cannot be visualized to detect the presence of bacteria.

74

Figure 41. Group A1 tip portion of Implantium anodized implant (x20,000). Note the
adhesion of single P. micra along the rim of the pore. Shadow of rounded object
observed inside the pore, however, a connection between the pore and the object was
noted.

75

Figure 42. Group A1 head portion of Implantium anodized implant (x5000)

Figure 43. Group C1 head portion of Implantium SLA+ implant (x5000)
P. micra appeared to adhere to peaks of the implant surface.

76

Figure 44. Group C1 head portion of Implantium SLA+ implant (x20,000). Individual
bacterium can be observed rather than clusters. Some bacteria were in the process of
division.

77

Figure 45. Group A1 head portion of Branemark machined implant (x5000). P. micra
distributed evenly throughout the surface. Bacteria appeared to be flat and presented with
increase contact surface area to the implant. The adhesion may be clustered or separately.

Figure 46. Group A1 head portion of Branemark machined implant (x20,000)

78

Figure 47. Group B2 tip portion of Branemark machined implant (x5000). When
coating of bacteria was observed on the implant surface, bacteria appeared to be
following the striation observed in the control.

79

Figure 48. Group A1 middle portion of TiUnite implant (x5000). Uneven distribution of
bacteria observed on the TiUnite surface. Majority of adhesion occurred on the side or
top of volcano shaped irregularities. None of the sample showed P. micra adhesion into
the volcano holes.

80

Figure 49. Group B1 tip portion of TiUnite implant (x5000). With direct view into the
porous structure on the surface, no bacteria noted inside the visible range of the pore.

81

Chapter 4: Discussion
To the best of our knowledge, this is the first study that has investigated P. micra
adhesion onto titanium surfaces in vitro with the aid of scanning electron microscopy. In
this experiment, P. micra was found to be capable of adhering to all implant surfaces
tested including both rough and smooth surfaces as hypothesized.

Rough surface and bacterial adhesion
Higher percentages of bacterial adhesion on rough surfaces have been previously
reported among various types of bacteria and this appeared to be a general trend
throughout the existing literature. In this study, an increased adhesion percentage was
noted in rough surfaces (smooth with 12% and rough with 16%). This phenomenon can
be explained by the sheltering effect provided by rough surfaces protecting the microbes
against shear forces, thus facilitating the transformation of reversible adhesion into
irreversible adhesion and subsequent biofilm formation (Quirynen and Bollen, 1995).
This result, as with many previous studies demonstrate the advantages of smooth surfaces
in decreasing bacterial adhesion.
Amoroso et al. (2006) investigated the adhesion of Porphyromonas gingivalis on
four different kinds of titanium surfaces, ranging from very smooth to very rough.
Titanium surfaces were roughened by sandblasting with glass beads or aluminum oxide
82

with pressure. They found significant reduction of P. gingivalis adhesion on the “very
smooth” surfaces compared to any other surfaces.
Yoshinari et al. (2000) used P. gingivalis and A. actinomycetemcomitans to
adhere to surface modified titanium in vitro. Titanium blasted group (Ra=0.47 µm,
blasted with 150 µm of titanium powder) showed significantly higher adhesion of both
bacteria when compared to titanium polished (Ra=0.07µm). The titanium striated group
(Ra=0.72 µm, striated with #320 SiC paper) also showed a higher adhesion of A.
actinomycetemcomitans than titanium polished group.
Pereira da Silva et al. in 2005 performed an experiment with smoothed surface
and 65µm and 250µm aluminum oxide blasted titanium surfaces, with Ra of 0.17µm,
1.14µm and 3.17µm, respectively. Streptococcus sanguis was incubated with the
modified titanium. The titanium specimens were vortexed in sterile saline, which were
then plated and incubated. Colony forming units were then counted. They found that the
amount of bacterial cells attached to titanium surface is directly correlated to the surface
roughness. 250µm aluminum oxide blasted titanium surfaces appeared to have the
highest count of bacterial cells (35x10
5
) when compared to smooth surface and 65µm
aluminum oxide blasted (49x10
3
and 11x10
4
cell counts respectively).
Wu-Yuan et al. in 1995 also conducted an experiment of bacterial attachment
onto various titanium surfaces. In this experiment, Streptococcus sanguis and
Actinomyces viscosus were selected due to their early subgingival colonization and
83

plaque development. P. gingivalis was selected for its frequent involvement in peri-
implantitis. Three types of titanium surfaces were created: smooth (polished), grooved
(hand ground with SiC papers), and rough (sandblasted with 50 micrometer of aluminum
oxide). The results found that S. sanguis and A. viscosus attached to rough surfaces more
often; however, P. gingivalis adhered to titanium surfaces regardless of their roughness.
They concluded that titanium surfaces influence the ability of bacterial adherence;
however, the degree of adherence may depend on the type of bacteria.
Drake et al. in 1999 tested S. sanguis adhesion to three different titanium surfaces:
polished with diamond paste of 1µm in size, polished with 800-grit silicon carbide
sandpaper and sandblasted. They found that rougher surfaces including 800-grit and
sandblasted surfaces presented with greater levels of bacterial colonization when
compared with 1µm smooth surfaces. They speculated that rough surfaces increased the
surface area for bacterial adhesion, or increased the wettability of titanium surface
promoting adhesion of bacteria. Their results showed no differences between the above
two rough surface groups. In addition, the authors confirmed that S. sanguis prefer to
colonize hydrophobic sites.
Clinically, Rams et al. in 1991 compared hydroxyapatite-coated implants placed
adjacent to pure titanium implants. Microbial samples were obtained 7-10 months after
prosthetic loading and examined by direct phase-contrast microscopy and plated for
culturing. In this study, although statistical significance was not reached, a higher mean
84

percentage of Parvimonas micra were found in the healthy hydroxyapatite-coated
fixtures (17.4%) than in the healthy pure titanium fixture (9.8%).
Quirynen et al. in 1993 compared the difference between roughened and standard
titanium abutments (Ra of 0.8 and 0.3µm) in humans. Twenty five times more bacteria
were harbored subgingivally on the rough abutments when compared to the standard ones.
On the other hand, rough abutments harbored fewer cocci supragingivally (an indication
of more mature plaque).
In supragingival studies on humans by Rimondini et al. (1997) the smoothest
titanium discs contained less bacteria than the roughest group. This was reflected by the
adherent microbial biomass and SEM observation.
Quirynen et al. (1996) studied four different roughnesses of titanium implants
intraorally in their patients. Their Ra values ranged from 0.05 to 0.21 µm. Subgingivally
after one month, only the two roughest abutments tested positive for spirochetes, a group
of periogenic bacteria also found in peri-implantitis. After three months, spirochetes
were still present on the roughest surface. Again more cocci were found on smoothed
surfaces, however the difference was not significant. Interestingly, the most frequently
isolated microorganisms were F. nucleatum and P. micra. This study also confirmed the
“threshold roughness” as an Ra of 0.2µm, in which any number lower than this threshold
(ie smoother surfaces) will no longer diminish bacterial adhesion/colonization.
85

Bollen et al. (1995) was the first to report the concept of a “threshold Ra” of
0.2µm where any surfaces with Ra values below this threshold can provide no further
reduction of microbial load. However from the available data, none of the implants used
in the present study had a known Ra lower than the threshold Ra.
Wennerberg et al. (2003) examined four types of titanium abutments intraorally in
ten patients for four week periods. The Sa value ranged from 0.3 to 1.87 µm and the
method of roughening the surface was blasting with aluminum oxide particles of various
sizes. Even though statistically insignificant, abutment surfaces blasted with the largest
particles (250µm) appeared to have the highest numbers of bleeding on probing.
Presence of plaque was also higher compared to other smoother surfaces.
The in vitro and in vivo studies presented above demonstrate that roughened
surfaces have the tendency to harbor more bacteria and increase plaque accumulation.
This is consistent with the results from the present study.

Acid etched surface and bacteria adhesion
In this study, implants processed with acid had higher P. micra adherence
percentage (23.2%) when compared to anodized or smooth implants (5.7%, and 12.6%,
respectively). This observation was also noted by Baldi et al. (2009). By using machined
abutment and Osseotite implants (3mm machined collar) and dual acid etched abutment
with full Osseotite (no machined collar) in eight patients, they found significantly higher
86

plaque accumulation on dual acid-etched surfaces. Higher amount of plaque
accumulation was also observed in the acid etched surfaces immediately after oral
hygiene sessions. This result was confirmed by both microbial testing as well as SEM
imaging. However, in this short term study (4 months after implant insertion), clinical
parameter such as bleeding on probing and histological analysis was not found to be
different in two surfaces.
Bürgers and associates (2010) used sand-blasted with acid-etched implant surface
and found higher bacterial adherence and plaque accumulation. In their study, sand-
blasted acid-etched and smooth titanium discs were bonded to the patient’s teeth for
twelve hours, higher percentage (8.5% v.s. 1.1%) of biofilm covering the sand-blasted
acid-etched titanium discs was found. On the other hand, when S. sanguinis was used to
access the adhesion on different surfaces, 11% of the sand-blasted acid-etched surface
was covered by S. sanguinis while only 3% of smooth surface was recovered. The
authors concluded that Ra values were of great influence on bacterial adhesion. This
observation coincides with the present study as acid treated implants appeared to have
higher adhesion percentage in this study. However, direct comparison cannot be made
since we did not have the same implants as presented in Bürger’s study.
Method of titanium alteration by means of acid etching with or without sand
blasting was shown to not only create different surface roughness (Ra) but also altered
the thickness of titanium oxide layers, surface contact angles, and pattern of protein
adherence (Sela, 2007). Acid etching appeared to be a method that modifies titanium
87

properties in addition to simply creating roughness by removal of titanium structure.
These property changes may have influence on the pattern of bacterial adhesion that will
need further investigation.

Anodized implants and bacterial adhesion
Anodized implants presented with a low percentage of P. micra adhesion both
individually and as a group in this study. This observation is in contrary to the
observation of Puckett et al. (2010). Puckett et al. created a nanotubular titanium surface
(anodization in 1.5% HF for 10 minutes), nanotexture titanium surface (anodization in
1.5% HF for 1 minute) and nanorough surface (electron beam evaporation). They found
the highest adherence of Staphylococcus aureus, Staphylococcus epidermidis, and
Pseudomonas aeruginosa on the nanotubular surface, followed by nanotexture surface
compared to nanorough and conventional titanium without alterations. A similar pattern
was observed for protein absorption. Puckett and colleagues in the same study also found
an increase in fluorine content on the surface of nanotexture and nanotubular which they
suspect to promote the bacterial and protein adhesion. This study demonstrated that the
type of acid solution used as well as the timing of anodization can significantly alter
surface properties. In our study, the exact timing of anodization was unclear in both
anodized implants used. In addition, TiUnite may use phosphoric acid instead of
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hydrofluoric acid to anodize, which may create different surface properties. However,
this may require further research to elucidate.
Anodized commercially pure titanium and titanium alloy were shown to have
photocatalytic bactericidal effect when exposed to near UV light of 350-380nm
(Muraleedharan et al., 2003 and Gopal et al., 2004). One of the crystalline forms of
titanium oxide, anatase, was known as a semiconductor with strong photocatalytic ability
(Gopal et al., 2004). Gopal et al. (2004) found decreased Micrococcus sp. and
Pseudomonas sp. adherence on anodized titanium surfaces when compared with dual acid
etched titanium with exposure to near UV light. They found gram negative Pseudomonas
sp. with lipoproteinaceous outer membrane to be more susceptible to the photocatalytic
effect while gram positive Micrococcus was not as susceptible. These bactericidal effects
on anodized surfaces may explain the results we obtained from our study.
According to Jarmar and co-workers in 2007, the majority of TiO
2
found on
TiUnite was anatase, which shown by Gopal et al. to have photocatalytic properties.
Also of note, the microbiology laboratory utilized in this experiment had abundant
exposure to sunlight, which includes the wavelengths known to activate the
photocatalytic ability of anatase. Therefore one cannot rule out the possibility that a
photocatalytic bactericidal effect created by titanium anatase and exposure to sunlight
occurred in our in vitro study. This may explain the decreased adherence of P. micra on
anodized implants in this study.
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Machined implants and bacterial adhesion
A surprisingly high percentage of P. micra adherence rate on machined implants
was noted in this study. The pattern of bacterial adhesion may to be related to the
striations noted on the surface (Figure 31, 32, 45, 46). Adhesion of bacteria onto smooth
titanium surface had been reported in many other studies. Kuula and associates (2004)
reported that P. gingivalis, P. intermedia and two F. nucleatum subspecies to be capable
of adhering to smooth surface titanium, however, the Ra values were not specified in that
study. Harris and Richards (2004) also reported the bacterial adherence onto
mechanically, electrically and chemically polished titanium with Ra values of 0.15, 0.18,
and 0.67µm. They reported the findings of scratches and grooves presented on the
smooth surfaces and suspected these irregularities provided the same sheltering effect of
the roughened surfaces. This is also suspected in our current findings as grooves and
striations were frequently noted in the SEM images.

Comparisons between same implant types by different manufacturers
In this study, differences in P. micra adhesion were found among two machined
surface implants from two companies. For Branemark machined implants, the SEM
images showed P. micra adhering to the surface in flattened forms with more surface area
contacting the implant while on 3i machined implants the bacteria appeared more
rounded with less contact area (Figure 31, 32, 45, 46, 47). The percentages of bacterial
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adhesion were different as well. Branemark presented with a higher P. micra adhesion
rate almost two times more than 3i machined surface (16.5% v.s. 8.8%). This variation
observed in these two machined surfaces could be due to the manufacturer’s polishing
method, the average roughness value, or differences in surface free energy.
The method of polishing may influence the surface and its ability to adhere
bacteria. Harris and Richards (2004) found that polished titanium surfaces did not
decrease the amount of Staphylococcus aureus adhesion. Chemically polished titanium
with a lower Ra (0.67) had higher adhesion rates when compared to other polishing
methods (electropolished and mechanically polished). The surface chemistry appeared to
be altered during chemical polishing. Both Biomet 3i and Branemark claim to have
machined surfaced implants, however, the material used to polish or cleanse these
implants may differ, possibly influencing their surface characteristics and subsequent
bacterial adhesion.
The definition of machined and turned surface requires further clarification in
many studies, as mentioned in a review by Wennerberg and Albrektsson (2010). The
same author in 2000 suggested a smooth surface to have a Sa value of < 0.5µm.
However, this is not consistently observed, as Branemark machined surface has a Sa of
0.9 µm and a Sdr of 40%, while 3i machined surface has values of 0.4 µm and 17%. In
other words, the Branemark smooth surface implants are considered rough according to
the guidelines suggested by Wennerberg and Albrektsson (2000). This may explain the
high P. micra adhesion percentage observed in this machined category.
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In addition, researchers also reported variations among smooth surfaces from
different companies. Lucchini et al. (1996) found that the mean roughness (Spa) of
NobelPharma machined implants and SERF machined implants were 0.4 µm and 0.26,
respectively. Minor differences were also observed in thread morphology.
Stout et al. (1990) described different methods of manufacturing “machined
implants.” Machined surfaces can be turned, polished, fly-cut, bored, electro-discharged,
end milled, ground, sand-blasted, or slap-milled.
The materials used to construct implants themselves may also vary. Harris and
Richards (2004) noticed differences between the two brands of commercially pure
titanium. They suspected that these surfaces were produced in different manufacturing
sites using methods with minor differences. All these differences in material and
processing methods may have altered the characteristics of the two machined implants
used in this study, perhaps providing another explanation for their differing results. In
addition, the difference in titanium properties may contribute to the difference in P. micra
adhesion among different brands assuming each company uses different sources of
commercially pure titanium.
Differences in surface free energy (SFE) may also explain the differences in
adhesion among same types of implants. The SFE of titanium smooth surfaces from
different companies could be different even though the topography may appear similarly
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on SEM images. This may be reflected by the morphological appearance and amount of
P. micra adhesion.
Generally speaking, the higher the surface free energy, the more adhesion of oral
bacteria can be observed due to the high SFE nature of oral bacteria (Quirynen and
Bollen, 1995). This phenomenon was reflected in Tsibouklis and associates’ study in
1999. They reported the low surface energy of smoothed surfaces decreased bacterial
adhesion.
Although SFE values were not tested in the present experiment, the values on 3i
machined and Branemark machined implants are suspected to be different. Branemark
machined implants may have a higher SFE compared to 3i due to the flatter appearance
of P. micra as well as the increased percentage of adhesion. Future research involving
measurements of SFE values on implant surfaces may be helpful in clarifying this issue.
Nevertheless effects of SFE appear to be much less significant when compared in
regard to surface roughness. Bürgers et al. (2010) studied the biofilm formation on two
different titanium implants, sand-blasted acid-etched titanium and machined titanium.
The sand-blasted, acid-etched titanium presented with a low SFE but a high Ra value.
However, their study showed high bacterial adhesion on the sand-blasted, acid etched
implants indicated the dominant effect of roughness over the effects of SFE. This same
phenomenon was also observed by Amoroso et al. (2006) and Quirynen et al. (1990).
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Differences in implant morphology were also noted when Implantium anodized
surfaces and Branemark TiUnite surfaces were compared. These two manufacturers
claim to use the same processing method; however, the appearances of two implants were
different in some details on SEM imaging (figure 12-15). The actual data on Sa and Sdr
for Implantium anodized surface cannot be located in the literature; therefore, the two
implants cannot be compared directly. Nevertheless, the bacterial adhesion percentages
were fairly low in both cases (Implantium anodized 6.7% and TiUnite 4.7%).
The relatively low rate of bacterial adhesion (4.7 % - 6.7 %) for either anodized
surface observed in this study is likely not directly related to surface roughness, Sa.
TiUnite implants presented with a much higher Sa value of 1.5 µm (Wennerberg and
Albrektsson 2010) when compared to Nanotite and Osseotite (0.5 µm and 0.68 µm,
respectively). The reason for this may be related to SFE values as mentioned earlier or it
may be related to the anatomical features of the individual implants.
The presence of volcano-appearing pores were observed on both anodized
implants. These pores may harbor bacteria that cannot be visibly counted under SEM
imaging technique. The size of the pores (less or equal to 4µm) is sufficient for P. micra
(size ranges from 0.3 to 0.7µm) to enter and possibly replicate undiscovered. Implantium
anodized implants tended to have smaller pores and occasionally substances inside of
pores were noted (Figure 40-41). However, these substances cannot be confirmed to be P.
micra. In addition, these substances were usually found connected to the sidewall of the
pore, and most likely represent incomplete formation of pores during the manufacturing
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process. Deeper pores with wider openings of large size were noted in TiUnite (Figure
49) and visibility to the bottom of the pores was very poor to non-existent. Thus, there
may be a high probability of P. micra falling into and adhering to these un-evaluable
areas. These pores would provide perfect shelter to protect the P. micra from any
shearing forces present.

Inconsistency of reported roughness in implants
Differences in reported Sa values in many studies reflected the inconsistency of
implant surfaces manufactured. For Branemark machined implants, Wennerberg and
Albrektsson (2010) measured Sa of 0.9 µm. However, Jarmar et al. (2008) reported a Sa
of 1.53 µm. Jarmar et al. attribute this difference in Sa value to measurement from
different portions of the implant. On the other hand, TiUnite was measured to be 1.1 µm
in Wennerberg and Albretksson (2010). However it was reported to be 1.5 µm by Jamar
and co-workers (2008).
Clearly, variations exist even among the same implant type manufactured from
the same company. The manufacturing process may not be consistent and alterations of
the surface topography or chemistry may occur. This implant heterogeneity may cause
potential problems in evaluating bacterial adhesion as seen in this experiment and it may
contribute to inconsistencies in osseintegration of implants whose properties are
otherwise identical.
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Comparison between implantium SLA + implant and Nanotitie implants
Both Implantium SLA+ and Nanotite implants involve the addition of calcium
phosphate. Clearly Nanotite implant surfaces had a higher adhesion percentage (42.1%)
compared to Implantium SLA+ (4.8%). The likely explanation may be in the differences
of CaP application technique of ion-beam assisted deposition (IBAD) and discrete
crystalline deposition (DCD). These two relatively new techniques aimed to solve
previous problems of non-uniform coatings of CaP and adhesive failure in plasma
sprayed hydroxyapatite coated implants.
Implantium SLA + surface utilizes the ion-beam assisted deposition (IBAD)
technique which deposits a layer of CaP ranging from 20 nm to 1µm (Park et al., 2005
and Coelho et al., 2009a). The specific measurement for Implantium SLA+ CaP coatings
was 500nm as measured by surface profiler (Yoon, 2009). A trend of decreasing
roughness (measured by Ra value) was noted (Coelho et al., 2009a) after application of
IBAD (Ra decreased from 0.66µm to 0.54µm). This lowering of the Ra value may
decrease the potential for P. micra adhesion.
Morphologically, Implantium SLA+ possesses a coating covering the sharp edges
of the implant surface (Figure 10 and 11). One important point noted during the random
sampling of SEM images of Implantium SLA+ was that the CaP was absent in some
portions of the implant, appearing as uncoated surfaces. This is may be explained by two
scenarios. First, incomplete coating occurred with use of the IBAD method. Second,
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there could have been a rapid dissolution of CaP layer (Coelho et al., 2009b). However,
images of Implantium SLA+ implants with bare surfaces without CaP were found in
control samples; therefore, the first speculation proposed would be more plausible. The
impact of this finding on host healing after implant placement or its effects on bacterial
adhesion may need further investigation.
Nanotite implants utilize a different technique called discrete crystalline
deposition (DCD). DCD deposits nano-meter sized CaP onto a previously treated dual
etched surface. Morphologically, the sharp edges created by the acid etching were not
completely masked by the CaP particles. Thus, a more complex structure of CaP was
created that was separate from the Osseotite surface after the DCD processing (figure 8
and 9). While Mendez et al. (2007) suspected this complex surface structure promoted
bonding of bone, it may also promote the adhesion of P. micra.
In addition to rougher surfaces, Nanotite implants may have a higher surface free
energy that promotes the adhesion of P. micra. However, this is only speculation as no
measurement of surface free energy was conducted. Future research will be needed in
this area.
The effect of calcium on titanium surface and its interaction with bacteria had
been discussed. Yamashita et al. in 1991 showed that calcium ions contributed to P.
gingivalis adhesion. It is also been documented that albumin has a fairly high affinity for
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calcium, perhaps accelerating pellicle formation on calcium coated implants and
subsequent bacterial colonization.
In a previously mentioned study by Yoshinari et al. (2000), P. gingivalis adhesion
was related to the amount of calcium ion adsorption as well as surface energy, while A.
acetomycetemcomitans was not affected by surface energy. Both bacteria were found in
increased numbers on calcium implanted titanium and P. gingivalis adhesion was
decreased in alumina coated surfaces. The authors speculated that “the ease of bacterial
adhesion to titanium is related to the ease of osseointegration to titanium. “ In other
words, the ease with which calcium phosphate and serum proteins are adsorbed on a
titanium surface are the same properties that may lead to a more rapidly osseointegration
of titanium surfaces. In the oral cavity, calcium ions and pellicle in saliva are adsorbed
onto the negatively charged titanium surface which may promote oral bacteria adherence.
The authors thus believed that calcium rich surfaces may promote protein adsorption in
saliva as well as bacterial adhesion, while being beneficial to osseointegration. The
authors also believed that the decreased P. gingivalis adhesion on seen on the alumina
coated titanium was due to the inhibition of calcium ion adsorption. The positively
charged alumina coated on the surface did not attract the similarly charged calcium ions.
The above mentioned studies show the potentially hazardous promotion of
bacterial adhesion with the presence of calcium phosphate. In the present study, P. micra
appears to preferentially adhere to Nanotite implants; however, more research is required
to confirm this result.
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Conditions of P. micra survival and proliferation
Even though P. micra is classified as an anaerobic gram positive bacteria, it is
known to be aero-tolerant with unknown extent. From the results from this experiment, P.
micra was found to be capable of surviving and adhering to implant surfaces within 4 to
24 hours in an atmospheric environment. An increase in adhesion percentage and
possible proliferation of P. micra were noted in all three subgroups of group B. The
appearance of P. micra proliferating as noted by the fused appearance of bacteria was
observed in some samples after 24 hours incubation time (Figure 39).
It was generally believed that exposure to oxygen or room air during surgery
results in decreased viability of anaerobic bacteria through a change in oxidation-
reduction potential of infected tissue (the sunshine theory). Therefore the use of
antibiotic was discouraged unless positive culture with specific anaerobic organisms was
found (Stone, 1975). However, aerotolerance was observed in many species of bacteria
such as Bacteriodes, Fusobacterium, Clostrodium, and Peptostreptococcus (Loesch,
1969). Hardin et al. (1982) found all anaerobic bacteria isolated from induced rat
abdomen abscess and human intra-abdominal sepsis survived after 24 hours exposure to
the room air and the decrease of anaerobic bacteria occurs in parallel with the decrease in
number of aerobic bacteria. They attributed the decrease in both bacteria to the wash-out
mechanism of the surgery. In addition, they stressed the need to administer antimicrobial
therapy to patients with potential anaerobic or mixed bacterial infections.
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This aero-tolerance observed in bacteria, specifically P. micra implies that while
treating peri-implantitis, the simple surgical procedure of exposing the bacteria to the
atmospheric environment may not be sufficient to eliminate the bacteria. Other means of
treatment may be required.

Effect of anaerobic conditions and nutrient broth on P. micra
The hypothesis that nutrient broth and anaerobic conditions promotes growth and
proliferation of P. micra cannot be proven in this study. The nutrient broth contains
thiotone E pepton which produced by digestion of animal tissue by enzymes. It contains
a variety of peptide sizes including ones with large molecular weights which can support
fastidious organisms. It also provides amino acids, vitamins, as well as nitrogen
(http://www.bd.com/ds/technicalCenter/inserts/Thiotone_E_Peptone.pdf). Tryptose,
which provides a large amount of nitrogen, is also a protein produced by enzymatic
breakdown. These broths and culture mediums provide the essential nutrient for the
growth of P. micra in anaerobic environments. (http://www.condalab.com/pdf/1614.pdf).
Nevertheless, incubation with nutrient broth as in group C did not increase the
amount of P. micra adhesion. This is contrary to the study by Almaguer-Flores and
colleagues (2009). In their study, amorphous carbon film and titanium films were tested
for bacterial adhesion under incubation with culture media and saliva. P. micra was
found to increase its adhesion when culture media was used. Authors suspected the
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nutrients and proteins found in culture media promoted the growth while saliva contained
lysozyme, lactoferrin, and secretory IgA which may impair the adhesion.
An additional explanation offered was that large amounts of bacterial adhesion
and replication may had occurred and created a large colony of P. micra. However, due
to the weight of this biofilm, the entire biofilm may have detached from the implant while
rinsing prior to SEM processing. Another explanation may be due to the small sample
size of an in vitro experiment which does not reflect the actual nature of P. micra
adhesion in anaerobic environments provided with nutrients.

P. micra adhesion mechanism and P. micra morphology
According to the SEM images provided in this current study, P. micra appears
spherical. None of the images demonstrated signs of fibril like structures. One can
concluded that the P. micra morphotype used in this experiment was the smooth type.
Therefore, the mechanism of adhesion was most likely based on extracellular
polysaccharides on the cell as illustrated by Kremer et al. 1999.
Different subtypes of bacteria may behave differently under similar condition.
Ökte et al. (1999) showed that different strains of same bacteria may possess different
adherence affinity to the same surface. In their study, A. actinomycetemcomitans was
used and serotype a demonstrated the highest adherence rate. It is likely that variations in
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P. micra subtypes can influence the adherence property. Therefore different
morphotypes of P. micra will need to be tested.

Effect of Serum
Treatment with serum in this current experiment was done to simulate the
presence of gingival crevicular fluid in the oral cavity. As hypothesized, serum
treatments were shown in this study to alter the adhesion pattern of P. micra as observed
in the difference between groups A1 to A2, B2 to B3 and C2 to C3 (Table 5).
Nevertheless, a consistent effect of serum was not observed throughout this experiment.
Serum treatments were effective in inhibiting P. micra adhesion in group A and C, while
increasing adhesion and proliferation in group B.
Contradictory results are found in much of the literature regarding the effect of
serum/crevicular fluid on the bacterial adhesion. In Lima’s study in 2008, they did not
find any beneficial effect of serum coating on bacterial adhesion on titanium. The
titanium non-coated group had similar adhesion of Streptococcus mutans when compared
with the serum coated group. As for Actinomyces naslundii adhesion, a slight increase in
adhesion was observed in the serum coated group (1.43x10
4
cells in non coated group and
1.85x10
4
cells in serum coated group). In their analysis of pellicles on titanium, titanium
with serum coating presented with IgA, IgG, fibronectin, fibrinogen, and albumin.
Amylase, cystatin S, and cystatin SN were not found. Therefore Lima et al. postulated
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that this difference could be the result of a specific serum protein. From Lima’s study, it
appeared that serum’s effect may be bacteria dependent, and specific serum proteins
interacting with bacteria may be an important factor.
In addition, Lima et al. found that in the salivary coated titanium, no albumin was
detected. Salivary coated titanium had IgA, IgG, fibronectin, fibrinogen and amylase
while serum coated titanium had IgA, IgG, fibronectin, fibrinogen, albumin. Discrepancy
in protein content between saliva and serum may also influence bacterial adhesion.
Cimasoni and McBride (1987) tested adherence of Treponema denticola on
modified hydroxyapatite. Hydroxyapatite beads were coated with saliva, gingival
crevicular fluid and serum. All three coatings were not effective in preventing the
adhesion of T. denticola. Only the coating of lysosomal enzymes prepared from human
polymorphonuclear leucocytes (PMN) was effective in consistently preventing the
adhesion of T. dentcola. They attributed this to the non-selective covering of binding
sites on HA surfaces by enzymes and nucleic acids. They theorized that these enzymes
could hydrolyze and eventually inactivate the binding sites on the bacterial surface or on
the HA coated surfaces.
Kohavi et al. in 1995 conducted a study on the adsorption of salivary proteins
onto the titanium components of oral implants. They found amylase as well as albumin
adheres on the titanium surfaces. They postulated that the adsorption of proteins enables
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attachment of some specific bacteria and may later changes the composition of the plaque
buildup.
In certain studies, the effect of serum and protein appeared to be dependent upon
the source of bacteria used in the study. Steinberg et al. (1993) reported that glucans
formed on the acquired pellicle served as a binding site for adhesion of A. viscosus from
rodents while reducing the adhesion of A. viscosus from humans.
In a review written by Hannig and Hannig (2009), based on both in vivo and in
situ studies, they concluded that protein in serum “masks the physicochemical surface
properties of dental materials to a certain extent; however, bacterial adhesion and tenacity
of the bacterial biofilm is considerably influenced by long-range forces transferred
through the pellicle layer.” Thus, serum/proteins may have certain effects on the
bacterial adhesion. The underlying texture of the surface also appeared to be influential
as well as the type of microorganism.
Therefore, the effect of serum or GCF fluid on bacterial adhesion may not be as
straight forward as one would expect as conflicting results were frequently found. The
composition of the fluid, the underlying surface and the characteristics of the bacteria
tested all can be influential in the adhesion of bacteria on the titanium surfaces. Advance
analysis of the bovine serum used in this study may be necessary to explain the results of
this current study. In addition, for future reference, human PICF/GCF or saliva samples
could be used to reflect the actual human oral environment.
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An additional reason for the inconsistency of serum treatment observed in this
current research was the time of exposure to serum. In this study, implants were exposed
to serum for one minute; protein in the serum may not have had enough time to bind to
the titanium surface. In the previous mentioned study by Cimasoni and McBride (1987),
HA beads were placed into coating material for two hours prior to immersion in bacterial
suspension.
As for the effect of serum on P. micra, Cowan’s study (1992) found high protein
content in P. micra. Due to lack of appendage structures, they suspected P. micra
absorbed these proteins extrinsically during its contact with serum protein. This
absorbance of protein content alters the hydrophobicity of P. micra, which may in turn
change the adherence pattern of P. micra. Nevertheless, P. micra was not saturated in
serum in the present study and the possibility that P. micra absorbed protein from serum
treated titanium implant is unlikely.
In reality, serum immersion of implant prior to bacterial suspension does not
replicate the constant washing of GCF or PICF in an in vivo situation and thus the actual
effect of the serum may have been diluted or washed off during the experimental
incubation period. Therefore, the effect of P. micra in serum will need to be investigated
as well.

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Effect of saline rinsing
In this experiment, saline rinsing showed ability to reduce P. micra adhesion
except for Nanotite in group B. The present study utilized a simple monojet curved
syringe to remove P. micra from a one inch distance with hand pressure. The results
from this study suggest that a homecare water jet cleaning device or simple monojet
curved tip syringe used in this study may be potentially useful in removing bacteria on
implant surfaces. Water jet cleaning devices have been advocated for their effective use
on in vitro biofilm removal (Gorur et al., 2009). A 99% reduction in biofilm was found
after application of water jet. However, similar to the present study, Gorur et al.
performed their study in a laboratory setting; many parameters in the oral cavity were not
replicated.
In addition, in an in vitro setting of experiment, direct access and manipulation of
the water jet tip directly towards the tested surfaces is possible. While in an actual
patient’s mouth, direct access to the implant site is often limited and thus the efficacy of
water rinsing may be reduced. This problem may be complicated even more by the
position of the implant in relation to the soft tissue. Irrigation subgingivally has proven
to be difficult. The penetration of solution by irrigation only reaches up to half the
pocket depth (Eakle et al., 1986). One can hypothesize that if the implant is exposed
supragingivally, irrigation of water may aid in removal of bacteria; however, more
research will be needed to test the efficacy of subgingival irrigation of implant surfaces.
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The American Academy of Periodontology position paper (2005) argued against
the solo usage of water irrigation supragingivally without toothbrushing. Many studies
have reflected the benefit on the combination of brushing and irrigation of water. Chaves
et al. (1994) found lower gingival index and mean bleeding on probing values in patients
utilizing brushing with irrigation with water by using WaterPik. Barnes et al. (2005) also
showed the benefit of brushing and water jet use. The sole effect of water jet cleaning
device is questionable. In addition, more studies on the effect of water rinsing on
different implant surfaces are warranted. As for rough implant surfaces such as Nanotite
implant, adjunctive therapy may be needed to improve the ability to remove bacteria.
Machined implants were more readily cleansed free of P. micra during saline
rinsing in the limited samples provided in this current study. The ease of cleansing noted
in smooth surfaced implants was in agreement with Dennison’s study (1994). Dennison
used different chemicals including water, citric acid and CHX soaked in gauze to burnish
the implant surface. They found all methods, including gauze soaked with water,
facilitated the removal of P. gingivalis endotoxin on smooth surfaces. This ease of
cleansing in smooth surfaces determined by mean of microbial and SEM analysis was
also noted in an in vivo study by Baldi et al. (2009).
Nanotite implants with bacterial adhesion appeared to have more resistance to
saline irrigation. This may be explained by surface irregularities which are proven to
protect the bacteria against removal forces (Teughel et al.,2006). It was also reported that
surface roughness becomes important during the presence of active larger shear forces
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(Hannig 1999). One suspects that in order to remove bacteria from rough surfaces such
as Nanotite implants, adjunctive therapy may be needed in addition to water rinsing.
Further studies will help answer this question.

Limitation of the study:
Several limitations exist in the present study. First of all, this experiment was an
in vitro study. The experiment failed to simulate every aspect of the oral cavity including
the presence of saliva, GCF/PICF and their washing effects. PICF was simulated by the
soaking of bovine serum prior to implant treatment, however, constant washing and
flushing of the PICF was not simulated in this study.
Second, periodontal pockets harbor multiple species of bacteria. In this
experiment, only P. micra was used. The interaction between different bacteria to create
the protective effects seen in oral biofilm is a well known phenomenon. This in vitro
experimental condition is unable to simulate a normal human microbial environment and
a single bacterium does not recreate the protective effect found in biofilm. In addition,
bacteria may behave differently in vitro versus in vivo.
Third, P. micra observed on the SEM images provided in this experiment may not
have been viable at the time the samples were fixed. In order to overcome this problem,
Yoshinari et al. (2000) radio-labeled P. gingivalis and A. actinomycetemcomitans by
growing them in a broth with [
3
H] thymidine or [
3
H] uridine. Bacteria were then counted
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by direct scintillation counting after processing on the desired surfaces. Morphology of
adherence was observed through SEM. Harris and Richards (2004) used a similar
technique in their experiment. S. aureus were stained with fluorescent material for
visualization under fluorescence microscope and computer software was used to count
the density of living bacteria adhering to the surface. Amoroso et al. (2006) utilized
acridine orange to stain P. gingivalis, observed them under fluorescent microscopy and
quantified their adhesion through a computer program.
These labeling and staining methods increase the likelihood that the bacteria
observed on the surface are alive and active. The use of computer programs may further
decrease human error during counting.
Wu Yuan et al. (1995) utilized a method similar to the present study by counting
the bacteria on SEM. According to their method, the bacteria was counted in each field
which was calculated to be 2.817x10
3
µm
2
and the mean number of bacteria in five fields
was determined and calculated as cell numbers per mm
2
. This method is comparably
similar to the method utilized in the present experiment. The process of randomization in
selecting SEM images in such studies is extremely important to have an accurate and
non-biased count of bacteria.
Different methods to quantify the amount of bacteria present on a given surface
are also available. One way is to swab the surface of biofilm, then plate and count the
colonizing units of bacteria (Drake et al. 1999). Another way is to “vortex” the bacteria
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off the tested surfaces and culture the bacteria to count the colony-forming units (Pereira
da Silva et al. 2005).
When a biofilm environment is created, different bacteria can be identified and
quantified. Almaguer-Flores and co-workers (2009) included 9 different kinds of
bacteria and tested the bacterial adhesion on different surfaces. In their experiment, a
bacterial adhesion test was conducted after 7 days of anaerobic incubation, colony
forming units (CFU) were visually counted to calculate the initial number of bacteria
attached. In order to determine the proportion of each bacterial strain in the surface,
checkerboard DNA-DNA hybridization was utilized. This method can be used in future
study when a biofilm is simulated.
Another important difference between the present study and other in vitro studies
was the usage of actual implants in this study. Full unaltered manufacturer implants were
used while other researchers used small pieces or discs of titanium. The entire surface
can be counted and processed easily when small pieces of titanium is used. However, the
3D cylindrical shaped titanium implants used here can only be examined on one side at a
time, increasing the difficulty of obtaining bacterial counts.
In addition, more quantitative information such as surface free energy of the
implants needs to be obtained in future experiments to make direct comparisons with
other studies. In addition, the roughness values on some implants are lacking (i.e.
110

Implantium implants) further complicating comparison. More information on P. micra’
ability to adhere to hard surfaces may help with the analysis of this study as well.
Finally, the major drawback of this pilot experiment is the small number of
implant samples included in each group. The limited number of implants tested results in
a wide range of standard deviations as well as insignificance in the statistical analysis
performed. Future research will require larger sample sizes, improved methodology and a
closer mimicking of the oral environment.
111

Chapter 5: Conclusion
In conclusion, P. micra was capable of adhering to all the implant surfaces tested
in this study. Interestingly, it appeared that any treatment modification to the implant
surface, smooth or rough, was capable of affecting P. micra adhesion to some degree.
Therefore, based on the findings of this study, a recommendation for thorough research
on newly developed surfaced implants to ensure safety prior to clinical patient usage
seems justified. In addition, in spite of lack of statistical significance in this pilot study,
the potential of saline rinsing as a method of bacterial removal stands out as having
important implications for further research into simple and economical treatments.
Finally, due to the in vitro nature and small number of samples dictated by available
resources, further research will be required to clarify results of the present pilot study.

112

References
Absolom, DR., Lamberti FV., Policova Z., Zingg W., van Oss., CJ, and Neumann AW.,
(1983). “Surface thermodynamics of bacterial adhesion.” Appl Environ Microbiol 46:
90-97.
Albrektsson, T., Isidor, F. (1994). “Consensus report: implant therapy.” Proceedings of
the 1
st
European Workshop on Periodontology Berlin: Quintessence. 365–369.

Albrektsson T., Wennerberg A., (2004). “Oral implant surfaces: Part 1–review focusing
on topographic and chemical properties of different surfaces and in vivo responses to
them.” Int J Prosthodont 17: 536–543.
Alcoforado GA., Rams TE., Feik D., Slots J., (1991). “Microbial aspects of failing
osseointegrated dental implants in humans.” J Parodontol 10(1): 11-8.
Almaguer-Flores A., Ximénez-Fyvie LA., Rodil SE., “Oral bacterial adhesion on
amorphous carbon and titanium films: effect of surface roughness and culture media.” J
Biomed Mater Res B Appl Biomater 92(1):196-204.
American Academy of Periodontology: Position paper, (2005). “The role of supra- and
subgingival irrigation in the treatment of periodontal diseases.” J Periodontol 76: 2015-
2027.
Amoroso PF., Adams RJ., Waters MG., Williams DW., (2006). “Titanium surface
modification and its effect on the adherence of Porphyromonas gingivalis: an in vitro
study.” Clin Oral Implants Res. 17(6):633-637.
Augthun M., Tinschert J., Huber A., (1998). “in vitro studies on the effect of cleaning
methods on different implant surfaces.” J Periodontol 69(8): 857-64.
Baldi D., Menini M., Pera F., Ravera G., Pera P., (2009). “Plaque accumulation on
exposed titanium surfaces and peri-implant tissue behavior. A preliminary 1-year
clinical study.” Int J Prosthodont 22(5):447-55.
Bang JS., Cimasoni G., (1971). “Total protein in human crevicular fluid.” J Dent Res
50(6): 1683.
Barnes CM., Russell CM., Reinhardt RA., Payne JB., Lyle DM., (2005). “Comparison
of irrigation to floss as an adjunct to tooth brushing: effect on bleeding, gingivitis, and
supragingival plaque.” J Clin Dent 16(3): 71-77.
113

Bida DF., (1991). “Current applications of the dental laser in implantology.” Implant
Soc 2: 8–10.
Block CM, Mayo JA, Evans GH. Effects of the Nd:YAG dental laser on plasma-sprayed
and hydroxyapatite-coated titanium dental implants: Surface alteration and attempted
sterilization. Int J Oral Maxillofac Implants 1992; 7: 441–449.
Bollen CML., Papaioannou W., VanEldere J., Schepers E., Quirynen M., Van
Steenberghe D., (1996). “The influence of abutment surface roughness on plaque
accumulation and peri-implant mucositis.” Clin Oral Impl Res 7: 201-211.

Brånemark PI., Hansson BO., Adell R, Breine U, Lindström J, Hallén O, Ohman A.
Osseointegrated implants in the treatment of the edentulous jaw. Experience from a 10-
year period. Scand J Plast Reconstr Surg Suppl. 1977;16:1-132.

Burgers R., Gerlach T., Hahnel S., Schwarz F., Handel G., Gosau M., (2010). “in vivo
and in vitro biofilm formation on two different titanium implant surfaces.” Clin Oral
Impl Res 21: 156-164.

Carlsson J., Larsen JT., Edlund MB., (1993). “Peptostreptococcus micros has a
uniquely high capacity to form hydrogen sulfide from glutathione.” Oral Microbiol
Immunol 8(1): 42-45.
Chairay JP., Boulekbache H., Jean A., Soyer A., Bouchard P., (1997). “Scanning
electron microscopic evaluation of the effects of an air-abrasive system on dental
implants: a comparative in vitro study between machined and plasma-sprayed titanium
surfaces.” J Periodontol 8(12): 1215-1222.
Chaves ES., Kornman KS., Manwell MA., Jones AA., Newbold DA.,Wood RC., (1994).
“Mechanism of irrigation effects on gingivitis.” J Periodontol 65: 1016-1021.
Cimasoni G., McBride BC., (1987). “Adherence of Treponema denticola to Modified
Hydrozyapatite.” J Dent Res 66(12): 1727-1729.
Coelho PG., Granjeiro JM., Romanos GE., Suzuki M,Silva NRF., Cardaropoli G.,
Thompson VP., Lemons JE., (2009a). “Review: basic research methods and current
trends of dental implant surfaces.” J Biomed Mater Res Part B: Appl Biomater 88:
579–596.

Coelho PG., Cardarpoli G., Suzuki M., and Lemons JE., (2009b). “Early healing of
nanothickness bioceramic coatings on dental implants. An experimental study in dogs.”
J Biomed Mater Res Part B: Appl Biomater 88: 387-393.
114

Cooper LF., (2000). “A role for surface topography in creating and maintaining bone at
titanium endosseous implants.” J Prosthet Dent 84(5): 522-534.
Cowan MM., van der Mei HC., Rouxhet PG., Busscher HJ., (1992). “Physico-chemical
and structural properties of the surfaces of Peptostreptococcus micros and Streptococcus
mitis as compared to those of mutans streptococci, Streptococcus sanguis and
Streptococcus salivarius.” J Gen Microbiol 138(12):2707-2714.
Curtis DA., Kao R., Plesh O., Finzen F., Franz L., (1997). “Crevicular fluid analysis
around two failing dental implants: A clinical report.” J Prosthod 6: 210-214.
Curtis MA., Griffiths GS., Price SJ., Coulthurst SK., Johnson NW., (1988). “The total
protein concentration of gingival crevicular fluid. Variation with sampling time and
gingival inflammation.” J clin Periodontol 15: 628-632.
Davies JE., (2003). “Understanding peri-implant endosseous healing.” J Dent Educ
67(8): 932-949.
De Groot D., Wolke JGA., Jensen JA., (1998). “Calciums phosphate coatings for
medical implants.” Preceedings of Institution of Mechnical Engineers. Journal of
Engineering in Medicine 212(H): 137-147.
Dennison DK., Huerzeler MB., Quinones C., Caffesse RG., (1994). “Contaminated
implant surfaces: an in vitro comparison of implant surface coating and treatment
modalities for decontamination.” J Periodontol 65(10): 942-948.
Deppe H., Horch HH., Henke J., Donath K., (2001). “Peri-implant care of ailing
implants with the carbon dioxide laser.” Int J Oral Maxillofac Implants 16(5): 659-667.
Drake DR., Paul J., Keller JC., (1999). “Primary bacterial colonization of implant
surfaces.” Int J Oral Maxillofac Implants 14(2): 226-232.
Duarte PM., Reis AF., de Freitas PM., Ota-Tsuzuki C., (2009). “Bacterial adhesion on
smooth and rough titanium surfaces after treatment with different instruments.” J
Periodontol 80(11): 1824-1832.
Eakle WS., Ford C., Boyd RL., (1986). “Depth of penetration in periodontal pockets
with oral irrigation.” J Clin Periodontol 13(1): 39-44.
Esposito M., Hirsch JM., Lekholm U., Thomsen P., (1997). “Failure patterns of four
osseointegrated oral implant systems.” J Mater Sci Mater Med 8(12):843-847.

115

Ezaki T., Yabuuchi E., (1985). “Oligopeptidase activity of gram-positive anaerobic
cocci used for rapid identification.” J Gen Appl Microbiol 31: 255–266.
Glantz PO., (1970). “The surface tension of saliva.” Odontologisk Revy 21: 119-127.
Goené RJ., Testori T., Trisi P., (2007). “Influence of a nanometer-scale surface
enhancement on de novo bone formation on titanium implants: a histomorphic study in
human maxilla.” Int J Periodontics Restorative Dent 27: 211-219.
Gopal J., George RP., Muraleedharan P., Khatak HS., (2004). “Photocatalytic inhibition
of microbial adhesion by anodized titanium.” Biofouling 20(3): 167-175.
Gorur A., Lyle DM., Schaudinn C., Costerton JW., (2009). “Biofilm removal with a
dental water jet.” Compend Contin Ed Dent 30(Suppl 1): 1-6.
Hämmerle CH., Fourmousis I., Winkler JR., Weigel C., Brägger U., Lang NP., (1995).
“Successful bone fill in late peri-implant defects using guided tissue regeneration. A
short communication.” J Periodontol 66(4): 303-308.
Hannig C., Hannig M., (2009). “The oral cavity – A key system to understand
substratum-dependent bioadhesion on solid surfaces in men.” Clin Oral Investig 13:
123-139.
Hannig M., (1999). “Transmission electron microscopy of early plaque formation on
dental materials in vivo.” European Journal of Oral Science 107: 55-64.

Hardin WD., Aran AJ., Smith JW., Nichols RL., (1982). “Aerotolerance of common
anaerobic bacteria – fact or fancy?” Southern Medical Journal 75(9): 1051-1056.

Harris LG., Richards RG., (2004). “Staphylococcus aureus adhesion to different treated
titanium surfaces.” J Mater Sci Mater Med 15: 311-314.
Hultin M., Gustafsson A., Hallstrom H., Johansson LA., Ekfeldt A., Klinge B., (2002).
“Microbiological findings and host response in patients with peri-implantitis.” Clin
Oral Impl Res 13: 349-358.
Jarmar T., Palmquist A., Brånemark R., Hermansson L., Engqvist H., Thomsen P.,
(2008). “Characterization of the surface properties of commercially available dental
implants using scanning electron microscopy, focused ion beam, and high-resolution
transmission electron microscopy.” Clin Implant Dent Relat Res 10(1): 11-22.
Jovanovic SA., Kenney EB., Carranza FA Jr., Donath K., (1993). “The regenerative
potential of plaque-induced peri-implant bone defects treated by a submerged membrane
technique: an experimental study.” Int J Oral Maxillofac Implants 8(1):13-8.
116

Kim H., Choi SH., Ryu JJ., Koh SY., Park JH., and Lee IS., (2008). “The
biocompatibility of SLA-treated titanium implants.” Biomed Mater 3: 1-6.
Kohavi D., Klinger A., Steinberg D., Sela MN., (1995). “Adsorption of salivary
proteins onto prosthetic titanium components.” J Prosthet Dent 74(5): 531-534.
Kremer BH., Loos BG., van der Velden U., van Winkelhoff AJ., Craandijk J., Bulthuis
HM., Hutter J., Varoufaki AS., van Steenbergen TJ., (2000). “Peptostreptococcus
micros smooth and rough genotypes in periodontitis and gingivitis.” J Periodontol
71(2):209-218.
Kremer BHA., Magee JT., van Dalen PJ., van Steenbergen TJM., (1997).
Characterization of smooth and rough morphotypes of Peptostreptococcus micros.” Int
J Syst Bacteriol 47: 363-368.
Kremer BH., Herscheid AJ., Papaioannou W., Quirynen M., van Steenbergen TJ.,
(1999). “Adherence of Peptostreptococcus micros morphotypes to epithelial cells in
vitro.” Oral Microbiol Immunol 14(1):49-55.
Kremer BH., van Steenbergen TJ., (2000). “Peptostreptococcus micros coaggregates
with Fusobacterium nucleatum and non-encapsulated Porphyromonas gingivalis.”
FEMS Microbiol Lett 182(1):57-62.
Kuula H., Könönen E., Lounatmaa K., Konttinen YT., Könönen M., (2004).
“Attachment of oral gram-negative anaerobic rods to a smooth titanium surface: an
electron microscopy study.” Int J Oral Maxillofac Implants 19(6): 803-809.
Lamster IB., Ahlo JK., (2007). “Analysis of gingival crevicular fluid as applied to the
diagnosis of oral and systemic diseases.” Ann N Y Acad Sci 1098: 216-229.
Le Guéhennec L., Soueidan A., Layrolle P., Amouriq Y., (2007). “Surface treatments of
titanium dental implants for rapid osseointegration.” Dent Mater 23(7): 844-854.
Lima EM., Koo H., Vacca Smith AM., Rosalen PL., Del Bel Cury AA., (2008).
“Adsorption of salivary and serum proteins, and bacterial adherence on titanium and
zirconia ceramic surfaces.” Clin Oral Implants Res 19(8):780-785.
Listgarten MA., Lai CH., (1999). “Comparative microbiological characteristics of
failing implants and periodontally diseased teeth.” J Periodontol 70(4): 431-437.
Loesche WJ., (1969). “Oxygen sensitivity of various anaerobic bacteria.” Appl
Microbiol 11: 723-727.
117

Lucchini JP., Aurelle JL., Therin M., Donath K., Becker W., (1996). “A pilot study
comparing screw-shaped implants. Surface analysis and histologic evaluation of bone
healing.” Clin Oral Implants Res 7: 397-404.

Muraleedharan P., Gopal J., George RP., Khatak HS., (2003). “Photocatalytic
bactericidal properties of an anodized Ti6Al4V alloy.” Curr Sci 84(2): 197-199.

Mendes VC., Moineddin R., Davies JE., (2007). “The effect of discrete calcium
phosphate nanocrystals on bone-bonding to titanium surfaces.” Biomaterials 28: 4748-
4755
Mendes VC., Moineddin R., Davies JE., (2009). “Discrete calcium phosphate
nanocrystalline deposition enhances osteoconduction on titanium-based implant
surfaces.” J Biomed Mater Res A 90(2): 577-585.
Mengel R., Buns CE., Mengel C., Flores-de-Jacoby L., (1998). “An in vitro study of the
treatment of implant surfaces with different instruments.” Int J Oral Maxillofac
Implants 13(1):91-96.
Mombelli A., Van Oosten MAC., Schurch E., Lang NP., (1987). “The microbiota
associated with successful or failing osseointegrated titanium implants.” Oral Microbiol
Immunol 2: 145-151.
Moore LV., Johnson JL., Moore WEC., (1986). “Genus Peptostreptococcus.” Bergey’s
manual of systematic bacteriology The Williams & Wilkins Co., Baltimore, Md. 2:
1083–1092
Murdoch DA., Mitchelmore IJ., (1991). “The laboratory identification of gram-positive
anaerobic cocci.” J Med Microbiol 34: 295–308.

Murdoch DA., Mitchelmore IJ., Tabaqchali S., (1988). “Peptostreptococcus micros in
polymicrobial abscesses.” Lancet I: 594.
Niimi A., Ueda M., (1995). “Crevicular fluid in the osseointegrated implant sulcus: A
pilot study.” Int J Oral Maxillofac Implants 4: 434-436.
Ofek I., Hasty DL., Sharon N., (2003). “Anti-adhesion therapy of bacterial diseases:
prospects and problems.” FEMS Immunol Med Microbiol 38(3):181-191.
118

Ökte E., Sultan N., Başak D., Asikainen S., (1999). “Bacterial adhesion of
Actinobacillus actinomycetemcomitans serotypes to titanium implants: SEM evaluation.
A preliminary report.” J Periodontol 70: 1376–1382.
Orsini G., Piattelli M., Scarano A., Petrone G., Kenealy J., Piattelli A., and Caputi S.,
(2007). “Randomized, controlled histologic and histomorphometric evaluation of
implants with nanometer-scale calcium phosphate added to the dual acid-etched surface
in the human posterior maxilla.” J Periodontol 78: 209-218.
Ota-Tsuzuki C., Alves Mayer MP., (2010). “Collagenase production and hemolytic
activity related to 16S rRNA variability among Parvimonas micra oral isolates.”
Anaerobes 16: 38-42.

Park YS., Yi KY., Lee IS., Han CH., Jung YC., (2005). “The effect of ion beam-
assisted deposition of hydroxyapatite on the grit blasted surface of endosseous implants
in rabbit tibiae.” Int J Oral Maxillofac Implants 20: 31-38.

Pereira da Silva CH., Vidigal GM., de Uzeda M., de Almeida Soares G., (2005).
“Influence of titanium surface roughness on attachment of Streptococcus sanguis: an in
vitro study.” Implant Dent 14(1): 88-93.
Persson LG., Araújo MG., Berglundh T., Gröndahl K., Lindhe J., (1999). “Resolution
of peri-implantitis following treatment. An experimental study in the dog.” Clin Oral
Implants Res 10(3): 195-203.
Persson S., Edlund MB., Claesson R., Carlsson J., (1990). “The formation of hydrogen
sulfide and methyl mercaptan by oral bacteria.” Oral Microbiol Immunol 5(4): 195-201.
Persson LG., Berglundh T., Sennerby L., Lindhe J., (2001). “Reosseointegration after
treatment of peri-implantitis at different implant surfaces. An experimental study in the
dog.” Clin Oral Impl Res 12: 595–603.

Puckett SD., Taylor E., Raimondo T., Webster TJ., (2010). “The relationship between
the nanostructure of titanium surfaces and bacterial attachment.” Biomaterials 31: 706-
713.
Quirynen M., Marechal M., Busscher HJ., Weerkamp AH., Darius PL., van Steenberghe
D., (1990). “The influence of surface free energy and surface roughness on early plaque
formation. An in vivo study in man.” J Clin Periodontol 17: 138-144.

119

Quirynen M., Bollen CML., (1995). “The Influence of Surface Roughness and Surface-
free energy on Supra- and Subgingival Plaque Formation in Man – A review of the
literature.” J Clin Periodontol 22: 1-14.

Quirynen M., van der Mei HC., Bollen CM., Schotte A., Marechal M., Doornbusch GI.,
Naert I, Busscher HJ., van Steenberghe D., (1993). “An in vivo study of the influence of
the surface roughness of implants on the microbiology of supra- and subgingival
plaque.” J Dent Res 72(9):1304-9.
Quirynen M., Bollen CM., Papaioannou W., Van Eldere J., van Steenberghe D., (1996).
“The influence of titanium abutment surface roughness on plaque accumulation and
gingivitis: short-term observations.” Int J Oral Maxillofac Implants 11(2): 169-178.

Quirynen M., De Soete M., van Steenberghe D., (2002). “Infectious risks for oral
implants: a review of the literature.” Clin Oral Impl Res 13: 1–19.
Rams TE., Roberts TW., Feik D., Molzan AK., Slots J., (1991). “Clinical and
microbiological findings on newly inserted hydroxyapatite-coated and pure titanium
human dental implants.” Clin Oral Impl Res 2: 121-127.
Rimondini L., Farè S., Brambilla E., Felloni A., Consonni C., Brossa F., Carrassi A.,
(1997). “The effect of surface roughness on early in vivo plaque colonization on
titanium.” J Periodontol 68(6): 556-562.
Sela MN., Badihi L., Rosen G., Steinberg D., Kohavi D., (2007). “Adsorption of human
plasma proteins to modified titanium surfaces.” Clin Oral Implants Res 18(5): 630-638.
Salcetti JM., Moriarty JD., Cooper LF., Smith FW., Collins JG., Socransky SS.,
Offenbacher S., (1997). “The clinical, microbial, and host response characteristics of
the failing implant.” Int J Oral Maxillofac Implants 12(1): 32-42.
Schou S., Holmstrup P., Jorgensen T., (2003). “Implant surface preparation in the
surgical treatment of experimental peri-implantitis with autogenous bone graft and
ePTFE membrane in cynomolgus monkeys.” Clin Oral Impl Res 14: 412-422.
Song Y., Finegold SM., (2007). “Anaerobic bacteria – Peptostreptococcus, Finegoldia,
Anaerococcus, Peptoniphilus, Veillonella, and other anaerobic cocci.” Manual of
Clinical Microbiology 9
th
Edition. ASM Press, Washington, DC. 862-888
Steinberg D., Kopec LK., Bowen WH., (1993). “Adhesion of actinomyces isolates to
experimental pellicles.” J Dent Res 72(6):1015-1020.

120

Stone HH., Kolb LD., and Geheber CE., (1975). “Incidence and significance of
intraperitoneal anaerobic bacteria.” Ann Surg 5: 705-714.

Stout KJ., Davis EJ., Sullivan PJ., (1990). Atlas of Machined Surfaces. Chapman and
Hall: London.
Stübinger S., Etter C., Miskiewicz M., Homann F., Saldamli B., Wieland M., Sader R.,
(2010). “Surface alterations of polished and sandblasted and acid-etched titanium
implants after Er:YAG, Carbon Dioxide and diode laser irradiation.” Int J Oral
Maxillofac Implants 25(1): 104-111.
Sul YT., Johansson CB., Jeong Y., Wennerberg A., Albrektsson T., (2002). “Resonance
frequency and removal torque analysis of implants with turned and anodized surface
oxide.” Clin Oral Impl Res 13: 252-259.
Sul YT., Johansson C., Albrektsson T., (2006). “Which Surface properties enhance
bone response to implants? Comparison of oxidized magnesium, TiUnite, and Osseotite
implant surfaces.” Int J Prosthdont 19: 319-328.
Tabanella G., Nowzari H., Slots J., (2009). “Clinical and microbiological determinants
of ailing dental implants.” Clin Implant Dent Relat Res 11(1):24-36.
Tam YC., Chan EC., (1985). “Purification and characterization of hyaluronidase from
oral Peptostreptococcus species.” Infect Immun 47(2): 508-513.
Tam YC., Noble PB., Chan EC., (1987). “Effects of extracellular products of a
presumed gingival pathogen, the bacterium Peptostreptococcus 84H14S, on cultured
human fibroblasts and HeLa cells.” Arch Oral Biol 32(4): 303-305.
Tanabe S., Bodet C., Grenier D., (2007). “Peptostreptococcus micros cell wall elicits a
pro-inflammatory response in human macrophages.” J of Endotoxin Res 13: 219-226.
Tanner J., Robinson C., Söderling E., and Vallittu P., (2005). “Early plaque formation
on fiber-reinforced composites in vivo.” Clin Oral Invest 9: 154-160.
Teronen O., Konttinen YT., Lindqvist C., Salo T., Ingman T., Lauhio A., Ding Y.,
Santavirta S., Sorsa T., (1997). “Human neutrophil collagenase MMP-8 in peri-implant
sulcus fluid and its inhibition by clodronate.” J Dent Res 76(9): 1529-1537.
Teughels W., Van Assche N., Sliepen I., Quirynen M., (2006). “Effect of material
characteristics and/or surface topography on biofilm development.” Clin Oral Imp Res
17(Suppl. 2): 68–81.
121

Tsibouklis J., Stone M., Thorpe AA., Graham P., Peters V., Heerlien R., Smith JR.,
Green KL., Nevell TG., (1999). “Preventing bacterial adhesion onto surfaces: the low-
surface-energy approach.” Biomaterials 20(13):1229-1235.
Van Dalen PJ., van Steenbergen TJM., Cowan MM., Busscher HJ., de Graaff J., (1993).
“Description of Two morphotypes of Peptostreptococcus micros.” Int J Syst Bacteriol
43: 787–793.
Waerhaug J., (1956). “Effect of rough surfaces upon gingival tissue.” J Dent Res 35:
323-325.
Weerkamp AH., van der Mei HC., Busscher HJ., (1985). “The surface free energy of
oral streptococci after being coated with saliva and its relation to adhesion in the
mouth.” J Dent Res 64(10):1204-1210.
Wennerberg A., Albrektsson T., (2000). “Suggested guidelines for the topographic
evaluation of implant surfaces.” The Int J Oral Maxillofac Implants 15: 331-344.
Wennerberg A., Sennerby L., Kultje C., Lekholm U., (2003). “Some soft tissue
characteristics at implant abutments with different surface topography. A study in
humans.” J Clin Periodontol 30(1): 88-94.
Wennerberg A., Albrektsson T., (2009). “Effects of Titanium Surface Topography on
bone integration: a systematic review.” Clin Oral Impl Res 20(Suppl 4): 172-184.
Wennerberg A., Albrektsson T., (2010). “On implant surfaces: A review of current
knowledge and opinions.” The Int J Oral Maxillofac Implants 24: 63-74.
Wu-Yuan CD., Eganhouse KJ., Keller JC., Walters KS., (1995). “Oral bacterial
attachment to titanium surfaces: a scanning electron microscopy study.” J Oral
Implantol 21(3):207-213.
Yamashita Y., Kunimori A., Takehara T., (1991). “Effect of calcium ions on cell
surface electrostatics of Bacteriodes gingivalis and other oral bacteria.” Zentralbl
Bakteriol 275: 46-53.

Yoshinari M., Oda Y., Kato T., Okuda K., Hirayama A., (2000). “Influence of surface
modifications to titanium on oral bacterial adhesion in vitro.” J Biomed Mater Res
52(2):388-394.

122

Yoon HJ., Son JE., Um YJ., Chae GJ., Chung SM., Lee IS., Jung UW., Kim CS., Choi
SH., (2009). “Effect of calcium phosphate coating to SLA surface Implants by the ion-
beam-assisted deposition method on self-contained coronal defect healing in dogs.”
Biomedical Materials 4: 1-6.
Zitzman NU., Berglundh T., (2008). “Definition and prevalence of peri-implant
diseases.” J Clin Periodontol 35 (Suppl. 8): 286–291.

Abstract (if available)

Abstract Objectives: Despite the high success rate of implant dentistry, failure of implants due to peri-implantitis occurs and may present difficult treatment decisions. Many bacteria have been associated with peri-implantitis, including P. micra. P. micra is considered to be a putative peri-implantitis bacterium which is often isolated from active peri-implantitis. The aim of this in vitro study is to evaluate the ability of P. micra to adhere to different implant surface textures that are currently on the market. In addition, similar surfaced implants between different manufacturing companies were also compared.

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Asset Metadata

Creator Chiu, Ching Hsiu Ketty (author)

Core Title Parvimonas micra adhesion on different implant surfaces: an in vitro pilot study

School School of Dentistry

Degree Master of Science

Degree Program Craniofacial Biology

Publication Date 06/23/2010

Defense Date 06/09/2010

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag bacterial adhesion,dental implant,implant modifications,OAI-PMH Harvest,Parvimonas micra,titanium surface,water rinsing

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Nowzari, Hessam (committee chair), Navazesh, Mahvash (committee member), Rich, Sandra (committee member)

Creator Email chinghsc@usc.edu,kettychiu1009@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m3150

Unique identifier UC187715

Identifier etd-Chiu-3872 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-351253 (legacy record id),usctheses-m3150 (legacy record id)

Legacy Identifier etd-Chiu-3872.pdf

Dmrecord 351253

Document Type Thesis

Rights Chiu, Ching Hsiu Ketty

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu