© 2016 christie lauren eastmanufdcimages.uflib.ufl.edu/uf/e0/04/99/73/00001/eastman_c.pdf ·...
TRANSCRIPT
DEVELOPMENT OF A NOVEL RAT MODEL OF POLYMICROBIAL PERI-IMPLANTITIS INFECTION
By
CHRISTIE LAUREN EASTMAN
A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
2016
© 2016 Christie Lauren Eastman
To my family who has been my strength and guidance, always encouraging my goals and future endeavors
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ACKNOWLEDGMENTS
I thank my boyfriend, friends and family for their support, unconditional love and
enthusiasm throughout my education. I am obliged to my mentor, Dr. L. Kesavalu, for
becoming my mentor halfway through the project and for allowing me the opportunity to
work under his instrumental guidance. I am also obliged to my original mentor, Dr. T.
Koutouzis, who was instrumental in the creation of this project and who performed all
surgical procedures. I would also like to thank my committee members for their advice
and supervision in the completion of this thesis. Lastly, I would like to express my
deepest appreciation and gratitude to faculty of the Department of Periodontology for
their invaluable guidance and continued advancement of my education and clinical
abilities.
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TABLE OF CONTENTS page
ACKNOWLEDGMENTS .................................................................................................. 4
LIST OF TABLES ............................................................................................................ 7
LIST OF FIGURES .......................................................................................................... 8
LIST OF ABBREVIATIONS ............................................................................................. 9
ABSTRACT ................................................................................................................... 10
CHAPTER
1 INTRODUCTION .................................................................................................... 12
Peri-Implant Health ................................................................................................. 12 Peri-Implant Diagnosis ............................................................................................ 14 Peri-Implant Disease............................................................................................... 14 Animal Studies: Ligature-Induced Defect Model ..................................................... 17 Animal Models: Peri-Mucositis ................................................................................ 18 Animal Models: Peri-Implantitis ............................................................................... 18 Animal Studies: Animal Selection ........................................................................... 19 Study Aim ............................................................................................................... 21 Study Hypothesis .................................................................................................... 21
2 MATERIALS AND METHODS ................................................................................ 23
Rats ........................................................................................................................ 23 Surgical Procedures ............................................................................................... 23 Bacterial Strains and Polymicrobial Inocula ............................................................ 24 Peri-Implantitis Induction with P. gingivalis, T. denticola and T. forsythia ............... 24 Gingival Plaque Sampling ....................................................................................... 25 Detection of Bacterial Genomic DNA in Gingival Plaque Samples ......................... 25 Euthanasia and Tissue Specimen Collection .......................................................... 26 Serum IgG and IgM Antibody Analysis ................................................................... 26 Morphometric Analysis of Mandibular Lingual Alveolar Bone Resorption ............... 26 Microcomputed Tomography (microCT) Peri-Implant Bone Levels ........................ 27 Histological Evaluation of Gingival Inflammation .................................................... 27 Statistical Analysis .................................................................................................. 28
3 RESULTS ............................................................................................................... 34
Clinical Results of Implant and Healing Abutment Placement ................................ 34 Peri-Implantitis Induction with Periodontal Bacteria ................................................ 34 Polymicrobial Infection Elicits Humoral Antibody Response ................................... 34
6
Polymicrobial Infection Elicits Horizontal Mandibular Alveolar Bone Resorption .... 35 MicroCT Imaging Analysis of Peri-Implant Bone ..................................................... 35 Peri-Implant Histology ............................................................................................. 35
4 DISCUSSION ......................................................................................................... 45
LIST OF REFERENCES ............................................................................................... 50
BIOGRAPHICAL SKETCH ............................................................................................ 56
7
LIST OF TABLES
Table page 1-1 Methodological procedure of standard animal ligature-induced defect models ....... 22
3-1 Remaining implants and healing abutments in infected and sham-infected groups according to microCT images ..................................................................... 37
3-2 Distribution of rat gingival microbial samples positive for Pg/Td/Tf by PCR ............ 38
3-3 Mean peri-implant bone levels in infected and sham-infected rats .......................... 39
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LIST OF FIGURES
Figure page 2-1 Experimental design. ............................................................................................... 29
2-2 Surgical protocol. .................................................................................................... 30
2-3 Implant & healing abutment.. .................................................................................. 31
2-4 Morphometric evaluation of horizontal alveolar bone resorption. ............................ 32
2-5 Representative microCT image of an implant-abutment complex.. ......................... 33
3-1 Rat maxillae showing presence or absence of healing abutments at conclusion of polymicrobial infection. ....................................................................................... 40
3-2 Polymicrobial infection induced bacterial specific IgG and IgM antibody response.. ............................................................................................................... 41
3-3 Mandibular lingual alveolar bone resorption.. .......................................................... 42
3-4 Peri-implant microCT alveolar bone resorption ....................................................... 43
3-5 Histological staining of the peri-implant tissues.. ..................................................... 44
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LIST OF ABBREVIATIONS
ABC
ABR
BC
BIC
BOP
CAL
CEJ
CT
GT
IgG
IgM
IP
JE
M1
M2
M3
MicroCT
PD
PDL
PE
Pg
PIE
PMN
Td
Tf
Alveolar bone crest
Alveolar bone resorption
Bone contact
Bone to implant contact
Bleeding on probing
Clinical attachment level
Cementoenamel junction
Connective tissue
Granulation tissue
Immunoglobulin G
Immunoglobulin M
Implant platform
Junctional epithelium
Maxillary 1st molar
Maxillary 2nd molar
Maxillary 3rd molar
Microcomputed tomography
Probing depth
Periodontal ligament
Pocket epithelium
Porphyromonas gingivalis
Peri-implant epithelium
Polymorphonuclear
Treponema denticola
Tannerella forsythia
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Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science
DEVELOPMENT OF A NOVEL RAT MODEL OF POLYMICROBIAL
PERI-IMPLANTITIS INFECTION
By
Christie L. Eastman
May 2016
Chair: Lakshmyya Kesavalu Major: Dental Sciences – Periodontics
Peri-implantitis is a complex polymicrobial-induced inflammatory osteolytic
infection that can lead to implant failure. The aim of this study was to develop a novel
peri-implant rat model by using an established rat model of polymicrobial periodontitis.
Wistar rats were used in this study. One month following bilateral extraction of maxillary
first molars (M1), mini titanium machine-surfaced implants were placed into the
edentulous sites. Two months later, implants were uncovered and abutments were
connected. Rats were divided into two groups: 1) Group A: polymicrobial-infected, n=6;
and 2) Group B: sham-infected, n=6. One week following uncovery, rats were infected
with Porphyromonas gingivalis (Pg), Treponema denticola (Td), and Tannerella
forsythia (Tf) for 12 weeks. Three rats with 4 implants in Group A and 2 rats with 3
implants in Group B were available for analysis. Bacterial genomic DNA was detected
along with infection induced serum IgG and IgM antibodies against all three bacteria,
indicating bacterial colonization around the implants. Polymicrobial infection in the
implanted rats resulted in a greater mean distance from the implant platform (IP) to the
first bone to implant contact (BIC) when compared to the sham-infected, implanted rats
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(0.80 ± .37 mm vs. 0.48 ± 0.13 mm, respectively). Polymicrobial infected peri-implant
tissues showed extensive peri-implantitis with advanced alveolar bone resorption (ABR)
and extensive inflammation with granulation tissue and polymorphonuclear (PMN)
leukocyte infiltration. This is the first study to develop a novel rat model for peri-
implantitis with polymicrobial infection with human pathogens and may provide
important insights into the initiation and progression of peri-implantitis.
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CHAPTER 1 INTRODUCTION
Since osseointegrated implants were introduced by Brånemark et al, extensive
research has been carried out and dental implants are now considered a reliable
treatment option for predictable tooth replacement1. Over the past 30 years, research
has validated the success of osseointegrated implants as an alternative to removable or
tooth-borne prosthetic restorations, improving the quality of life of millions of patients
and preventing disuse atrophy of the alveolar bone2. The total literature available today
suggests that over a period of 10 years, roughly 1 of 20 implants is lost3. Greater than
the risk for total implant failure are, however, the odds that a technical complication or
an inflammatory condition of the peri-implant tissues may arise. Biological complications
such as peri-implant diseases do occur and are common4. The incidence of peri-implant
diseases is currently a controversial issue. Conflicting statements have been made with
regard to the magnitude and long-term consequences of this problem. Nevertheless, as
the total number of global individuals that undergo restorative therapy through dental
implants increases, so does the importance of considering peri-implantitis as a major
and growing problem in dentistry. Understanding the pathogenesis and etiology of peri-
implantitis, thus, will help clinicians develop clinical strategies to both prevent and treat
this disease.
Peri-Implant Health
Effective endosseous dental implant survival assumes the successful integration
of three peri-implant tissues: bone, connective tissue (CT) and epithelium.
Transmucosal implants in the oral cavity penetrate the soft tissues and form a seal at
the soft tissue interface, which ensures the integrity of the fixture. The implant-gingival
13
tissues have a similar barrier function as the dento-gingival epithelium: to provide a soft
tissue seal to help maintain tissue health5.
The soft tissue that surrounds an implant is referred to as the peri-implant
mucosa while the soft tissue that surrounds a tooth is referred to as the marginal
gingiva. These two types of tissue have several similarities. The junctional epithelium
(JE) and CT attachment of the implant are developed during the healing of the ridge
mucosa, which occurs after implant placement. The epithelial cells are able to attach to
the implant surface via basal lamina and hemidesmosomes in order to establish an
epithelial barrier, or sulcus6. This sulcus is lined by epithelium that is only a few cell
layers thick and is continuous apically with the JE7. Common features between peri-
implant mucosa and marginal gingiva include: pink color, firm consistency, keratinized
oral epithelium, JE or barrier epithelium and hemidesmosomal attachment.
There are also several differences between peri-implant mucosa and marginal
gingiva. The soft tissues around implants form a tight CT barrier, containing a larger
proportion of collagen (85%) and a lower proportion of fibroblasts (15%) than the tissue
adjacent to natural teeth7. Because of the lack of cementum for collagen fiber insertion,
the fibers around dental implants lack a true attachment to the implant. Instead, they are
oriented parallel to the implant surface and attach to the periosteum whereas the fibers
around teeth are oriented perpendicular and are attached to the cementum7-9.
The term biologic width is used when describing the soft tissue dimensions
around teeth and implants5. In 1961, Gargiulo et al identified the biologic width of the
periodontium to be around 3 mm10. The biologic width of a tooth is composed of a
sulcus averaging 0.69 mm, a JE averaging 0.97 mm, and a CT attachment averaging
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1.07 mm. Conversely, the peri-implant mucosa is made up of JE, which is on average 2
mm, and a CT zone which is usually about 1-1.5 mm long, resulting in a biologic width
of 3-3.5 mm8, 11.
The vascular blood supply to a natural tooth also varies from that of a dental
implant. Unlike natural teeth, osseointegrated implants lack a periodontal ligament
(PDL). As a result, periosteum is the sole source of nutrients and this low vascularity of
the peri-implant tissues could be a reason for the increased inflammation around an
implant. Instead of a PDL, mineralized bone grows in close proximity along the length of
the implant surface, resulting in a direct structural and functional connection between
the bone and implant surface, otherwise known as osseointegration12.
Peri-Implant Diagnosis
The close monitoring of an implant is critical for its maintenance and treatment
over time. This is completed during a thorough clinical examination and with
radiographs. Periodontal probing successfully monitors the clinical attachment level
(CAL) changes in peri-implant mucosa and may correlate to the radiographic bone
levels. This is particularly true in sites with healthy gingiva and in the absence of
intrabony defects13. Probing depth (PD) increases are associated with increasing levels
of periodontal pathogens, especially implants with probing depths greater than 5 mm. 1
Suppuration around an implant is not usually seen until peri-implant disease has
progressed significantly. It is, however, associated with disease activity at inflamed
implant sites14.
Peri-Implant Disease
Peri-implant disease is a collective term for inflammatory reactions in the tissues
surrounding an implant and has been confirmed to be infectious in nature15. Peri-implant
15
disease represents the most common inflammatory complication in orofacial
implantology16. Peri-implant disease includes peri-implant mucositis and peri-implantitis.
Peri-implant mucositis describes the inflammatory change that surrounds an implant
and is confined to the soft tissue only with no signs of loss of supporting alveolar bone.
The classic signs of gingival inflammation include swelling and redness. Peri-implant
mucositis has been reported to occur in about 80% of subjects and involving 50% of
implant sites15, 17. Peri-implant mucositis is a reversible condition, but if left untreated, it
may progress to peri-implantitis17.
Peri-implantitis describes the progressive peri-implant alveolar bone resorption
that occurs in conjunction with soft tissue inflammatory lesions18. It has been reported to
occur in between 28% and 56% of subjects and involving 12% to 40% of implant sites15.
Clinically, it presents as the loss of crestal peri-implant bone in conjunction with
bleeding on probing (BOP). It is associated with increased peri-implant PD, mucosal
recession and suppuration. The clinical and histopathologic features of periodontitis and
peri-implantitis are similar and include erythema, pocket formation, mobility, BOP, CT
changes, inflammatory infiltrate and osteolytic bone loss19.
A risk indicator is a probable risk factor, but the cross sectional data is weaker
than the results of longitudinal studies. Some of the risk indicators for peri-implantitis are
poor oral hygiene, a history of periodontitis, diabetes, and smoking15, 20, 21. A meta-
analysis revealed that the odds for implant survival were significantly higher in subjects
without a history of periodontal disease22. Other contributing factors of peri-implantitis
include cement trapped below the gingiva, inadequate seating of the restoration on the
abutment, over contouring the restoration, implant malpositioning, occlusal overload,
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implant texture and composition, and endodontic pathology from a residual granuloma
or a neighboring tooth2, 15, 20, 23-25.
Despite research into peri-implantitis, the exact mechanisms involved have not
been elucidated. Available evidence suggests that pathogenic organisms that inhabit
oral biofilms are important triggers to this disease, whereas much of the tissue
destruction is the result of adaptive and innate host-mediated immune responses to
biofilm pathogens16. Kumar et al recently examined the microbial signatures of the peri-
implant microbiome in health and disease26. The peri-implant microbiome was found to
differ significantly from the periodontal community in both health and disease in that it
demonstrated significantly lower diversity, however, several species, including
previously unsuspected and unknown organisms, were unique to this niche.
Peri-implantitis was established as a microbially heterogeneous infection with
predominantly gram-negative species, and was noted to be less complex than
periodontitis26. The predominant species in peri-implant communities belonged to the
genera Butyrivibrio, Campylobacter, Eubacterium, Prevotella, Selenomonas,
Streptococcus, Actinomyces, Leptotrichia, Propionibacterium, Peptococcus,
Lactococcus, and Treponema. When compared to healthy implants, peri-implant
disease was associated with lower levels of Prevotella and Leptotrichia and higher
levels of Actinomyces, Peptococcus, Campylobacter, non-mutans Streptococcus,
Butyrivibrio and Streptococcus mutans. When compared to periodontitis-associated
biofilms, these communities also demonstrated lower levels of Prevotella, non-mutans
Streptococcus, Lactobacillus, Selenomonas, Leptotrichia, Actinomyces and higher
17
levels of Peptococcus, Mycoplasma, Eubacterium, Campylobacter, Butyrivibrio,
Streptococcus mutans, and Treponema.
While peri-implant mucositis represents the host response of the peri-implant
tissues to the bacterial infection that is not fundamentally different from gingivitis, the
initiation and progression of peri-implantitis may differ from periodontitis. The progression
of peri-implantitis occurs more quickly and more extensively than its periodontitis
counterpart27. Additionally, peri-implantitis may differ from periodontitis both in the extent
and the composition of extensive inflammatory cells in the lesion28, 29. The treatment of
these two disease entities should be similar; debridement with implant-safe hand
instruments and strict home care oral hygiene22.
Animal Studies: Ligature-Induced Defect Model
The association between bacterial plaque biofilm formation and the pathogenesis
of peri-implant diseases has been demonstrated in animal studies using a ligature-
induced defect model. In this experimental model, peri-implant mucositis and peri-
implantitis lesions are induced by terminating the plaque control regimen followed by the
placement and exchange of ligatures around the implant neck in a circumferential,
submucosal position9, 30. At peri-implantitis sites, this active breakdown period is usually
terminated by ligature removal, which is in the majority of sites associated with
spontaneous disease progression (i.e. progression period)4. The basic methodological
procedure of the ligature-induced defect model includes the phases listed in Table 1-131.
It is important to realize that the ligature-induced breakdown is a technical part of
an experimental invasive procedure to achieve a certain degree of breakdown – the final
result of which will mimic a natural peri-implantitis lesion. Thus, it is not an ideal model to
study progression, because the investigator can control the ligature-induced breakdown
18
process. The type of ligature, the coronal-apical position of the ligature and how often the
ligature is replaced during the plaque formation period determine the rate and the amount
of breakdown. Thus, ligature-induced tissue destruction does not accurately represent in
vivo peri-implant pathogenesis32.
Animal Models: Peri-Mucositis
The soft-tissue reaction to undisturbed plaque formation at osseointegrated
titanium implants has been evaluated mainly in the canine model but also in nonhuman
primates30, 33-36. In these studies, peri-implant mucositis lesions were commonly induced
by terminating the oral hygiene regimen, thus allowing undisturbed accumulation of
plaque. Clinical observation at mucositis lesions confirmed the presence of gross
amounts of bacterial plaque biofilms in both supra- and submucosal compartments
adjacent to the implant surface. This was commonly associated with pocket formation
and concomitant inflammation (redness, edema and BOP) of the soft tissues.
Histological evaluation revealed an ulcerated pocket epithelium (PE) and the
development of an inflammatory cell infiltrate, which was comparable in both size and
composition to experimental gingivitis lesions. In particular, the CT components at
inflamed implant sites revealed a decrease in collagen and an increase in vascular
structures, fibroblasts, leukocytes and residual tissue30.
Animal Models: Peri-Implantitis
Most research studies on the pathogenesis of ligature-induced peri-implantitis
used the dog as the experimental animal of choice. Similarly, most studies have
employed the dog for research on the treatment of peri-implantitis. The mean active
breakdown period (mean ligature application/exchange) was 12.0 ± 5.0 weeks. The
mean bone loss in these studies was 41.6 ± 16.1% relative to the original implant
19
length32. Microbiologic analysis revealed an increased level of P. gingivalis, Prevotella
intermedia and T. forsythia37, 38. Histologically, peri-implantitis lesions in dogs were
characterized by the presence of a large inflammatory cell infiltrate residing in the peri-
implant mucosa, but also extending into the alveolar bone. This was associated with
ulceration of the PE and loss of implant-supporting alveolar bone. In comparison with
experimental periodontitis lesions, the inflammatory cell infiltrate at implants revealed
small amounts of collagen, but a proportionally higher vasculature and larger volumes of
PMN leukocytes and plasma cells. In particular, the percentage volume of the
inflammatory cell infiltrate revealed the following major components: collagen, vascular
structure, fibroblasts, macrophages, lymphocytes, plasma cells, PMN leukocytes and
residual tissue9.
Animal Studies: Animal Selection
The literature reporting on experimental studies aimed at investigating the
pathogenesis and therapy of both peri-implant mucositis and peri-implantitis commonly
employ large animal models. These mainly include canine models, but also include
nonhuman primate and swine (mini pig) models. The canine model is the large animal
model most frequently used to investigate either the pathogenesis or treatment of peri-
implant disease. Most breeds of dog exhibit a natural susceptibility to periodontitis39. The
bone composition (i.e. water, organic, volatile inorganic and ask fractions) of canine
alveolar bone is very similar to that of human bone. The specific anatomic
characteristics of the canine jaw bone usually facilitate the insertion of common dental
implants (i.e. length = 10 mm; diameter = 3-4 mm).
Nonhuman primates possess similar oral structures to humans and also exhibit
naturally occurring bacterial plaque biofilms and a susceptibility to develop gingivitis.
20
Interestingly, in the case of undisturbed accumulation of plaque, spontaneous
progression of the inflammatory cell infiltrate and subsequent ongoing CAL loss was
found to be limited to implants and ankylosed teeth and did not occur in relation to
control teeth exhibiting an intact PDL40. Accordingly, nonhuman primates have been
frequently employed for studies on the pathogenesis and treatment of peri-implant
disease. However, the specific anatomic characteristics of the nonhuman primate jaw
bone do require the insertion of diameter-reduced dental implants. As a result of strict
regulatory requirements and demanding acquisition and maintenance costs, nonhuman
primates are nowadays rarely used in dental implant research40.
Both the macro- and microstructure of pig bone are moderately similar to those of
human bone. Most similarities between pig and human bone were observed for either
bone composition or bone remodeling (1.2–1.5 μm/day for pig vs. 1.0–1.5 μm/day for
human)41. Similarly to the dog, the specific anatomic characteristics of the pig jaw bone
facilitate the insertion of dental implants of the same dimensions as those used in
humans42.
Rodent models have several useful features for investigating molecular
mechanisms involved in the pathogenesis of infection-driven inflammatory diseases43.
In relation to the process of periodontal disease, rodent models have demonstrated that
periodontal bacteria play an essential role in initiating gingival inflammation and
periodontal bone loss and that the acquired immune response contributes significantly
to periodontal tissue destruction44, 45. Much of this critical insight into mechanisms
through which periodontal pathogens lead to attachment loss and ABR would have
been considerably more difficult or expensive in other animal models. A rodent model
21
may have a great potential to enhance our knowledge on molecular and cellular
mechanisms for the pathogenesis of peri-implant diseases.
Recently, it has been demonstrated that P. gingivalis, T. denticola, and T. forsythia
can colonize the rat and mice gingival cavity, induce gingival inflammation with induction
of enhanced IgG and IgM immune responses, and cause significant ABR, characteristic
of polymicrobial periodontitis46-50. To the best of our knowledge, there is limited data from
rodent models available for evaluating the biofilm-mediated pathogenesis of peri-implant
diseases19.
Study Aim
The aim of this study is to develop a novel peri-implantitis model in rats by using
a previously established rat model of polymicrobial infection.
Study Hypothesis
The hypothesis is that major periodontal bacteria P. gingivalis, T. denticola and
T. forsythia will colonize around titanium implants, induce gingival inflammation, and
cause ABR, leading to peri-implantitis in a rat model.
22
Table 1-1 Methodological procedure of standard animal ligature-induced defect models Clinical Phase Clinical Procedure 1 Tooth extraction 2 Healing period I 3 Placement of implants 4 Healing period II 5 6
Implant uncovery Healing period III
7 Active breakdown period: ligature placement
8 Progression period: after ligature removal
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CHAPTER 2 MATERIALS AND METHODS
Rats
This study protocol was approved by the Institutional Animal Care and Use
Committee of the University of Florida (Protocol #201408568). Twelve 5-week-old male
Wistar rats (weighing around 150 grams) were used. Rats were maintained in groups,
housed under microisolator conditions, fed standard powdered chow, given H2O ad
libitum and kept at 25°C with alternating 12h periods of light and darkness. The design
of the study is described in Fig. 2-1. The rats were divided into two experimental groups.
Rats of Group A (n=6) were exposed to polymicrobial infection and rats of Group B
(n=6) served as sham-infection.
Surgical Procedures
Surgical protocol is described with surgical photos in Fig. 2-2. During all surgical
procedures, general anesthesia was induced by intramuscular injection of 1 mg/kg of a
solution containing 100 mg/ml ketamine and 20 mg/ml xylazine. Bilateral extraction of
maxillary first molars was performed. One month after healing, mucoperiosteal flaps
were elevated at the edentulous maxillary molar region and osteotomies were prepared
with a 1.7 mm diameter reamer. Custom made, pure titanium, machined surface
implants with 1.5 mm diameter and 2 mm length were installed (JMR, Niigata, Japan)
(Fig. 2-3). The alveolar mucosa was reapproximated with absorbable sutures (5.0
chromic gut) and implants were submerged. Two months following implant installation,
implants were uncovered and 1 mm custom healing abutments were fixed to the
implants using cyanoacrylate in order to prevent abutment loosening (Fig. 2-3). One
24
week following healing, oral infection with 3 bacteria was initiated for Group A rats.
Sham-infection was initiated for Group B rats with the control vehicle.
Bacterial Strains and Polymicrobial Inocula
P. gingivalis ATCC 53977, T. denticola ATCC 35404, and T. forsythia ATCC
43037 were routinely cultured anaerobically as well as maintained for infection as
described previously46, 50, 51. Bacterial concentration was determined by OD600 and the
organisms were suspended in equal proportions in reduced transport fluid at 1010
cells/ml. To prepare the bacterial suspension for infection, P. gingivalis (3.3 x 109) cells
were gently mixed with an equal volume of T. denticola (3.3 x 109) cells and allowed to
interact for 5 min. Subsequently, T. forsythia (3.3 x 109) cells were added to the tube
and gently mixed for 1-2 min and allowed to interact for an additional 5 min. The
polymicrobial mixture was then mixed thoroughly with an equal volume of 4% (w/v)
sterile low viscosity carboxymethylcellulose (CMC; Sigma-Aldrich, St. Louis, MO, USA)
and 0.5 ml (5 x 109 bacteria/ml) was administered by oral lavage to Group A rats using
isoflurane inhalation anesthesia.
Peri-Implantitis Induction with P. gingivalis, T. denticola and T. forsythia
All rats were given kanamycin (500 μg/ml) for 4 days in the drinking water46, 51, 52.
After a 3-day antibiotic washout period, the gingival surface was swabbed with 0.12%
chlorhexidine gluconate (Peridex, Proctor and Gamble, Cincinnati, OH) mouth rinse to
inhibit the endogenous organisms and to promote subsequent colonization of P.
gingivalis, T. denticola and T. forsythia46, 51, 52. One day after swabbing with chlorhexidine,
polymicrobial inocula were administered by oral lavage for 4 consecutive days per week
on 6 alternate weeks for a total of 24 inoculations during 12 weeks of the experimental
infection period (Group A). Control rats (Group B) received vehicle (sterile 4% CMC) only.
25
Gingival Plaque Sampling
To confirm the presence of P. gingivalis, T. denticola, and T. forsythia on the
gingival surface of implant, infected rats as well as the absence of these species on the
gingival surface of implant, sham-infected rats, gingival plaque samples were collected
using sterile veterinary cotton swabs gently rubbed around the abutment surfaces and
immersed in Tris-EDTA buffer46. In order to determine gingival colonization of bacteria
with minimal disruption of the biofilms, a total of two post-infection samples were
collected 3 days following the 4th and 5th infections (at the 8th and 10th week) from
infected and sham-infected rats.
Detection of Bacterial Genomic DNA in Gingival Plaque Samples
Polymerase chain reaction (PCR) was performed to detect the presence of
bacterial species in the gingival plaque samples as described previously46, 51, 52. Colony
PCR was carried out with a Bio-Rad thermal cycler and by using the following 16S
rRNA bacterial gene species-specific oligonucleotide forward and reverse primers: (P.
gingivalis): 5’-TGTAGATGACTGATGGTGAAAACC-3’ (forward), 5’-
ACGTCATCCCCACCTTCCTC-3’ (reverse); (T. denticola) 5’-
TAATACCGAATGTGCTCATTTACAT-3’ (forward), 5’-
CTGCCATATCTCTCTGTCATTGCTCTT-3’ (reverse); (T. forsythia) 5’-
AAAACAGGGGTTCCGCATGG-3’ (forward), 5’-TTCACCGCGGACTTAACAGC-3’
(reverse). PCR products were separated by agarose (1.5%) gel electrophoresis. The
genomic DNA extracted from P. gingivalis, T. denticola, and T. forsythia served as
positive controls and PCR performed with no template DNA served as the negative
control.
26
Euthanasia and Tissue Specimen Collection
After 6 infection cycles, rats were euthanized via CO2 asphyxiation and blood
was collected by cardiac puncture. Sera were separated and stored at -20°C for
bacterial specific immunoglobulin G (IgG) and immunoglobulin M (IgM) antibody
analysis.
Serum IgG and IgM Antibody Analysis
Serum collected from the rats was used to determine IgG and IgM antibody
concentrations against each of the 3 bacteria using a standard Enzyme-linked
immunosorbent assay (ELISA) protocol53. Briefly, P. gingivalis, T. denticola and T.
forsythia cells were treated overnight with 0.5% formalin in buffered saline. These cells
were then washed, diluted to OD600 of 0.3 and coated in microtiter plate wells. Diluted
rat sera (1:100 for IgG and 1:100 for IgM) were allowed to react with the bacterial
antigen for 2h at room temperature. After washing, the goat anti-rat IgG and IgM
secondary antibodies conjugated to alkaline phosphatase (1:5,000) (Bethyl
Laboratories, Montgomery, TX) were added to the plates and the assay was developed
with p-nitrophenolphosphate (Sigma-Aldrich). By using 3M NaOH, the assay reactions
were terminated and analyzed at OD405 using a Bio-Rad microplate reader.
Morphometric Analysis of Mandibular Lingual Alveolar Bone Resorption
The horizontal ABR was measured on the lingual of mandibular molars by
morphometry as described previously53. Briefly, the mandibles were autoclaved and
defleshed and immersed in 3% (vol/vol) hydrogen peroxide for 3h. Cementum was not
stained with methylene blue prior to measuring ABR. Digital images of lingual root
surfaces of all mandibular molar teeth were captured under a 10X stereo dissecting
microscope (SteReo Discovery V8; Carl Zeiss Microimaging Inc, Thornwood, NY). The
27
line tool was used to measure the horizontal ABR from the cementoenamel junction
(CEJ) to the alveolar bone crest (ABC). The surface perimeters of CEJ and ABC were
traced using the calibrated line tool (AxioVision LE 29A software version 4.6.3.) and the
horizontal ABR area in mm2 was instantly imprinted over the digital image (Fig. 2-4).
Two blinded examiners performed all measurements twice at separate times.
Microcomputed Tomography (microCT) Peri-Implant Bone Levels
Three-dimensional peri-implant bone levels in the implant sham-infected and
implant infected groups were evaluated by microCT radiographic analysis. The maxillae
were fixed in 10% buffered formalin in phosphate-buffered. The specimens were
transferred to 70% ethanol. Each rat maxilla was scanned using a high resolution
microCT100 system (Scano Medical, Brüttisellen, Switzerland) at isotropic voxel size:
17.2μm, energy: 90kVp, current: 200μA, integration time: 300ms and filter: 0.5mmAI.
The peri-implant bone volume was measured using ImageJ software (NIH, Bethesda,
MD, USA) in frontal (buccal/lingual) and sagittal (mesial/distal) sections of the implants
as the distance from the implant platform (IP) to the first bone to implant contact (BIC)
(Fig. 2-5).
Histological Evaluation of Gingival Inflammation
The maxillae of the implant sham-infected and implant infected groups were fixed
in 10% buffered formalin and then decalcified in phosphate-buffered saline containing 0.4
M EDTA and 2% formaldehyde. The solution was changed every other day for six weeks.
Implants with healing abutments were then gently removed by unscrewing with pliers.
The specimens were then processed for embedding in paraffin and sectioning using
routine histology protocols. The specimens were sectioned (7 µm) in the mesio-distal
direction. Every fifth slide from each block of sections was stained with hematoxylin and
28
eosin. Stained sections were used to assess the migration of peri-implant epithelium
(PIE), CT, ABC and the level of inflammation by light microscopy. Using a digital camera,
images were captured from stained sections and transferred to Adobe® Photoshop®.
Statistical Analysis
Statistical analysis of bacterial-specific serum antibody levels and horizontal ABR
was performed using Graph Pad Prism Software for Windows, version 5. A value of
p<0.05 was considered statistically significant. Antibody analysis and horizontal ABR
data are presented in figures as means ± standard deviations. Unpaired, two-tailed
Student’s t test was used to compare the infected and sham-infected groups.
For antibody analysis, mean bacterial-specific antibody titers of infected rats
were divided by mean antibody titers of sham-infected rats and the quotient represents
the fold change in mean bacterial specific antibody titers produced as a result of
polymicrobial infection. The graphs demonstrate mean fold increase of bacterial-specific
antibody titers of the infected rats compared to the sham-infected rats.
MicroCT measurements were analyzed by the Mann-Whitney U test, using
implants as the statistical unit, to evaluate differences between the two groups
regarding peri-implant bone levels. Statistical significance was given at a value of
p<0.05.
29
Figure 2-1 Experimental design. After all surgical procedures, polymicrobial or sham infection was administered for 4 consecutive days per week on 6 alternate weeks for a total of 24 inoculations. Six polymicrobial infections (Infection I - VI) are indicated at 1, 3, 5, 7, 9 and 11 weeks.
30
Figure 2-2 Surgical protocol. A) Bilateral maxillary M1 extractions. B) After 1 month of
healing, full thickness flap was elevated in previous extraction site and osteotomies were prepared. C) Implant placement was performed in osteotomies. D) After 2 months of healing, full thickness flap was elevated and healing abutments were connected to implants. Photos courtesy of author.
31
Figure 2-3 Implant & healing abutment. A) Machined surface implant. Measurements are shown in millimeters. B) Healing abutment. C) Comparison of extracted M1 molar adjacent to implant. Photos courtesy of author.
32
Figure 2-4 Morphometric evaluation of horizontal alveolar bone resorption.
Representative left mandible showing the lingual horizontal alveolar bone resorption area by morphometry in A) infected and B) sham-infected rats. The area outlined between CEJ-ABC represents the area of horizontal ABR in mm2. M1, M2, M3 are molars. Rat jaw images captured at 10X magnification. Photos courtesy of author.
33
Figure 2-5 Representative microCT image of an implant-abutment complex. Landmarks used to measure peri-implant bone level in the A) sagittal (mesial, distal) and B) frontal (buccal, lingual) planes. IP indicates implant platform. BIC indicates first bone to implant contact. Photos courtesy of author.
34
CHAPTER 3 RESULTS
Clinical Results of Implant and Healing Abutment Placement
One rat in Group A (polymicrobial infection) and two rats in Group B (sham-
infection) were euthanized due to corneal infection following the mini-implant installation
surgical procedure. In addition, 2 more rats in Group B died due to aspiration of blood
after the implant installation procedure and during the recovery time. At the time of
implant uncovery, two rats in Group A and one rat in Group B had one implant fail to
integrate each. At the end of the study, two more rats in Group A had both implants fail.
Thus, at the end of the experiment, 3 rats in the Group A with 4 implants and 2 rats in
the Group B with 3 implants were available for microCT and histologic evaluation.
Additionally, there was 1 rat in Group A with 2 healing abutments and 2 rats in Group B
with 3 healing abutments (Fig. 3-1; Table 3-1).
Peri-Implantitis Induction with Periodontal Bacteria
The implanted, polymicrobial-infected rats (Group A) were swabbed 3 days after
the 4th and 5th infections to confirm infection by three periodontal bacteria. PCR using
bacterial-specific primers revealed all rats positive for P. gingivalis, T. denticola and T.
forsythia genomic DNA, indicating successful colonization of all three periodontal
bacteria on the gingival surface around the implants (Table 3-2). None of the implanted,
sham-infected rats (Group B) showed genomic DNA for the three bacteria.
Polymicrobial Infection Elicits Humoral Antibody Response
Serum levels of IgG and IgM antibodies in implanted, polymicrobial-infected rats
(n=5) and implanted, sham-infected rats (n=2) were measured. Polymicrobial infection
induced significant serum IgG and IgM antibodies (p<0.05) against all the three bacteria
35
compared to sham-infected rats (Fig. 3-2). These results corroborated the successful
generation of humoral immune response after polymicrobial infection in implanted rats.
Polymicrobial Infection Elicits Horizontal Mandibular Alveolar Bone Resorption
Mandibular lingual horizontal ABR area was measured to confirm ABR following
polymicrobial infection. The mandibles of the polymicrobial-infected rats (n=5) had
significant (p<0.05) ABR when compared to those of the sham-infected rats (n=2) (Fig.
3-3). These results indicate statistically significant induction of ABR in the infected rats.
MicroCT Imaging Analysis of Peri-Implant Bone
To understand the effects of polymicrobial infection on peri-implant bone levels,
microCT imaging was performed on all rat maxillae and bone levels were measured
from the IP to the first BIC. Mean peri-implant bone levels for the implants in the
polymicrobial-infected rats (n=4) and the implants in the sham-infected rats (n=3) are
illustrated in Table 3-3. MicroCT images for representative infected and sham-infected
implants are depicted in Fig. 3-4. There was a greater mean distance from the IP to the
first BIC in the infected rats compared to the sham-infected rats (0.80 ± 0.37 mm vs.
0.48 ± 0.13 mm, respectively). This result was not statistically significant (p<0.05).
Peri-Implant Histology
To evaluate the peri-implant inflammation, histology was performed on the
maxillae of the specimens in the control and infected groups. In the control group, three
implants were retained in two animals for histological assessment (75% implant survival
rate). Peri-implant epithelium was composed of extended keratinized oral epithelium with
no resemblance of typical tooth lining JE (Fig. 3-5 A-C). The CT immediately underneath
the epithelium was practically free of inflammation. The CT above the bone and below the
PIE that leans against the implant abutment had a mild inflammatory infiltrate with
36
capillaries and many PMN cells. Implant-bone contact was well preserved in all control
specimens (Fig. 3-5C).
In the infected group, four implants were recovered in three animals for histological
assessment (40% implant survival rate). The histological view of two implants in one rat in
this group was similar to the control implants (not shown). However, the other two
infected implants showed extensive peri-implantitis with advanced bone loss and
extensive inflammation (Fig. 3-5 D-F). Interestingly, the location of the PIE in these
specimens was comparable to the sham-infected specimens. However, the PIE appeared
to be more blunt and thickened in the infected group. There were no signs of formation of
PE in the infected samples. Intriguingly, bone contact (BC) to the implant threads was
often minimal and predominantly replaced by inflammatory granulation tissue (GT) with
PMN leukocytes (Fig. 3-5F).
37
Table 3-1 Remaining implants and healing abutments in infected and sham-infected groups according to microCT images
Rat # Implants Abutments
left right left right
Infe
cted
1 - + - -
2 - - - -
3 - + - -
4 + + + +
5 - - - -
Sha
m-
Infe
cted
1 - + - +
2 + + + +
38
Table 3-2 Distribution of rat gingival microbial samples positive for Pg/Td/Tf by PCR
Group Rats
No. of gingival microbial samples positive for PCR P. gingivalis T. denticola T. forsythia
8th week
10th week
8th week
10th week
8th week
10th week
Infected 5 4 5 4 5 3 5 Sham-Infected 2 0 0 0 0 0 0
39
Table 3-3 Mean peri-implant bone levels in infected and sham-infected rats Mean (mm) SD Implants and infection
(n=4 implants) 0.80 0.37
Implants and sham-infection (n=3 implants)
0.48 0.13
40
Figure 3-1 Rat maxillae showing presence or absence of healing abutments at
conclusion of polymicrobial infection. Red arrows indicate intact implant visible on microCT. A) Infected rats (n=5). Four implants and two healing abutments present. B) Control rats (n=2). Three implants and three healing abutments present. Photos courtesy of author.
41
Figure 3-2 Polymicrobial infection induced bacterial specific IgG and IgM antibody
response. The graphs demonstrate the fold change in mean bacterial-specific A) IgG antibody titer and B) IgM antibody titer as a result of polymicrobial infection in comparison to sham-infected rats. The humoral immune response in implanted, infected rats was significantly greater than implanted, sham-infected rats.
42
Figure 3-3 Mandibular lingual alveolar bone resorption. Polymicrobial infection induced
significant (p<0.05) mandibular lingual ABR around the molars of the infected rats compared to the control rats.
43
Figure 3-4 Peri-implant microCT alveolar bone resorption. Representative microCT
images of peri-implant bone levels in two rats – implant + polymicrobial-infected and implant + control. Left and right implants are visible in the sagittal plane. M2, M3 represent remaining molars. Implants in the infected group had a greater mean distance from the IP to the first BIC compared to those in the control group. Photos courtesy of author
44
Figure 3-5 Histological staining of the peri-implant tissues. Histological representative
images in the control (A-C) and infected (D-F) rats showing bone loss and inflammation. A) Overview of a control implant showing normal PIE (magnified in B), mild inflamed CT and normal BC with threads clearly visible (BC, magnified in C). D) Overview of an infected implant showing blunt and thickened PIE (magnified in E) but little apical migration. The CT and BC areas have been replaced by GT (D and E). Some thread sites that used to have BC have been predominantly replaced by inflammatory tissue with PMN leukocytes lining the surface that used to have implant contact (F). Arrows in panels B and E point to the termination of the PIE. Photos courtesy of author.
45
CHAPTER 4 DISCUSSION
Peri-implantitis has become a global clinical problem and affects 10-20%
implants at varying degrees17. The peri-implantitis microbiome differs from periodontitis
but still harbors many periodontal pathogens, including those used in this study26, 54.
The pathogenesis of peri-implant lesions remains relatively poorly understood due to
the lack of inexpensive animal models to study the disease. Most animal studies thus
far have been performed with dogs9. Limitations of the dog model include cost related to
the number of animals to be used and emotional dislike of using family pets for research
purposes. Having a small number of animals does not allow dissection of longitudinal
cellular and molecular events that lead to establishment of peri-implant lesions. In
addition, peri-implantitis lesions need to be initiated with ligatures around the implants
that may not truly mimic initiation of peri-implantitis, as discussed.
Animal studies and cross-sectional human histological data collectively suggest
that peri-implantitis lesions differ from periodontitis lesions at least in three aspects55.
First, the formation of barrier PE is limited at the apical aspect of the peri-implantitis
lesion and thus it does not “insulate” the biofilm from the inflammatory infiltrate. This
means that bacteria are in direct contact with inflammatory cells. In our rat model, the
PIE showed practically no migration to form a true PE in the infected peri-implantitis
specimens, thus supporting the previous findings. Second, there are more PMN cells in
the peri-implantitis lesions compared to those with periodontitis. This finding was also
corroborated in our rat model as PMN cells were found lining the implant surface. It is
not surprising, therefore, that peri-implantitis lesions are frequently associated with
suppuration (dead PMN and other leukocytes and bacteria). Third, inflammation
46
extends to the alveolar crest in peri-implantitis while in periodontitis, inflammation is
rarely seen within 1 mm of bone. This finding was also supported in our rat model. In
summary, the rat model with multi-species infection with human pathogens replicates
well the histological observations made previously with the dog models and human
histology data.
The disadvantage of the current rat model was substantial loss of implants or
healing abutments during the course of the experiment. There were a few more
implants lost in the infected group that may reflect loosening of the implant due to peri-
implantitis. However, the difference was not significant so it is more likely that in the
rodent model, more implants are lost due to function and trauma. Softer diet may offer
better implant survival in rodents. The fact that peri-implantitis was advanced in the
infected group 22 weeks after implant placement indicates that shorter experimental
timeframe should be used that may lead to preservation of more implants with initial
peri-implantitis lesions. It is conceivable that 12 weeks will be sufficient for introduction
of initial to moderate peri-implant lesions. Understanding the development of initial to
moderate lesions is likely more important for understanding the pathological process
than further advancement of moderate lesions.
There are several advantages with the current rat model. Increasing the number
of animals for these types of experiments is relatively easy and less expensive than
using larger animals. In addition, the clear advantage is the natural process of disease
without mechanical trauma with ligatures, thus better mimicking natural peri-implantitis
in man. Like in dogs, it is anticipated that not all animals are susceptible to initiation or
advancement of peri-implantitis.
47
In the present study, the peri-implant tissue reaction of implants exposed to
polymicrobial infection was analyzed. It was observed that the amount of bone loss that
occurred at the end of the experimental period was greater for implants exposed to
polymicrobial infection compared to control implants. The histological analysis showed
that implants subjected to polymicrobial infection had peri-implant bone replaced by
granulation tissue with PMN leukocytes lining the surface that used to be in contact with
bone. It is suggested that polymicrobial infection with P. gingivalis ATCC53977, T.
denticola ATCC 35404 and T. forsythia ATCC 43037 was able to induce an
inflammatory reaction to peri-implant tissues and subsequent bone loss in rats.
The present study addressed the feasibility of inducing peri-implantitis in rats
utilizing a polymicrobial infection model. The majority of the information regarding peri-
implant infections is derived from larger animals utilizing a ligature-induced peri-
implantitis model18. However, the ligature-induced breakdown is a technical part of an
experimental invasive procedure to achieve a certain degree of breakdown with the
investigator controlling the ligature-induced breakdown process. The type of ligature,
the coronal-apical position of the ligature, and how often the ligature is replaced during
the plaque formation period determine the rate and the amount of breakdown. Thus,
ligature-induced tissue destruction does not accurately recapitulate in vivo peri-implant
pathogenesis32.
Rodents have been utilized in order to evaluate peri-implant bone reactions
under local conditions such as excessive loading and systemic conditions such as
osteoporosis and diabetes56-59. There is limited information for evaluation of peri-
implant soft and hard tissue responses to bacterial challenge in rodents. Freire et al.
48
utilized implants containing an established biofilm contaminated with A.
actinomycetemcomitans and implanted in rats19. It was demonstrated that after 6
weeks, implants containing the A. actinomycetemcomitans biofilm expressed greater
inflammation assessed by clinical means and greater bone volume loss assessed by a
microCT. In the current experiment, we have utilized a polymicrobial infection model
with P. gingivalis, T. denticola and T. forsythia that has been extensively used in the
induction of periodontitis in rats and mice46-50, 53. This consortium of bacteria has
been identified as the hallmark of the pathogenic biofilm in deep periodontal pockets in
chronic periodontitis and stimulates maxillary and mandibular ABR in rats.46, 60 We
documented for the first time that this polymicrobial consortium has the potential to
induce peri-implant bone resorption and inflammation in rats.
Although earlier studies have reported putative periodontal pathogens such as P.
gingivalis, P. intermedia and A. actinomycetemcomitans to be present in the majority of
peri-implant lesions examined, more current studies have reported distinct differences in
the microbiological profiles of periodontal and peri-implant lesions26, 54, 61. Thus, in
peri-implant areas, Staphylococci, enterococci and yeasts were found almost as
frequently as periodontopathogens, indicating differences as compared to the
microbiota around affected teeth. The polymicrobial infection model used in this study
may facilitate better understanding of the role of different bacteria in the pathogenesis of
peri-implantitis.
In the present study, machined surface implants were used. It has been reported
that implant surface characteristics can influence the progression of peri-implantitis, with
modified implant surfaces demonstrating greater rate of ligature-induced peri-implantitis
49
progression compared to machined surfaces55, 62. However, the influence of implant
surface characteristics on peri-implantitis progression has not been fully understood in
physiological infection-induced models or in clinical observations. Thus, a physiological
induction of peri-implantitis model such as a polymicrobial infection model is highly
relevant in further understanding the effect of implant surface characteristics on the
pathogenesis and progression of peri-implantitis.
The amount of bone loss observed for implants subjected to polymicrobial
infection was greater compared to control implants, but the difference was not
statistically significant. This may be due to the small sample size of this pilot study. The
experiment initiated with 6 rats per group having 12 implants total. At the end of the
experiment, only 3 animals with 4 implants in the test group and 2 animals with 3
implants in the control group were available for analysis. As this is a major limitation of
the study, the current findings need to be confirmed with further studies. The histological
observations, however, confirm the inflammatory changes in the peri-implant area and
validate the findings despite the small sample size.
In summary, within the limitations of this study, it was demonstrated that it is
feasible to induce peri-implant infection with subsequent peri-implant bone loss utilizing
a polymicrobial infection model with P. gingivalis, T. denticola and T. forsythia.
Additional studies with a larger sample size and shorter duration of infection period
should be explored. The current rat model for peri-implantitis with polymicrobial infection
with human pathogens could offer significant advantages for studies of initiation and
progression of peri-implantitis and the role of implant, host and microbial characteristics
on the peri-implantitis disease process.
50
LIST OF REFERENCES
1. Branemark PI, Adell R, Breine U, Hansson BO, Lindstrom J, Ohlsson A. Intra-osseous anchorage of dental prostheses. I. Experimental studies. Scand J Plast Reconstr Surg 1969;3:81-100.
2. Isidor F. Influence of forces on peri-implant bone. Clin Oral Implants Res 2006;17 Suppl 2:8-18.
3. Mombelli A, Muller N, Cionca N. The epidemiology of peri-implantitis. Clin Oral
Implants Res 2012;23 Suppl 6:67-76. 4. Zitzmann NU, Berglundh T, Ericsson I, Lindhe J. Spontaneous progression of
experimentally induced periimplantitis. Journal of clinical periodontology 2004;31:845-849.
5. Cochran DL, Hermann JS, Schenk RK, Higginbottom FL, Buser D. Biologic width
around titanium implants. A histometric analysis of the implanto-gingival junction around unloaded and loaded nonsubmerged implants in the canine mandible. J Periodontol 1997;68:186-198.
6. Lindhe J, Berglundh T. The interface between the mucosa and the implant.
Periodontol 2000 1998;17:47-54. 7. Berglundh T, Abrahamsson I, Lindhe J. Bone reactions to longstanding functional
load at implants: an experimental study in dogs. Journal of clinical periodontology 2005;32:925-932.
8. Berglundh T, Lindhe J, Ericsson I, Marinello CP, Liljenberg B, Thomsen P. The
soft tissue barrier at implants and teeth. Clin Oral Implants Res 1991;2:81-90. 9. Lindhe J, Berglundh T, Ericsson I, Liljenberg B, Marinello C. Experimental
breakdown of peri-implant and periodontal tissues. A study in the beagle dog. Clin Oral Implants Res 1992;3:9-16.
10. Gargiulo AW. Dimensions and Relations of the Dentogingival Junction in
Humans. Journal of Periodontology 1961:261-267. 11. Berglundh T, Lindhe J. Dimension of the periimplant mucosa. Biological width
revisited. Journal of clinical periodontology 1996;23:971-973. 12. Rose L, Mealey BL. Periodontics Medicine, Surgery, and Implants. St-Louis,
Elsevier Mosby: E. Mosby; 2004. 13. Quirynen M, van Steenberghe D, Jacobs R, Schotte A, Darius P. The reliability of
pocket probing around screw-type implants. Clin Oral Implants Res 1991;2:186-192.
51
14. Rams TE, Roberts TW, Tatum H, Jr., Keyes PH. The subgingival microbial flora associated with human dental implants. J Prosthet Dent 1984;51:529-534.
15. Lindhe J, Meyle J, Group DoEWoP. Peri-implant diseases: Consensus Report of
the Sixth European Workshop on Periodontology. Journal of clinical periodontology 2008;35:282-285.
16. Offenbacher S, Barros SP, Singer RE, Moss K, Williams RC, Beck JD.
Periodontal disease at the biofilm-gingival interface. J Periodontol 2007;78:1911-1925.
17. Zitzmann NU, Berglundh T. Definition and prevalence of peri-implant diseases.
Journal of clinical periodontology 2008;35:286-291. 18. Berglundh T, Stavropoulos A, Working Group 1 of the VEWoP. Preclinical in vivo
research in implant dentistry. Consensus of the eighth European workshop on periodontology. Journal of clinical periodontology 2012;39 Suppl 12:1-5.
19. Freire MO, Sedghizadeh PP, Schaudinn C, et al. Development of an animal
model for Aggregatibacter actinomycetemcomitans biofilm-mediated oral osteolytic infection: a preliminary study. J Periodontol 2011;82:778-789.
20. Heitz-Mayfield LJ. Peri-implant diseases: diagnosis and risk indicators. Journal of
clinical periodontology 2008;35:292-304. 21. Rocchietta I, Nisand D. A review assessing the quality of reporting of risk factor
research in implant dentistry using smoking, diabetes and periodontitis and implant loss as an outcome: critical aspects in design and outcome assessment. Journal of clinical periodontology 2012;39 Suppl 12:114-121.
22. Safii SH, Palmer RM, Wilson RF. Risk of implant failure and marginal bone loss
in subjects with a history of periodontitis: a systematic review and meta-analysis. Clin Implant Dent Relat Res 2010;12:165-174.
23. Quirynen M, Vogels R, Alsaadi G, Naert I, Jacobs R, van Steenberghe D.
Predisposing conditions for retrograde peri-implantitis, and treatment suggestions. Clin Oral Implants Res 2005;16:599-608.
24. Misch CE, Perel ML, Wang HL, et al. Implant success, survival, and failure: the
International Congress of Oral Implantologists (ICOI) Pisa Consensus Conference. Implant Dent 2008;17:5-15.
25. Isidor F. Histological evaluation of peri-implant bone at implants subjected to
occlusal overload or plaque accumulation. Clin Oral Implants Res 1997;8:1-9.
52
26. Kumar PS, Mason MR, Brooker MR, O'Brien K. Pyrosequencing reveals unique microbial signatures associated with healthy and failing dental implants. Journal of clinical periodontology 2012;39:425-433.
27. Lang NP, Berglundh T, Working Group 4 of Seventh European Workshop on P.
Periimplant diseases: where are we now?--Consensus of the Seventh European Workshop on Periodontology. Journal of clinical periodontology 2011;38 Suppl 11:178-181.
28. Pesce P, Menini M, Tealdo T, Bevilacqua M, Pera F, Pera P. Peri-implantitis: a
systematic review of recently published papers. Int J Prosthodont 2014;27:15-25. 29. Kantarci A, Hasturk H, Van Dyke TE. Animal models for periodontal regeneration
and peri-implant responses. Periodontol 2000 2015;68:66-82. 30. Berglundh T, Lindhe J, Marinello C, Ericsson I, Liljenberg B. Soft tissue reaction
to de novo plaque formation on implants and teeth. An experimental study in the dog. Clin Oral Implants Res 1992;3:1-8.
31. Schwarz F, Iglhaut G, Becker J. Quality assessment of reporting of animal
studies on pathogenesis and treatment of peri-implant mucositis and peri-implantitis. A systematic review using the ARRIVE guidelines. Journal of clinical periodontology 2012;39 Suppl 12:63-72.
32. Schwarz F, Sculean A, Engebretson SP, Becker J, Sager M. Animal models for
peri-implant mucositis and peri-implantitis. Periodontol 2000 2015;68:168-181. 33. Abrahamsson I, Berglundh T, Lindhe J. Soft tissue response to plaque formation
at different implant systems. A comparative study in the dog. Clin Oral Implants Res 1998;9:73-79.
34. Ericsson I, Berglundh T, Marinello C, Liljenberg B, Lindhe J. Long-standing
plaque and gingivitis at implants and teeth in the dog. Clin Oral Implants Res 1992;3:99-103.
35. Ericsson I, Persson LG, Berglundh T, Marinello CP, Lindhe J, Klinge B. Different
types of inflammatory reactions in peri-implant soft tissues. Journal of clinical periodontology 1995;22:255-261.
36. Zitzmann NU, Abrahamsson I, Berglundh T, Lindhe J. Soft tissue reactions to
plaque formation at implant abutments with different surface topography. An experimental study in dogs. Journal of clinical periodontology 2002;29:456-461.
37. Leonhardt A, Berglundh T, Ericsson I, Dahlen G. Putative periodontal pathogens
on titanium implants and teeth in experimental gingivitis and periodontitis in beagle dogs. Clin Oral Implants Res 1992;3:112-119.
53
38. Nociti FH, Jr., Cesco De Toledo R, Machado MA, Stefani CM, Line SR, Goncalves RB. Clinical and microbiological evaluation of ligature-induced peri-implantitis and periodontitis in dogs. Clin Oral Implants Res 2001;12:295-300.
39. Weinberg MA, Bral M. Laboratory animal models in periodontology. Journal of
clinical periodontology 1999;26:335-340. 40. Schwarz F, Sahm N, Mihatovic I, Golubovic V, Becker J. Surgical therapy of
advanced ligature-induced peri-implantitis defects: cone-beam computed tomographic and histological analysis. Journal of clinical periodontology 2011;38:939-949.
41. Pearce AI, Richards RG, Milz S, Schneider E, Pearce SG. Animal models for
implant biomaterial research in bone: a review. Eur Cell Mater 2007;13:1-10. 42. Schwarz F, Sager M, Becker J. Peri-implantitis defect model. Osteology
guidelines for oral and maxillofacial regenerations Preclinical models for translational research 2011:1-56.
43. Graves DT, Fine D, Teng YT, Van Dyke TE, Hajishengallis G. The use of rodent
models to investigate host-bacteria interactions related to periodontal diseases. Journal of clinical periodontology 2008;35:89-105.
44. Teng YT, Nguyen H, Hassanloo A, Ellen RP, Hozumi N, Gorczynski RM.
Periodontal immune responses of human lymphocytes in Actinobacillus actinomycetemcomitans-inoculated NOD/SCID mice engrafted with peripheral blood leukocytes of periodontitis patients. J Periodontal Res 1999;34:54-61.
45. Teng YT, Nguyen H, Gao X, et al. Functional human T-cell immunity and
osteoprotegerin ligand control alveolar bone destruction in periodontal infection. J Clin Invest 2000;106:R59-67.
46. Kesavalu L, Sathishkumar S, Bakthavatchalu V, et al. Rat model of polymicrobial
infection, immunity, and alveolar bone resorption in periodontal disease. Infect Immun 2007;75:1704-1712.
47. Verma RK, Rajapakse S, Meka A, et al. Porphyromonas gingivalis and
Treponema denticola Mixed Microbial Infection in a Rat Model of Periodontal Disease. Interdiscip Perspect Infect Dis 2010;2010:605125.
48. Verma RK, Bhattacharyya I, Sevilla A, et al. Virulence of major periodontal
pathogens and lack of humoral immune protection in a rat model of periodontal disease. Oral diseases 2010;16:686-695.
54
49. Nahid MA, Rivera M, Lucas A, Chan EK, Kesavalu L. Polymicrobial infection with periodontal pathogens specifically enhances microRNA miR-146a in ApoE-/- mice during experimental periodontal disease. Infect Immun 2011;79:1597-1605.
50. Rivera MF, Lee JY, Aneja M, et al. Polymicrobial infection with major periodontal
pathogens induced periodontal disease and aortic atherosclerosis in hyperlipidemic ApoE(null) mice. PLoS One 2013;8:e57178.
51. Kesavalu L, Vasudevan B, Raghu B, et al. Omega-3 fatty acid effect on alveolar
bone loss in rats. J Dent Res 2006;85:648-652. 52. Sathishkumar S, Meka A, Dawson D, et al. Extracorporeal shock wave therapy
induces alveolar bone regeneration. J Dent Res 2008;87:687-691. 53. Chukkapalli SS, Rivera-Kweh MF, Velsko IM, et al. Chronic oral infection with
major periodontal bacteria Tannerella forsythia modulates systemic atherosclerosis risk factors and inflammatory markers. Pathog Dis 2015;73.
54. Dabdoub SM, Tsigarida AA, Kumar PS. Patient-specific analysis of periodontal
and peri-implant microbiomes. J Dent Res 2013;92:168S-175S. 55. Carcuac O, Abrahamsson I, Albouy JP, Linder E, Larsson L, Berglundh T.
Experimental periodontitis and peri-implantitis in dogs. Clin Oral Implants Res 2013;24:363-371.
56. Nagasawa M, Takano R, Maeda T, Uoshima K. Observation of the bone
surrounding an overloaded implant in a novel rat model. The International journal of oral & maxillofacial implants 2013;28:109-116.
57. Zhang X, Duyck J, Vandamme K, Naert I, Carmeliet G. Ultrastructural
characterization of the implant interface response to loading. J Dent Res 2014;93:313-318.
58. Glosel B, Kuchler U, Watzek G, Gruber R. Review of dental implant rat research
models simulating osteoporosis or diabetes. The International journal of oral & maxillofacial implants 2010;25:516-524.
59. Javed F, Romanos GE. Impact of diabetes mellitus and glycemic control on the
osseointegration of dental implants: a systematic literature review. J Periodontol 2009;80:1719-1730.
60. Socransky SS, Haffajee AD. Periodontal microbial ecology. Periodontol 2000
2005;38:135-187. 61. Leonhardt A, Renvert S, Dahlen G. Microbial findings at failing implants. Clin Oral
Implants Res 1999;10:339-345.
55
62. Albouy JP, Abrahamsson I, Persson LG, Berglundh T. Spontaneous progression of peri-implantitis at different types of implants. An experimental study in dogs. I: clinical and radiographic observations. Clin Oral Implants Res 2008;19:997-1002.
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BIOGRAPHICAL SKETCH
Christie Eastman was born in Bradenton, Florida. She completed her
undergraduate degree with a Bachelor of Science in biology at Rollins College in May
2008. She continued her education at the University of Florida, College of Dentistry, and
graduated in May 2013 with a Doctor of Dental Medicine. Currently, she is enrolled in
her final semester of her post-doctoral residency in Periodontology at the University of
Florida and anticipates graduating in June 2016 with a Certificate in Periodontics and a
Master of Science. She plans to return home to her family and begin practicing clinical
periodontology in Bradenton, Florida.