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

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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

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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

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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

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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

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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

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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.