Investigating batch-to-batch variability and stability of Enamel Matrix Proteins Preparations Used in Periodontal Surgery (Emdogain®)

by

Jacob Richard Swiderski

A thesis submitted in conformity with the requirements for the degree of Master of Science Discipline of , Faculty of University of Toronto

© Copyright by Jacob Richard Swiderski 2019

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Investigating batch-to-batch variability and stability of Enamel Matrix Proteins Preparations Used in Periodontal Surgery (Emdogain®)

Jacob Richard Swiderski

Master of Science

Discipline of Periodontology, Faculty of Dentistry

University of Toronto

2019

Abstract

Emdogain®, a porcine-derived (EMD), is a promising biological mediator used in the regeneration of lost , a direct result of periodontitis. However, several clinical studies have demonstrated inconsistencies regarding Emdogain®’s therapeutic results. We hypothesize that a reason for inconsistent clinical results stems from a variability in the product’s protein content resulting from the inherent variability in the harvested enamel matrix, presence of enzymes and effects of storage conditions. Emdogain® protein composition was assessed using tandem mass spectrometry (TMS), silver stained SDS-PAGE and western blotting. Enzyme activity was examined using a fluorescence enzyme assay. TMS identified all major Enamel matrix proteins (EMPs) and enzymes. No definitive effects of temperature and time were observed through SDS-PAGE, western blotting or TMS. Nonetheless, TMS analysis did show differences in abundances of certain EMPs between batches. Enzymatic activity was detected in most samples, with significant differences in activity between batches.

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Acknowledgements

My Masters journey began with the trials and tribulations at the laboratory bench, full of questions and uncertainty. However, in the short period of three years, the mentorship, support and vision of a special group of people allowed me to complete a thesis that I am proud of. This team of peers, professors and family to whom I’m forever grateful, include the following:

To my supervisors, Dr. Bernhard Ganss and Dr. Howard Tenenbaum: A heart-felt thank you for all the time and effort spent on molding me into the graduate I am today. I was but an inexperienced student, but the mentorship and care taken into guiding me through both the laboratory work, the committee meetings, thesis writing and finally the defense itself is the reason I succeeded. But it is not just the gratitude for the end result – rather, your leadership and inspiration throughout my journey amounted to the enlightenment and enjoyment that I experienced by being under the wings of such great teachers. The humor and kindness that both of you have graced me throughout my career as a graduate student is unmatched, and a dominant reason for why I will look back at my graduate studies with a smile.

To James Holcroft: Thank you immensely for teaching me all there is to know about laboratory work. Your time and dedication in helping me accomplish all my tasks (and simultaneously preventing any chemical catastrophes from occurring) is the reason I was able to complete all my work and data collection with high quality and satisfaction. I will cherish my time in the lab thanks to you and your extensive knowledge of biochemical procedures, as well as movie quotes and comical impersonations.

To Dr. Michael Goldberg and Dr. Karina Carneiro, my advisory committee members: Thank you for supporting this project, and providing insightful guidance and valuable feedback at our meetings.

To Dr. Suzette Guo and Dr. Dianna Malkin: It was a pleasure sharing the past three years with you as co-residents. I am sure that you both will make great periodontists, and I am excited to see what the future holds for us.

To my wife-to-be (Patricia), my parents (Richard and Joanna) and my sisters (Kamila and Karolina): I am grateful that I had this rock-solid support system throughout my three years of residency and graduate studies. My journey was far from easy, and at times it was even overwhelming – but I was never alone. I had this great team that I could come back home to, knowing very well that I would be listened to, and that I'd be encouraged and motivated. It is these amazing individuals that kept me grounded and focused, and reminded me each step of the way of why I took this task upon myself. Patricia, I know you’re very happy to be this well versed in enamel matrix proteins!

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Table of Contents Acknowledgements……………………………………………………………….…….iii Table of contents………………………………………………………………...... iv List of Figures…………………………………………………………………………..vi List of Appendices……………………………………………………………………...vii List of Abbreviations…………………………………………………………………...viii 1. SYNOPSIS/DIRECTION OF RESEARCH………………………………………..1 2. BACKGROUND……………………………………………………………………..3 2.1. The Periodontium in health…………………………………………………………3 2.1.1 Structures and components of the healthy periodontium………………….3 2.1.2 Origin and development of the periodontium……………………………..7 2.1.3. Enamel Matrix Proteins and Amelogenesis………………………………8 2.1.4. EMPs and root development……………………………………………...10 2.2. The Periodontium in disease………………………………………………………..12 2.2.1. Periodontitis………………………………………………………………12 2.3. Treatment of …………………………………………………...14 2.3.1. Surgical and non-surgical periodontal treatments………………………...14 2.3.2 Regeneration……………………………………………………………….15 2.3.3. Biological mediators………………………………………………………15 2.3.4. Emdogain®………………………………………………………………..17 2.3.4.1. Cellular effects of Emdogain®………………………………….18 2.3.4.2. Emdogain® and wound healing…………………………………21 2.3.4.3. Emdogain®; Effects on Bacteria………………………………..23 2.3.4.4. Histological findings…………………………………………….23 2.3.5. Clinical application of EMD………………………………………………25 2.3.5.1. Treatment of Intraosseous defects………………………………25 2.3.5.2. Treatment of ………………………………..27 2.3.5.3. Treatment of Furcation defects………………………………….28 3. AIM, OBJECTIVES AND HYPOTHESIS………………………………………...30 3.1. Statement of the problem…………………………………………………………....30 3.2. Objectives…………………………………………………………………………...30 3.3. Hypothesis…………………………………………………………………….…….31

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4. MATERIALS AND METHODS……………………………..…………………….32 4.1. Materials…………………………………………………………………………....32 4.2. Methods…………………………………………………………………………….33 4.2.1. Sample treatment………………………………………………………....33 4.2.2. Primary antibody production (anti-porcine AMTN)……………………..33 4.2.3. Tandem Mass Spectrometry (TMS)……………………………………...34 4.2.4. SDS-PAGE……………………………………………………………….35 4.2.5. EnzChek® Protease Assay……………………………………………….37 5. RESULTS……………………...…………………………………………………….38 5.1. TMS………………………………………………………………………………...38 5.2. Silver stained SDS-PAGE………………………………………………………….40 5.3. Western Blotting…………………………………………………………………...43 5.4. Fluorescence Enzyme assay………………………………………………………..44 6. DISCUSSION………………………………………………………...……………..46 7. CONCLUSION AND FUTURE DIRECTIONS………………………………….53 Bibliography…………………………………………………………………………….55 Appendices……………………………………………………………………………...71

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List of Figures

FIGURE 1: Cross-sectional diagram of the periodontium…………..…………… 3

FIGURE 2: Gingival fiber groups of the lamina propria………………………… 5

FIGURE 3: Fiber network of the ………….………………………….. 6

FIGURE 4: Histological depiction of developing human ………….………. 7

FIGURE 5: EMP abundances from TMS analysis (run 1)……………………….. 39

FIGURE 6: EMP abundances from TMS analysis (run 2)……………………….. 40

FIGURE 7: Silver-stained SDS-PAGE lane with labeled bands………………… 41

FIGURE 8: Band intensities versus molecular weight……………………..…….. 42

FIGURE 9A: Western Blot membrane treated with anti p-AMTN………..…….. 43

FIGURE 9B: Line graph of normalized band intensities versus time……………. 43

FIGURE 10: Enzyme assay fluorescence of Emdogain® batches………….…….. 44

FIGURE 13: Porcine AMTN amino acid sequence with potential MMP 20 and KLK 4 cleavage sites…………………………...…… 49

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List of Appendices

APPENDIX A: Emdogain® batch LOT numbers and expiry dates…….………. 71

APPENDIX B: Western Blot of porcine AMTN dilution series……………...... 72

APPENDIX C: TMS reporter ion intensities for select proteins (run 1)………… 73

APPENDIX D: TMS reporter ion intensities for select proteins (run 2)………… 74

APPENDIX E: Silver-stained SDS-PAGE gels of Emdogain ®……..…………. 75

APPENDIX F: Histograms of band intensities versus molecular weight………... 80

APPENDIX G: anti-pAMTN treated Western Blots of Emdogain® …………… 84

APPENDIX H: Normalized AMTN band intensities vs time of storage…….….. 88

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List of Abbreviations

AAP American Academy of Periodontology AEP Acquired Enamel Pellicle ALP Alkaline phosphatase AMBN Ameloblastin AMEL Amelogenin AMTN Amelotin BMP Bone Morphogenic Proteins BSA Bovine Serum Albumin BSP Bone Sialoprotein CAF Coronally Advanced Flap CAL Clinical Attachment level CTG Connective Tissue Graft DFDBA Demineralized freeze-dried bone allograft DEJ Dental Enamel Junction EBL External Basal Lamina EMD Enamel Matrix Derivative EMP Enamel Matrix Proteins ENAM Enamelin FDBA Freeze-dried bone allograft FGF Fibroblast Growth Factor GTR Guided Tissue Regeneration HA Hydroxyapatite HERS Hertwig’s Root Sheath HMVEC Human Microvascular Endothelial Cells IBL Internal Basal Lamina IC Intermediate Cementum IGF Insulin Growth Factor IL Interleukin

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JE Junctional KLK Kallikrein-related Peptidase LRAP Leucine-rich Amelogenin Peptide MCP Monocyte Chemo-attractant MGJ MIST Minimally Invasive Surgical Techniques MMP Matrix Metalloproteinase OC Osteocalcin ODAM Odontogenic Ameloblast-Associated OE Oral Epithelium OFD Open Flap OPG Osteoprotegerin OPN Osteopontin PD Probing Depth PDGF Platelet-derived Growth Factor PDL Periodontal ligament PDVF Polyvinylidene Difluoride PGA Propylene Glycolic Alginate

RANK Receptor activator of nuclear factor kappa-Β

RANKL RANK Ligand SDS-PAGE Sodium Dodecyl-sulfate Polyacrylamide Gel Electrophoresis SE TBST Tris-Buffer Saline Tween TGF Transforming Growth Factor TIMP Tissue inhibitor of metalloproteinases TMS Tandem Mass Spectrometry TNF Tissue Necrosis Factor TRAP Tyrosine-rich amelogenin peptide VEGF Vascular Endothelial Growth Factor

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1. SYNOPSIS/ DIRECTION OF RESEARCH

Emdogain® is a commercially available enamel matrix derivative (EMD), composed of a purified extract from the embryonic enamel of unerupted molars in six-month old piglets within a propylene-glycolic-alginate carrier (Maycock et al., 2002). Its main indication is in periodontal tissue regeneration. Enamel matrix proteins (EMPs) are thought to be the active agents in Emdogain®. Their individual characteristics and roles in the developing enamel during amelogenesis are still under investigation and their role or mechanism of action at the level of periodontal regeneration is even more obscure. It has been suggested that EMPs are secreted during periodontal development by cells of Hertwig’s root sheath (HERS), and play a role in the formation of the acellular cementum (Hammarstrom, 1997). Studies have shown that EMD, a complex mixture of enamel (and other) proteins, is capable of promoting early wound healing and resolving inflammation through reduction of interleukin expression and production, which may reduce local bone resorption and favor the formation of new bone (Miron et al., 2015). It has even been used as an aid for enhancement of the effects of non-surgical management on peri- implant mucosal disease (Kashefimehr et al., 2017). Moreover, it can enhance the maturation phase of wound healing and initiate periodontal regeneration by directly stimulating gingival fibroblast, osteoblast and cementoblast proliferation and activity (Zeldich et al., 2007; Weinberg et al., 2010).

Clinically, addition of EMD in the treatment of intra-bony defects, furcation involvements and some root coverage procedures has yielded superior results when compared to more classical surgical approaches (Heijl et al., 1997; McGuire and Nunn, 2003). However, root coverage procedures where a connective tissue graft was used resulted in further increased root coverage beyond that of EMD alone (McGuire and Nunn, 2003). With regard to periodontal regeneration, there was no clinically significant difference between either guided tissue regeneration (GTR) or EMD for treatment of intra-bony defects (Pontoriero et al., 1999). Although histological analyses have shown that EMD can induce periodontal regeneration, it appears that GTR was more consistent in this regard (Sculean et al., 1999).

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In addition to a wide array of things that might influence outcome of treatment (ranging from the type of defect being treated to the skills of the practitioner), it can also be suggested that some possible factors contributing to inconsistent clinical results found with the use of Emdogain® might include potential variations in the content of proteins contained in this agent on a batch-to-batch basis. It is also conceivable, given that proteins are the proposed biologically active agents, that storage conditions might affect Emdogain® content composition, potentially altering clinical outcomes. The assembly and the structure of the proteins contained within Emdogain®, and thus their biological activity, may be sensitive to various factors, including temperature. The therapeutic effectiveness relies in part on the native conformation of its protein constituents.

The inherent variability in protein composition of the enamel matrix during development can be an obstacle in obtaining samples with a consistent protein composition. At a single point in time, different sections of the same will be at different phases of amelogenesis, and thus there will be a heterogeneous mixture of protein content harvested at any given time. To confound the issue, the inherent presence of proteinases in the enamel matrix brings forth the possibility of time-dependent degradation of certain proteins within the product during storage, which could directly affect the clinical efficacy of Emdogain®. The aim of this Master’s project is to examine the protein composition of Emdogain®, assess for differences in protein and enzyme abundances amongst different batches of the product, and to discern if the components are sensitive to changes in storage temperature and time. To the author’s knowledge, the lack of published data in this regard would indicate that this would be the first public analytical investigation into this product’s compositional variability and the possible effect of the temperature and time of storage on protein composition. This investigation may unravel previously unappreciated compositional variabilities in Emdogain® and lead to rational recommendations for optimal storage conditions that would benefit the shelf life of Emdogain®. Beyond this investigation, the research project may form the basis for correlating compositional variations between batches of Emdogain® with variations in clinical outcomes.

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2. BACKGROUND

2.2. The Periodontium in health

2.1.1 Structures and components of the healthy periodontium

The supporting structures of the dentition are collectively termed the periodontium. Its main functions include support and attachment of teeth to the jaw. The periodontium is composed of the epithelium and the associated connective tissue of the gingiva, the alveolar bone, cementum and periodontal ligament fibers (Lang and Lindhe, 2015).

Figure 1: Cross-sectional diagram of the periodontium. OE: oral epithelium, SE: sulcular epithelium, JE: , CEJ: Cemento-enamel junction (source: Lang and Lindhe, 2015).

The gingiva is part of the masticatory mucosa, and covers the and surrounds the cervical portion of teeth (Lang and Lindhe, 2015). It is composed of an epithelial layer and an underlying connective tissue portion, termed the lamina propria. The epithelial portion shows regional morphologic variations, which result from tissue adaptation to the tooth and alveolar bone. These variations include the oral epithelium (OE), sulcular epithelium (SE), and junctional epithelium (JE). OE and SE are largely

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protective in function, however, the JE serves both to facilitate gingival attachment to the tooth, and also as a barrier (e.g. against oral microbes), and is of considerable importance in regulating tissue health (Shimono et al., 2003). The OE is composed of stratified squamous keratinized cells, is located on the exterior of the periodontal apparatus and is exposed to the oral cavity. The OE extends from the mucogingival junction (MGJ) apically, to the free coronally. Located interiorly, past the free gingival margin, the SE is a stratified squamous non-keratinized tissue that lines the inside of the (Schroeder and Listgarten, 1997). The base of sulcus and the location of the epithelial attachment to the mineralized surface of the tooth is a non-keratinized bilayer of epithelial cells known as the JE (Schroeder and Listgarten, 1971). The JE bilayer is attached to the underlying connective tissue via the external basal lamina (EBL) on one side, and to the mineralized tooth surface via the internal basal lamina (IBL) on the other side (Schroeder, 1963). Cells at both the EBL and IBL use a similar cellular attachment apparatus, consisting of hemidesmosomes bound to integrins to attach to components of the basal lamina (Pakkala et al., 2002). Though the EBL is a typical basement membrane containing type IV collagen (Oksonen et al., 2001), the IBL at the epithelium/tooth is also termed the dental cuticle, but, unlike the EBL, little is known of its molecular composition (Bosshardt and Lang, 2005).

The underlying connective tissue or lamina propria, is the predominant component of the gingiva and is composed of collagen fibers (60% of connective tissue volume), cells (fibroblasts, 5% of connective tissue volume) and ground substance with associated nerves and vessels (35% volume of connective tissue) (Palmer and Lubbock, 1995). Though some of the collagen fibers appear to be haphazardly arranged, most are organized into five principal fiber groups based on their orientation: dentogingival, alveologingival, dentoperiosteal, circular and transseptal. In addition, there are six additional minor fiber groups woven in between (Hassel, 1993).

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Figure 2 Cross-sectional diagram depicting orientation and relationship of some of the gingival fiber groups within the gingival lamina propria (Wolters Kluwer. Lippincott Williams & Wilkins, 2013).

The lamina propria serves to support the overlying epithelium, both through supply of nutrients and physical attachment to underlying structures (Lang and Lindhe, 2015). The fibers serve to bind down the soft tissue to the underlying hard tissue of the alveolar bone and mineralized tooth structure. The vessels of the connective tissue also provide access to immune cells both for immune surveillance and in inflammatory processes (Palmer and Lubbock, 1995).

Cementum is a mineralized tissue that covers the surface of root dentin. Like bone, it contains a collagen fiber network embedded in an organic matrix. Cementum contains about 65% hydroxyapatite mineral by weight, slightly more than bone (Selvig, 1965). Unlike bone, cementum contains no vasculature or nerves and does not undergo physiologic remodeling. Instead, it is characterized by continuous deposition throughout life (Zander and Hurzeler, 1958). Though the tissue is quite thin and minute in amount, cementum has a very specialized function. It is less prone to resorption and aids in shielding the more vulnerable root dentin from resorption by odontoclasts (Selvig, 1965). It also possesses regenerative and growth potential (Zander and Hurzeler, 1958). In terms of the attachment apparatus of the tooth, it is the site of anchorage for principle fibers of

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the periodontal ligament (Selvig, 1965).

Periodontal ligament fiber-endings embedded in the mineralized tissue of cementum or the alveolar bone on either side are termed Sharpey’s fibers (Palmer and Lubbock, 1995). The periodontal ligament is a soft and richly vascular and cellular connective tissue that encompasses the roots of teeth. Forces from masticatory function can be distributed and dissipated by the alveolar bone via the periodontal ligaments, furthermore, these ligaments allow for and some sensory function, such as proprioception (Beertsen et al., 1997). These fibers, which run perpendicular to the root surface, are produced by fibroblasts and constitute the extrinsic fiber system. The intrinsic fiber system, however, is synthesized by cementoblasts and its fibers are generally oriented parallel to the long axis of the root (Selvig, 1965).

Figure 3: Orientation and relationship of the intrinsic and extrinsic fiber network associated with cementum and the periodontal ligaments. (source: https://www.tankonyvtar.hu/en/tartalom/tamop412A/2011-0095_fogaszat_angol/ch01s09.html)

Different types of cementum have been classified based on presence or absence of cells, as well as the origin and direction of collagenous fibers. The cervical third to two-thirds of human tooth roots are covered by acellular extrinsic fiber cementum, characterized by densely packed Sharpey’s fibers in a non-cellular ground substance (Hammarstrom, 1997). Cementum with neither cells nor fibers is termed acellular afibrillar and is typically found in the coronal-most portion of roots, but can also be part of the acellular extrinsic fiber cementum. At the apical-most third of roots we find cellular mixed

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stratified cementum, which is composed both of intrinsic and extrinsic fibers and irregularly distributed cells (Hammarstrom, 1997).

2.1.2. Origin and development of the periodontium

The development of the periodontium is concomitant to the development of the crown and root of the tooth. During embryonic development, migrating neural crest cells form a band of ectomesenchyme beneath the epithelium of the primitive oral cavity, or stomatodeum (Lang and Lindhe, 2015). This ectomesenchyme receives instructive signals secreted by the epithelium of the stomatodeum, forming the dental lamina – at this stage, tooth development proceeds through the bud stage, cap stage, bell stage and root development (Lang and Lindhe, 2015). The development of the periodontium involves interplay between the mesenchymal tissue of the dental sac and the epithelium cells of Hertwig’s epithelial root sheath (HERS). The mesenchyme of the dental sac and the pulp is derived from the cranial neural crest ectomesenchyme (Thomas, 1995). During tooth root development, the mesenchyme in the apical portion of the tooth germ proliferates. This proliferation will generate lineages that contribute to radicular pulp as well as the surrounding periodontium. Concomitantly, HERS grows downward, establishing a boundary between the pulp and surrounding periodontium (Thomas, 1995).

Figure 4: Diagram depicting a histological section through the developing human tooth, late bell-stage. OEE: Outer enamel epithelium, SR: Stellate reticulum, SI: Stratum intermedium, IEE: Inner enamel epithelium, CL: Cervical loop, EO: enamel organ, DP: dental papilla, DS: Dental sack, A: ameloblasts, EM: enamel matrix, D: dentin, OB: odontoblasts, UMC: undifferentiated mesenchymal cells (source: http://mandanadonoghue.blogspot.com/2015/02/oral-histology-diagrams-advanced-bell.html)

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Root formation involves the HERS-induced formation of root dentin by the mesenchymal cells of the dental papilla. Once the mantle pre-dentin is formed, the HERS disintegrates, exposing the root surface to the surrounding mesenchymal cells of the dental follicle, thus inducing cementogenesis (Hammarstrom, 1997). However, unlike in enamel and dentin, where the deposition of protein matrices and minerals are accomplished by an adjacent well-organized layer of cells, both the source of cells responsible for cementum development, as well as the finer details of the process itself are more enigmatic (Diekwisch and Thomas, 2001). Much of the uncertainty lies in the fact that both the HERS epithelial cells and the mesenchymal cells of the dental follicle are in proximity to the developing root surface (Diekwisch and Thomas, 2001). The classical theory of cementogenesis involves the mesenchymal cells of the dental follicle invading through the disintegrating HERS. These cells are believed to differentiate into cementoblasts and secrete cementum (Paynter and Pudy, 1958; Schroder, 1986). Based on microscopic observations of animal cementum as well as immunological similarities between enamel and cementum proteins, a crucial role of the epithelial lineage in cementogenesis has been proposed (Stahl and Slavkin, 1972; Slavkin and Boyde, 1974; Slavkin et al., 1989). Scanning electron microscope and auto-radiographic studies on monkey incisors have observed a secretory stage within the inner epithelial root sheath and the formation of an enamel-like secretion on root dentin prior to cementum formation (Lindskog 1982). Furthermore, an intimate relationship between coronal cementum and enamel exists in many mammals including humans and manifests as a thin superficial layer of acelluar cementum on the most cervical enamel (Listgarten 1967). The exposure of the developing enamel to the cells of the dental follicle seems to initiate the process of coronal cementogenesis and starts shortly after the completion of the secretory stage of amelogenesis (Hammarstrom 1997).

2.1.3. Enamel Matrix Proteins and Amelogenesis

As a prerequisite for enamel formation, odontoblasts begin secreting a pre-dentin matrix, which begins to mineralize from the outside (nearest to the future dentin-enamel junction, or DEJ ) and progresses inwards towards the pulp (Mjor and Ole, 1986). As a response to

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this initial dentin mineralization near the DEJ, differentiating pre-ameloblasts extend cytoplasmic projections through the interfacial basement membrane, removing it (Bartlett, 2013). Amelogenesis is composed of two main stages: secretory and maturation. During the former, EMPs are secreted by ameloblasts and are essential for enamel growth. During the maturation stage, organic content is removed from the enamel matrix and mineral content increases to ultimately reach over 96% hydroxyapatite. The secreted EMPs include the structural proteins of the amelogenin family (>90%), ameloblastin (<5%) and enamelin (<5%), as well as trace amounts of two proteolytic enzymes, matrix metalloproteinase -20 (MMP 20) that is predominantly expressed during the secretory stage, and Kallikrein-4 (KLK 4) that is predominantly expressed during the maturation stage (Bartlett, 2013).

Developmentally, amelogenesis is orchestrated by the enamel organ, which is composed of an outer epithelial layer, the stellate reticulum, the stratum intermedium, and the inner enamel epithelium. The inner enamel epithelium is located at the future DEJ and is composed of a single layer of ameloblasts, which are attached via hemi-desmosomes to the underlying basal lamina (Bartlett, 2013). The secretory stage begins with the transformation of pre-ameloblasts to secretory ameloblasts, morphologically distinguished as elongated columnar cells with Tomes’ processes at their apical extent. Ameloblasts secrete large amounts of EMPs as they move away from the dentin surface, and these EMPs induce the formation of parallel crystallite ribbons. During the secretory stage, MMP-20 activity leads to limited hydrolysis of EMPs. An ameloblast is responsible for creating a single rod or prism, which is composed of about 10 to 40 thousand individual crystal-forming ribbons (Daculsi et al., 1984). An interdigitated network of such prisms, each derived from a single ameloblast, forms the bulk of the enamel. By the end of the secretory stage, the enamel layer has achieved its full thickness (Bartlett, 2013).

As the process transitions from the secretory to the maturation phase, the ameloblasts lose their Tomes’ processes, and this occurs simultaneously with the formation of a smooth enamel surface with a final coating of densely mineralized, aprismatic enamel (Barlett, 2013). Morphologically, maturation stage ameloblasts appear non-polarized and more

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cuboidal, display the hallmarks of transport cells, and re-establish a new basal lamina-like layer at the interface with the enamel surface (Smith, 1998). A second proteinase, KLK- 4, is secreted mainly during this stage to completely degrade the previously secreted EMPs and their larger cleavage products, which are then removed from the enamel layer (Hu et al., 2000). This allows the rod and inter-rod crystallites to expand in volume, eventually interlocking and almost completely filling the enamel space (Bartlett, 2013).

Unlike the EMPs described above, Odontogenic ameloblast-associated (ODAM, formerly called Apin) and Amelotin (AMTN) are expressed during the maturation phase stage of enamel formation (Nakayama et al., 2005; Moffatt et al., 2008) and in the JE after (Nishio et al., 2010). In terms of amelogenesis, current studies have localized AMTN predominantly at the enamel/ameloblast interface of rodent incisors of rats (Moffatt et al., 2006; Somogyi-Ganss et al., 2011). It has been proposed from studies in transgenic mice that AMTN, possibly in conjunction with ODAM, is involved in the formation of the surface enamel, whereby there is a structural transition from bulk to the more compact superficial layer (Lacruz et al., 2012). Furthermore, both AMTN and ODAM play a key role in the attachment mechanism of the JE to the mineralized tooth surface (Moffatt et al., 2006; 2008), especially because AMTN and ODAM have been shown to interact with one another (Holcroft and Ganss, 2011). Thus, all these organic components of the enamel matrix play specific individual roles during amelogenesis. Overall, the molecular mechanisms that govern amelogenesis are remarkably conserved in different mammalian species (Robinson et al., 1988)

2.1.4 EMPs and root development

The role or even existence of EMPs in the vicinity of cementum has been historically debated. Ultra-structural findings in root development indicate both synthetic and secretory activity of the HERS (Slavkin et al. 1988). EMP expression has been observed on the developing root sheath (Slavkin et al., 1989; Fong et al., 1996; and Thomas et al., 1997), though it has been argued that HERS cells produce distinct cementum proteins (Bosshardt and Nanci 1997, 1998, 2000).

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Immunohistological studies done in animals have shown that the concentration of amelogenin rises with tooth development (Hammarstrom, 1997). Histologically, human teeth have a thin layer of highly mineralized enamel existing between the root dentin and the acellular cementum (Hammarstrom, 1997). Indeed, laminin and other epithelial- derived proteins are present on the developing root surface. Since epithelium products are well-known to induce bone formation in susceptible mesenchyme (Hall and Van Exan, 1982), it is reasonable to speculate that they may have a similar effect on the root dentin. Osteopontin and bone sialoprotein have been shown to be associated with the early stages of acellular cementogenesis, potentially underscoring the potential role of those two proteins in cell adhesion to the peripheral root surface (Thomas, 1995). However, some studies have shown an absence of amelogenin mRNA in HERS cells (Lou et al., 1991; Diekwisch, 2001), stating the possibility of immunological cross-reactivity of polyclonal amelogenin antibodies with keratin as the reason for the conflicting findings. Interestingly, through in situ hybridization, those same studies discovered low-level amelogenin hybridization signals in the odontoblast layer (Diekwisch, 2001). This supports previous findings of polypeptides containing an amino-terminal amelogenin fragment in dentin matrix extracts (Nebgen et al., 1999). Another EMP, enamelin, has also been detected in cells embedded in cementum (Fong et al., 1996).

The following may help explain the conflicting results with regard to the presence or absence of EMPs in cementogenesis: HERS cells synthesize and secrete enamel-related proteins along the forming root surface of mouse molars during in situ as well as long- term in in vitro organ culture (Slavkin et al., 1988). These proteins, though sharing certain similar domains with enamelin and amelogenin, do possess distinct amino acid sequences. In developing roots of mouse molars, cross-reactivity existed with both anti- mouse amelogenin and anti-mouse enamelin IgG antibodies. Several root-derived proteins were identified, including 72 kDa and 26kDa root-derived intermediate cementum (IC) proteins, which were identical with one another but distinct from either coronal enamelin or amelogenin (Slavkin et al., 1988). It was noted that these IC proteins contained a significantly higher amount of certain amino acids, especially valine, relative to those found in coronal EMPs. It was suggested that the observed cross-reactivity may

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be the result of selective overexpression of particular protein splice variants that are particularly rich in conserved peptide domains during amelogenesis as well as initial cementogenesis, resulting in the observed immuno-cross reactivity (Slavkin et al., 1988).

2.2. The Periodontium in disease

2.2.1. Periodontitis

The contains a very unique feature – the non-shedding surface of teeth. These surfaces are exposed to a plethora of elements and microbiota. Unlike with all other human surfaces where the constant sloughing of the exterior surface layer prevents or limits accumulation of microbes, the accumulation and metabolism of bacteria on these hard surfaces is considered the primary cause of dental caries, , periodontitis, peri-implant infections, and stomatitis (Lang and Lindhe, 2015).

Periodontitis is a bacteria-associated and host modulated inflammatory disease that affects the supporting structures of teeth. The bacterial biofilm initiates a host- inflammatory response, which can lead to gingival inflammation, known as gingivitis. Not all cases of gingivitis become periodontitis, but gingivitis is a prerequisite to establish the progression to periodontitis (Baelum et al., 1988). The interplay between the microbial challenge and the host immune-inflammatory response is in part responsible for the advancement to periodontal disease (Page, 1998). The resulting destruction of connective tissue and deregulated bone metabolism, the hallmarks of periodontitis, are further influenced by genetic, environmental and acquired risk factors. Though bacterial biofilms initiate tissue destruction, risk factors, such as smoking (Bergstrom and Preber, 1994; Tomar and Asma, 2000; Johnson, 2004), diabetes status (Grossi and Genco, 1998; Taylor and Borgnakke, 2008), and genetic background of affected individuals (Kornman et al., 1997; Papapanou, 1999; Diehl et al., 1999), indicate that the etiology of periodontitis is multi-factorial.

The 2017 World Workshop differentiated 3 forms of periodontitis based on distinct pathophysiology: Necrotizing periodontitis; Periodontitis as a direct manifestation of systemic diseases; and Periodontitis (Caton et al., 2018). Differential diagnosis of these

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three entities is focused on the history and specific signs and symptoms of necrotizing periodontitis or the presence of an uncommon systemic disease that alters the host immune system. However, the majority of clinical cases fall under the category of periodontitis (Tonetti et al., 2018). The pathophysiology of periodontitis is focused on the dysbiotic ecological changes in the microbiome as a response to gingival inflammatory proteins and tissue breakdown products (Tonetti et al., 2018). The influx of inflammatory proteins and tissue breakdown by-products provides vital nutrients and alters the micro- environment of the periodontal tissues allowing for these dysbiotic changes. These biofilm changes incite the host response, which is a crucial culprit of the consequences observed in periodontal disease.

Inflammatory cell recruitment and proliferation, as well as molecular signaling cascades, lead to the release of anti-bacterial products and host-tissue proteinases. Specifically, immune cells such as monocytes and macrophages respond to the presence of bacterial virulence factors with the release of pro-inflammatory cytokines, including interleukin-1 (IL-1) and tumor necrosis factor-alpha (TNF-α) (Kornman et al., 1997). These induce production of matrix metalloproteinases, which destroy the connective tissues of the gingiva and periodontal ligament, and prostaglandins, which mediate alveolar bone destruction (Page, 1998). Therefore, the unfortunate by-product of this immunological response is destruction of the surrounding periodontal tissue, and hence attachment loss around the tooth. Additionally, most periodontal pathogens secrete their own extracellular proteinases, which contribute to the pathogenesis of the disease. For example, , a gram negative anaerobic rod with significant ties to periodontal disease, secretes a class of enzymes termed gingipains. These enzymes not only degrade host proteins such as collagen, but also breakdown certain antibodies and pro-inflammatory cytokines, impairing the host immune response (Sheet et al., 2012). The apical migration of the junctional epithelium, resulting from the loss of marginal periodontal ligament fibers, allows for the apical migration of the bacterial biofilm. This in turn helps in the microbial evasion of mechanical debridement efforts, contributing to the chronicity of the inflammatory state (Page, 1988).

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Over time, the gradually-decreasing support leads to the premature loss of teeth, resulting in partial and even complete edentulism. As such, periodontitis has a profoundly negative effect on the quality of life and general health. The prevalence of periodontitis in the US population is approximately 47% in adults over the age of 30 years, and over 70% in adults over the age of 65 (Eke et al., 2012). The most severe forms of periodontitis affect approximately 10% of surveyed populations (Norderyd et al., 2015; Eke et al., 2012). The average annual attachment loss in the general population (both healthy and with periodontitis) is about 0.1mm (Needleman et al. 2017). However, this number drastically increases to an annual loss of 0.57mm in the subgroup of patients with periodontitis (Needleman et al. 2017).

2.3. Treatment of periodontal disease

2.3.1. Surgical and non-surgical periodontal treatments

Periodontal therapies are designed to arrest the progression of attachment loss, with subsequent treatments being implemented to remove destroyed periodontal tissues and to stimulate regeneration. Retention of dentition in function and comfort has been the classical goal of periodontal therapies. Both non-surgical and surgical therapies rely primarily on the eradication of microbial deposits (Pihlstrom et al., 1983). Non-surgical treatment includes both scaling and root planning with or without adjunctive pharmacological treatment (usually systemic or locally delivered antibiotics), as well as instructions to motivate and educate patients on proper home-care (Axelsson and Lindhe, 1981; Pihlstrom et al., 1983). Classically, surgical treatment is initiated when periodontal pocketing deepens to a level where non-surgical therapy becomes either inefficient or inaccessible. Indeed, it has been shown that the extent of residual sub- gingival (after non-surgical debridement) was directly related to pocket depth, and a significantly greater proportion of deposits were left following non-surgical therapy (Caffesse et al., 1986).

Surgical treatment, such as (OFD) provides critical access to evaluate and decontaminate root surfaces and establishes favorable architecture of the

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hard and soft tissue for long-term maintenance (Cortellini et al., 2007). Unfortunately, if periodontal defects are left empty after surgical debridement, they tend to become repopulated with fast growing and proliferating epithelial cells and fibroblasts, which generate a fibro-epithelial tissue attachment to the root surface (Trombelli, 2005). This tissue repair process does not allow time for cells of the natural supportive tissues (bone and PDL) to proliferate into the defect, and as such the original architecture and tissue organization is lost.

2.3.2. Regeneration

Regeneration of lost attachment is the most sought-after periodontal treatment outcome, however it is very case-specific and presently unpredictable. The American Academy of Periodontology (AAP) has defined periodontal regeneration as the restoration of lost periodontium, the formation of new bone, new cementum, and functionally oriented periodontal ligaments. To date, most methods of regeneration involve the concept of “Guided tissue Regeneration” (GTR), whereby certain cell populations are excluded from a defect in order to allow for colonization of wounds coronal to the alveolar crest by cells derived from periodontal ligament and bone (Melcher, 1976). Exclusion is usually performed through the use of either a resorbable (e.g. xenogenic collagen) or non- resorbable (e.g. poly-tetraflouroethylene) membrane. The ultimate goal is the reproduction or reconstitution of the lost part with the same tissue and architecture. Different materials have been introduced and used to promote regeneration and maintain the defect volume: different collagen and synthetic membranes; bone replacement fillers, including autografts, allografts, xenografts and synthetic minerals; and more recently biological mediators. The latter category includes extracellular matrix proteins, cell attachment factors, growth factors and differentiation factors (Bosshardt, 2008).

2.3.3. Biological mediators

The most studied class of biological mediators for regeneration appears to be growth factors. These molecules signal and regulate a multitude of cellular functions, including chemotaxis, differentiation and proliferation (Bosshardt, 2008). Examples include

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insulin-like growth factors (IGFs), fibroblast growth factors (FGFs), transforming growth factor-b (TGF-β), and bone morphogenic proteins (BMPs). The most promising of these are BMPs, which are part of TGF-β superfamily and help regulate bone formation and growth (Subramaniam et al., 2013).

In particular, BMP-2 and BMP-7 had demonstrated promising results in pre-clinical animal studies involving periodontal and furcation defects (Ripamonti et al., 1994; Sigurdsson et al., 1996; Cochran and Wozney, 1999). Though these animal studies demonstrated significant reconstitution of bone, cementum and periodontal ligament tissue in tested defects, the results were not entirely ideal. Given the unique action of BMPs on mineralized tissue formation, obliteration of periodontal ligament space and ankylosis were observed in a number of test subjects (Wikesjö et al., 1999). It should be noted further that there have been far fewer human clinical trials with BMPs for periodontal regeneration, as opposed to other areas of medicine such as in orthopaedics. In part, this can be justified by the fact that the periodontium is not comprised solely of bone alone. Rather, this structure can be regarded more as an ‘organ’ or at least a complex tissue structure containing various unique elements. The heterogeneity of the tissues that constitute the periodontium complicates attempts at its regeneration and could indicate that a variety of factors might be needed to induce regeneration, ranging from physical graft replacements to biological factors (like Emdogain®). Furthermore, even with knowledge of putative biological agents that could induce some aspects of regeneration, dosing issues would still need to be addressed (Bosshardt, 2008). Moreover, growth factors such as BMPs have a high clearance rate, and thus ideal carriers and release systems might need to be investigated to prolong the resident time of these molecules (Caffesse and Quinones, 1993) or perhaps even to make sure that these biomolecules do not persist to the point of causing detriment. These issues reflect the notion that given the putative need for multiple growth factors that could be needed to induce periodontal regeneration, different release kinetics need to be considered to optimize for the differential growth rate of the periodontal tissues, thus optimizing regeneration (Bosshardt, 2008).

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2.3.4. Emdogain ®

Clearly there are several agents and factors that could be analyzed in relation to their biocomposition and relevance to periodontal therapy. In this case the focus of the investigations reported in this thesis will be on Emdogain® (Straumann, Switzerland), a purified fraction of EMPs derived from the embryonal enamel layer of molars extracted from 6-month old piglets in a propylene-glycolic-alginate (PGA) carrier (Maycock et al., 2002). It has recently been given the working name “enamel matrix derivative” (EMD) (Miron et al. 2016). Very little detail regarding procedural steps, reagents, specific tooth type sources (if any) is available publicly due to the proprietary nature of EMD/Emdogain®. The presence of EMPs on the surface of developing tooth roots prior to formation of cementum and their potential role in cementogenesis (Lindskog 1981a, b, Lindskog & Hammarstrom 1981, Slavkin et al. 1989) has stimulated over two decades of subsequent research focused on the putative role of EMPs and their possible use for stimulation of periodontal regeneration (Hammarstrom et al. 1991, 1992, 1995).

The major components of EMD are amelogenins (AMEL), a family of hydrophobic proteins that account for more than 90% of the total protein content derived from different splice variants and post-secretory regulation, all controlled from the expression of a single gene (Lyngstadaas et al., 2009). The remaining 10% are proline-rich non- amelogenins, tuftelins and other serum proteins (Brookes et al., 1995). More specifically, the remaining protein content is composed of enamelin (ENAM), ameloblastin (AMBN), amelotin (ATMN), apin (ODAM) and certain proteinases, namely MMP-20 (Fukae et al., 1998) and KLK-4 (Simmer et al., 1998). Many of the enamel matrix proteins have been shown to auto-assemble into a large insoluble extracellular matrix that controls the structural organization of the developing enamel crystallites (Bartlett et al., 1996). While much has been learned about the roles of individual enamel proteins through transgenic animal studies, many fundamental questions remain unanswered. For example, the activity of peptide fragments of AMEL, AMBN, ENAM, produced by limited hydrolysis via MMP-20, remains largely enigmatic. Thus, although there is already a font of information regarding the individual components of EMD, the roles that they play within

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the context of Emdogain® are still unclear. However, there have been several investigations that have demonstrated that individual fractions of EMD are responsible for different cellular and tissue effects. If the levels/concentrations of the individual protein components vary, this could account for the variable clinical results that have been observed when EMD is used therapeutically (Miron et al., 2016).

2.3.4.1 Cellular effects of Emdogain®

EMD can promote epithelial cell adhesion as well as stimulates the polymerization of cytoskeletal actin (Kawase et al. 2001; Rincon et al. 2005). However, EMD was shown to inhibit epithelial cell proliferation. It was thus concluded that EMD has a cytostatic effect on epithelial cells (Kawase et al. 2000). Furthermore, EMD reduces epithelial cell DNA-synthesis (Kawase et al. 2002). However, EMD does not have the same effect on all epithelial cell lines. DNA synthesis by the epithelial cell rests of Malassez was increased significantly after EMD stimulation (Rincon et al., 2005). Of interest is the fact that this line of cells is known to respond to inflammatory mediators by proliferation and may be involved in periodontal regeneration (Rincon et al., 2005). In terms of expression of extracellular proteins, EMD treatment of human epithelial cells caused significantly increased secretion of platelet-derived growth factor (PDGF) compared to baseline levels. Similarly, treatment of epithelial cells derived from the epithelial cell rests of Malassez with EMD induced increased synthesis of osteopontin (OPN) (Lyngstadaas et al. 2001; Rincon et al., 2005).

EMD stimulates proliferation of human gingival fibroblasts in a dose-dependent manner as well as increases DNA synthesis (Kawase et al. 2002). EMD also stimulates the attachment and spreading of human gingival fibroblasts in culture. Interestingly this effect was more pronounced with PDL fibroblasts compared to gingival fibroblasts. Further, it was found that integrins play a key role in the interaction between EMD and fibroblasts (Van der Pauw et al. 2000; 2002). In terms of expression and synthesis of EMPs, EMD was shown to stimulate the release of TGF-β1 by human gingival fibroblasts (Van der Pauw et al. 2000). EMD also stimulates synthesis of both hyaluronan

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and OPN by human gingival fibroblasts (Haase and Bartold, 2001). Moreover, EMD stimulated the levels of alkaline phosphatase in human gingival fibroblasts significantly as compared to baseline – however, those levels were markedly lower as compared with human PDL fibroblasts (Van der Pauw et al. 2000)

EMD has been demonstrated to increase human PDL cell density and DNA synthesis in cultures (Lyngstadaas et al. 2001; Okubo et al. 2003; Davenport et al. 2003). PDL cell attachment rate was significantly increased when these cells were treated with EMD in culture (Lyngstadaas et al. 2001). Futhermore, in an in vitro model for cell attachment, EMD induced the synthesis of the cell surface receptor integrin ανβ3 by human PDL fibroblasts, thereby enhancing cell attachment (Suzuki et al. 2001). EMD has been shown to stimulate synthesis of total protein by human PDL fibroblasts (Gestrelius et al. 1997). EMD also induces increased production of autocrine/paracrine factors such as TGF-β1, interleukin 6 and PDGF-AB by PDL cells (Lyngstadaas et al. 2001). Moreover, human PDL fibroblasts responded to EMD by significantly increasing (in a dose-dependent manner) proteoglycan and hyaluronan synthesis (Haase and Bartold, 2001). EMD increases both BSP and OPN mRNA expression in Human PDL cells, whereas osteocalcin (OC) mRNA expression is decreased (Hakki et al. 2001). Moreover EMD stimulates both gene expression for and actual synthesis of IGF-1 and TGF-β1 proteins in PDL fibroblasts (Okubo et al. 2003). EMD significantly increased alkaline phosphatase (ALP) activity and increased in vitro mineralized nodule formation in human PDL fibroblasts (Rodrigues et al. 2007).

Human dental follicle cells treated with EMD in vitro had an increased expression of BMP-2, BMP-7, BSP, cementum attachment protein (CAP) and cementum protein-23 (CP-23), two putative cementum markers (Kemoun et al., 2007). Exposure of immortalized murine cementoblasts to EMD resulted in significantly enhanced cell proliferation (Tokiyasu et al. 2000, Viswanathan et al. 2003). EMD down-regulated OC and slightly up-regulated OPN mRNA expression, whereas gene expression for BSP was only slightly increased towards the end of the cultivation period of murine cementoblasts (Tokiyasu et al. 2000). In an in vitro experiment using porcine stromal cells placed on teeth treated with EMPs (porcine tooth germ origin), histology elucidated that a new

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cellular cementum-like tissue formed along EMP-treated root surfaces (Bosshardt 2005).

EMD increases the proliferation of human bone marrow stromal cells in a dose- dependent manner (Guida et al., 2007). EMD treatment of human osteoblastic cells (Saos-2) increased cell proliferation (Heng et al. 2007). Similarly, when mouse pre- osteoblastic cell line (MC3T3-E1) were treated with EMD, there was a significantly enhanced ALP activity, and an up-regulation of mRNA expression of type I collagen, BSP, OC, osteoprotegrin (OPG), and IGF-1 (He et al., 2004a). The fractioned enamel matrix extract from developing porcine teeth demonstrated an osteo-inductive fraction containing mainly 20- 23kDa proteins (Iwata et al., 2002). This fraction enhanced ALP activity and in vitro mineralized nodule formation, and up regulated OC, BSP, and ALP mRNA expression in ST2 cells, a mouse bone marrow stromal cell line. With respect to the RANKL/OPG system of bone homeostasis, EMPs have been shown to have an influence on this system by modulating the expression of OPG and RANKL. While a few studies suggest an up-regulation of RANKL (Otsuka et al., 2005), most studies show a down-regulation of RANKL and an up-regulation of OPG (Bosshardt, 2008). This suggests that EMPs modulate the RANK- RANKL-OPG system most likely towards bone apposition. Of interest in this context is the observation that amelogenin knockout mice show increased hard tissue resorption (Hatakeyama et al. 2003). Furthermore, it has to be taken into consideration that some of the growth factors and cytokines that are up- regulated by EMPs directly up-regulate OPG and down-regulate RANKL production (Bosshardt, 2008). Thus, EMPs appear to be indirectly involved in the regulation of bone remodeling.

All the above emphasizes that EMPs have significant effects on different cell types involved in both periodontal wound healing and regeneration. EMPs caused an increase in cell attachment of epithelial cells, gingival fibroblasts, and PDL fibroblasts. A promotion of adhesion of osteogenic cells also does occur, but appears to be dependent on the cell differentiation/ maturation state. Furthermore, cell–matrix adhesion appears to be mediated, at least in part, by integrins (Bosshardt, 2008). In terms of cell proliferation, EMPs favour PDL fibroblasts over gingival fibroblasts or epithelial cells. In fact, the effect of EMPs on epithelial cells appears to be cytostatic, but not cytotoxic. With regards

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to osteogenic cells and their progenitors, the influence of EMPs on cell proliferation appears to decrease with increasing cell differentiation/maturation state (Bosshardt, 2008). EMD’s effect of enhanced cell proliferation and migration results in accelerated wound-fill rates in vitro using PDL fibroblasts, gingival fibroblasts, and osteoblast-like cells (Hoang et al. 2000). EMD, and, in particular its vehicle PGA, have antibacterial properties. The antibacterial effect of PGA has been known for many years (Olitzky, 1965). EMPs stimulate an increase in the synthesis of total protein as well as the synthesis of specific extracellular matrix molecules (glycoproteins and proteoglycans).

There is very little in the literature with regards to the analysis of separate fractions of EMD and identifying its bio-active components. One such study looked at the treatment of cultured human PDL cells with different fractions of EMD (Villa et al., 2015). It was demonstrated that both high and low molecular weight proteins contribute to the bioactive effect of EMD, and that there was a differential profile of cytokine release with different fractions of EMD (Villa et al., 2015). Both vascular endothelial growth factor (VEGF) and IL-6 secretion was only stimulated by EMD components larger than 20 kDa (Viilla et al., 2015). IL-6 appears to have a crucial role during wound healing as demonstrated by the fact that IL-6 deficient mice have been shown a delayed skin-wound healing (Gallucci et al., 2000). This observed effect might explain the proliferative activity of amelogenin on microvascular endothelial cells (Johnson et al., 2009). Classical inflammatory cytokines, for example, IL-1β and IL-6, are released early during the inflammatory response of wound healing and stimulate the production of other cytokines and growth factors, such as VEGF, initiating a cascade of molecular events leading to inflammation, tissue formation, and remodelling. On the other hand, lower molecular weight components enhanced cell proliferation and secretion of Monocyte chemo- attractant protein 1 (MCP-1) and IL-8, while simultaneously reducing IL-4 release (Villa et al., 2015). Some of the lower molecular weight fractions containing leucine-rich amelogenin peptide (LRAP) and truncated forms of tyrosine-rich amelogenin peptide (TRAP), promoted enhanced proliferation of PDL cells (Villa et al., 2015).

2.3.4.2. Emdogain® and Wound healing

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Of special note should be EMD’s effects on wound healing, which some have attributed partial reasoning for its improvement in clinical results. EMD treatment of PDL cells, gingival fibroblasts, and an osteosarcoma cell line (MG-63) resulted in enhanced wound- fill rates (Hoang et al. 2000). It was noted that the effect was statistically greater for PDL cells than for both gingival fibroblasts and MG-63 cells at early time points in the experiment. An in vitro wound-fill model showed that EMD, but not recombinant amelogenin protein, significantly enhanced PDL cell migration at the wound edge (Chong et al., 2006).

Effects of EMD on human microvascular endothelial cells (HMVECs) were also investigated. Low concentrations of EMD resulted in significant stimulation of HMVEC proliferation and chemotaxis when PDL cells were present (Schlueter et al. 2007). Moreover all doses tested increased angiogenesis. The combined presences of HMVECs with EMD stimulated a 750% increase in PDL cell migration (Schlueter et al. 2007). An almost 400% increase in VEGF concentration was demonstrated by ALP-positive PDL cells as well as a significant increase in TGF-β production in both ALP-positive and ALP-negative PDL cells in EMD-stimulated conditioned media. EMD proteins bind to wound extracellular matrix proteins (Schlueter et al. 2007). It is this interaction that tended to favour fibroblast adhesion over epithelial cells (Narani et al. 2007). Similarly, a significant increase in secreted VEGF was observed in adult human dermal fibroblasts exposed to EMD (Mirastschijski et al., 2004). EMD also significantly increased release of MMP- 2 from the fibroblasts and from human microvascular endothelial cells (Mirastschijski et al., 2004). The expression of genes involved in early inflammatory events of wound healing in human PDL cells is down-regulated following EMD treatment, while genes encoding growth and repair-promoting molecules are upregulated (Parkar and Tonetti, 2004)

Overall, EMPs downregulate the expression of genes and their products involved in early inflammatory events of wound healing and upregulate the expression of genes and factors encoding growth and repair-promoting molecules (Bosshardt, 2008). EMPs tend to promote vascularization, probably by increasing the numbers of endothelial cells (Bosshardt, 2008).

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2.3.4.3. Emdogain®; Effects on Bacteria

An ex vivo experiment utilizing the from 24 patients with demonstrated that EMD reduced total bacterial plaque viability by 46%, but when EMD was combined with PGA, only 21.4% of the bacteria remained vital (Sculean et al., 2001). When the PGA carrier was used alone, further reduction was seen with only 19.6% of plaque bacteria remaining vital (compared to negative control NaCl and positive control that showed vitality of 76.8% and 32.3%, respectively). It was suggested that Emdogain® (EMD+PGA) has antibacterial effects and that PGA contributed heavily to this characteristic. An in vitro study evaluating the effect of EMD on the growth of gram-negative periopathogens (Aggregatibacter actinomycetemcomitans, Porphyromonas gingivalis and ) described a marked inhibitory effect of EMD +PGA on growth of these microbes (Spahr et al., 2002). Emdogain®’s antibacterial effect and the major contribution of PGA to this effect were confirmed further by other independent authors (Arweiler et al., 2002; Newman et al., 2003; Walter et al., 2006) and these effects are likely not insignificant insofar as the generally beneficial effects of EMD on periodontal wound healing (Bosshardt, 2008).

2.3.4.4. Histological findings

Earlier studies focused on the use of EMD as an adjunctive treatment for regenerative periodontal surgery involving surgically created recession defects treated with either coronally advanced flaps (CAF) alone or adjunctive use of EMD (Hammarstrom et al., 1997). After 8 weeks of healing, it was demonstrated at the histological level that treatment with EMD induced the formation of acellular cementum, periodontal ligament and alveolar bone in all defects, but not in control defects.

Histological assessments were performed on block biopsies derived 6 months following treatment of intraosseous defects in 10 human teeth with EMD (Yukna & Mellonig, 2000). Complete periodontal regeneration (new cementum, new bone, and new periodontal ligament) was observed in three specimens, new attachment (connective tissue attachment/ adhesion only) in three specimens and a long junctional epithelium in

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four specimens. No evidence of root resorption, ankylosis or untoward inflammation was found (Yukna & Mellonig 2000). In one rat study, EMD caused significant improvement in incisional wound healing as evidenced by increased blood vessel formation and collagen content in the connective tissue of the wound (Maymon-Gil et al., 2016).

With regard to bone wound-healing, one model in rat femurs (Kawana et al., 2001) and in rat parietal bone (Sawae et al., 2002), demonstrated significantly higher bone volume fraction of newly formed bone trabeculae 7 days after injury in the EMD groups compared with the PGA (carrier) controls. Significantly greater trabecular bone area was noted around implants treated with EMD compared to control (PGA carrier) at both 14 and 30 days following placement of endosseous implants in rat femurs (Shimizu-Ishiura et al., 2002).

Treatment of periodontal defects with either GTR or EMD led to significant reductions in PD (probing depth) and gain in CAL (clinical attachment level) 6 months post-surgery (Sculean et al., 1999). The clinical improvements were characterized histologically by a new connective tissue attachment, and to a varying extent, new bone formation, which were noted in both groups (Sculean et al., 1999). In the cases treated with EMD, the new connective tissue attachment was followed by substantial bone regrowth in only two cases while in four specimens, bone regeneration was either minimal (0.5 mm) or the reformed connective tissue attachment was not accompanied by any signs of bone regeneration. All defects in the GTR group demonstrated new bone, cementum and periodontal ligament formation (Sculean et al., 1999).

In summary, these results demonstrate that it is possible to achieve periodontal regeneration with EMD treatment, but this does not occur in all cases. As noted above, although several factors could affect or cause these variable treatment outcomes (e.g. wound integrity, infection, patient age and systemic conditions) (Tonetti et al., 1996, Sanz et al., 2004, Jepsen et al., 2008) it is also possible that variations in the composition of EMD extracts could also contribute to the variances in outcomes of treatment that have been observed.

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2.3.5. Clinical application of EMD

2.3.5.1. Treatment of Intraosseous defects

In a multi-center, randomized, placebo-controlled study, contra-laterally located intraosseous defects were treated with either OFD alone or with OFD plus EMD. When analysed 3 years after treatment with EMD, it was noted that there were significant increases in CAL gains and pocket depths after 3 years of follow-up. Based on radiographic assessment it was noted that defects treated with EMD had on average 2.6 mm bone gain (66% fill) compared to no significant bone gain in control defects (Heijl et al. 1997). Three separate clinical trials demonstrated significantly higher soft tissue density as an additional benefit of EMD use in regenerative procedures using (Trombelli et al. 2002, Yilmaz et al. 2003, Jentsch & Purschwitz 2008). Another study investigated the use of EMD in regenerative therapy of deep intrabony defects in 172 patients with advanced chronic periodontitis in 12 centres (Tonetti et al. 2002). After debridement, EMD was placed in the test subjects, whereas omitted in the controls. Defects treated with EMD gained 3.1 ±1.5 mm of CAL, while the control defects yielded a significantly lower CAL gain of 2.5 ± 1.5 mm. In addition, pocket reduction was significantly higher in the EMD group (3.9 ±1.7 mm vs 3.3 ±1.7 mm in controls).

Both GTR (with either non-resorbable or bio-resorbable membranes) or EMD treatment in intra-bony defects, demonstrated significant and comparable CAL gains and defect fills, compared to OFD alone (Pontoriero, 1999). An additional benefit of EMD use vs GTR is that the latter tend to be prone to certain post-surgical complications, namely membrane exposure and subsequent infections, whereas in EMD treatments such complications are quite limited (Sanz et al., 2004). Interestingly enough, when comparing minimally invasive surgical techniques (MIST) with or without concomitant EMD treatment, authors observed similar results in treating intra-bony defects in both groups (Cortellini and Tonetti, 2007). A recent systematic review concluded that most studies did not demonstrate superior results of EMD addition to GTR procedures in intra-bony defect treatments (Miron et al., 2016). Indeed, treatment of single self-containing intrabony defects with EMD, GTR or a combination of both, demonstrated that all three

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regenerative procedures resulted in a significantly higher improvement of the clinical parameters compared to OFD. However, no additional benefit could be observed for the combined treatment of EMD + GTR. (Sculean et al., 2001b). Moreover, even though EMD can induce periodontal regeneration, its results were not consistent, whereas GTR was more reliable in this regard (Sculean et al., 1999).

In regards to combinations of EMD with other grafting materials, there appears to be a lack of a consensus in the literature. One recent systematic review and meta-analysis on 12 studies (434 patients) found that the combination of material with EMD led to statistically significant better outcomes in CAL gain and mean PD reduction, when compared to solely using EMD (Matarasso et al. 2015). Though the results are statistically significant, the clinical significance of the differences was questionable. Furthermore, many other studies have shown no significant benefits of adding EMD to bone grafting material vs separate EMD or bone grafting treatment in periodontal regeneration procedures (Kuru et al., 2006, Guida et al., 2007, Trombelli and Farina, 2008).

The literature is also equivocal with respect to specific types of bone grafting materials. Autogenous bone grafting with or without adjunctive EMD treatment yielded no significant difference in one study (Guida et al., 2007), while a second study on 2 and 3 wall defects demonstrated significant benefits of combining autogenous bone with EMD (Yilmaz et al., 2010). Two studies (Gurinsky et al., 2004 and Hoidal et al., 2008) demonstrated no added benefit in combining EMD with demineralized freeze-dried bone allograft (DFDBA) in treating intrabony defects (of 3mm or greater depth). On the other hand, a statistically significant additional 1 mm of PD reduction was attained with the combined EMD and DFDBA treatment of intra-osseous defects (Aspriello, 2001). A comparison of both freeze-dried bone allograft (FDBA) and DFDBA with and without EMD in the treatment of 69 intrabony defects showed significantly better PD reductions and CAL gains in combined EMD+DFDBA treatments, and even greater improvements with EMD+FDBA treatments as opposed to EMD alone (Ogihara and Tarnow, 2014). The lack of consistent results in clinical trials was also evident when comparing xenografts with or without adjunctive EMD. One study showed a statistically significant

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improvement in PD reduction (3.43 vs 1.91mm) and CAL gain (3.13 vs 1.72mm) with combined EMD and deproteinized bovine bone material (DBBM) treatment of >6mm intrabony defects after 6 months (Lekovic et al., 2000). Many other studies, however, concluded no significant differences in PD reduction and CAL gain with respect to combined xenograft and EMD treatment versus sole EMD or DBBM treatment of intrabony defects (Scheyer et al. 2002, Sculean et al. 2002b, Velasquez-Plata et al. 2002, Zucchelli et al. 2003).

The review of current clinical studies yields a mixed picture in terms of advantages of EMD use in treating intrabony defects, within a periodontal regenerative objective. Indeed, many studies have demonstrated that the application of EMD in the context of surgical therapy of deep intrabony defects has led to significantly more clinical attachment gain and defect fill, then open flap debridement alone. (Heijl et al. 1997, Forum et al. 2001). However, GTR, EMD or a combination of both, therapies yielded equal results in terms of gain of attachment levels, when intrabony defects were treated (Pontoriero et al. 1999). Furthermore, there is quite a wide variability in results both in terms of the types of grafting materials studied, but also within individual groups.

2.3.5.2. Treatment of Gingival Recession

EMD application in treatment modalities for gingival recessions has been studied substantially. In the majority of studies it has been shown that addition of EMD to a CAF lead to more keratinized tissue (Miron et al., 2016). A randomized controlled study of Miller class 1 and II recession treatment with either EMD+CAF or CAF alone, demonstrated superior stability of complete root coverage results during a 2 year follow- up (53% vs 23%) (Spahr et al., 2015). The additional benefit of EMD application to CAF has been independently confirmed by multiple other studies (Castellanos et al., 2006; Pilloni et al., 2006; Cairo et al., 2008).

At single recession defects, the addition of a connective tissue graft (CTG) or EMD with the CAF improved complete root coverage and is considered the standard for sites in the maxillary anterior and premolar sites (Tonetti and Jepsen, 2014). Histological studies

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have demonstrated that application of EMD as an adjunct to CAF enhanced periodontal regeneration, as opposed to CAF alone or in conjunction with a CTG, which resulted in long junctional epithelium (Mcguire and Cochran 2003). The combination of EMD and a sub-epithelial CTG in the treatment of 42 Miller class 1 and 2 recessions resulted in higher mean root coverage then CAF alone, and in higher percent compete root coverage then CAF alone after a year post-surgery (Roman et al., 2013). However the study failed to discern any significant difference between the addition of EMD to CTG and CTG alone.

In conclusion, it appears that the addition of EMD to CAF alone improves soft tissue thickness and root coverage outcomes, and it is capable of enhancing regeneration. However, multiple publications have noted its variability in clinical results when used in conjunction with a CTG (Henriques et al 2010; Rasperini et al, 2011), which thus puts into question any additional benefit of EMD use with the already existing gold standard for soft tissue grafting (connective tissue graft).

2.3.5.3. Treatment of Furcation defects

EMD has also been used in treatments of furcation defects. In one study comparing 20 furcal defects, contralateral matched sites with class II mandibular furcations were treated with either OFD or EMD. Re-entry in 6 months demonstrated a 2mm horizontal defect reduction in the EMD treatment group vs 0.8mm in the OFD group (Chitsazi et al. 2007). In the regeneration of class II furcations, a significantly improved PD reduction was associated with the addition of EMD to GTR and DFDBA treatment (Jaiswal and Deo, 2013). A multi-centre, split-mouth randomized control study on 90 class II mandibular furcations treated with EMD or GTR demonstrated superior results with the EMD treatment in both reduction of horizontal probing (2.6mm vs 1.9mm) and in patient- assessed post-operative experiences (pain and swelling) at 14-months re-entry (Hoffmann et al. 2006). In terms of proximal maxillary furcations, mixed results were achieved. One study demonstrated superior results with EMD +OFD vs OFD alone in treating class II defects, however it was noted that complete furcation closure was rarely observed (Casarin et al., 2010). However, another study found no significant difference in adding

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EMD to an OFD+ hydroxyapatite treatment modality of class II furcations (Peres et al., 2013). In conclusion, EMD’s use in furcation defects shows promise, however more definitive evidence from larger well-controlled studies is required.

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3. Aim, objectives and hypothesis

3.1. Statement of the problem

It has been demonstrated in a variety of clinical trials that the effectiveness of Emdogain® is somewhat inconsistent. The factors that might influence the variabilities seen in treatment outcomes have already been discussed above including surgical factors, among other things. To the best of our knowledge, no literature exists that describes the consistency or lack thereof in the levels or concentrations of the various proteins within different batches of Emdogain®. It is also not well known how various storage regimes might affect the composition and activity of EMD.

It is suggested here that the assembly and the structure of the proteins contained within Emdogain® may be sensitive to various factors, including temperature. If the therapeutic effectiveness of EMD/Emdogain® relies in part on the native conformation and content of its protein constituents, it is important to clarify the consistency of protein content/conformation in general (e.g. from batch to batch) and also to understand the effects of storage conditions (in this case temperature) on the former as well.

3.2.Objectives

• To determine whether there are any differences in EMP quantities between batches of EMD, and how other factors such as time in storage, or fluctuations in storage temperatures affect proteins in EMD. • To quantify any differences in levels of AMTN content (as a ‘reporter’ protein, see immediately below) between batches, as well as the effects of storage time and impact of temperature during storage on this protein • To identify and compare proteinase activity between batches of Emdogain®

The rationale for the use of AMTN as a ‘reporter’ protein is based on its putative role played in cell adhesion, a critical factor in healing of periodontal flaps (Nakayama et al., 2005) but also because of its minute quantity within EMD; any variabilities shown in

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AMTN could imply that there could be even larger changes in relative percent abundances of other more prevalent proteins (their functionality notwithstanding).

3.3. Hypothesis EMD Emdogain® has inherent variabilities in its protein composition as a result of the harvesting process and presence of proteinases. Moreover, protein content will decrease with storage at longer times and higher temperatures.

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4. Materials and Methods

4.1. Materials

5 Emdogain® (Straumann, Switzerland) batches were obtained from Mount Sinai Hospital Department of Dentistry and the University of Toronto, Faculty of Dentistry. Different LOT numbers on the products (along with differing expiry dates) signified a unique batch (See Appendix A).

Preparation of samples for mass spectrometry was done using the 2-D clean-up kit (GE healthcare Life Sciences) according to the manufacturer’s protocol.

Lab consumables and reagents were purchased through the University of Toronto MedStore (https://www.uoftmedstore.com/index.sz) unless indicated otherwise. All materials for the sodium dodecyl sulphate – polyacrylamide gel electrophoresis (SDS- PAGE) were prepared using stock reagents and chemicals supplied by Sigma-Aldrich (Oakville, ON). BenchMark TM pre-stained protein ladder was obtained from Life Technologies TM l, and the Mini-Protean ® Electrophoresis cell (Bio-Rad) was used for SDS-PAGE. Gels were stained using the ProteoSilver TM Silver Stain kit.

Protein transfer for western blotting was accomplished using Immuno-Blot polyvinylidene difluoride (PDVF) membranes (Biorad) via a semi-dry Western blotting transfer apparatus (LTF Labortechnik). Primary anti-porcine AMTN antibody was synthesized using reagents and protocol by Fisher Scientific (Ottawa, ON). The polyclonal antisera were produced in two white New Zealand rabbits. A standard protocol by Fisher Scientific was used for purification. Goat anti-rabbit antibodies (IRDye® 800CW Secondary Antibody by LI-COR®, part number: 925-32211 ) was used as a secondary antibody. The fluorescence was then captured and image produced using the LI-COR® Odyssey Imaging system.

The enzymatic activity of batches was investigated using EnzChek® Protease Assay

(Thermo Fisher), using casein derivatives labeled with green-fluorescent BODIPY® FL

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(E6638). FLUOstar® Omega multi-mode microplate reader (BMG Labtech) was used to detect and quantify resulting fluorescence.

4.2. Methods 4.2.1. Sample treatment

20 µL aliquots of Emdogain® were placed in sealed 500 µL Eppendorf tubes and incubated at one of 3 different temperatures (4°C, 23°C and 37°C) for one of 5 different times (3 hours, 9 hours, 24 hours, 72 hours and 168 hours), resulting in a total of 15 samples per batch . These incubations were performed as follows:

4°C – samples were placed in the laboratory fridge (set at 4°C)

23°C – samples were submerged in a water bath (set at 23°C). The water bath unit was placed in a controlled refrigerated room (4°C).

37°C – samples were placed in an incubator (set at 37°C)

At the completion of the incubation, samples were centrifuged briefly to collect all liquid and stored at -80° C to be used for further experiments.

In order to enhance manageability of the viscous Emdogain® samples for all future experiments, they were diluted with Tris Buffered saline (TBS) at a ratio (volume/volume) of 1:4 and vortexed for 5 minutes.

4.2.2. Primary antibody production (anti-porcine AMTN)

The peptide H2N-GTPRGPFPTSSGTDDDFD-COOH, corresponding to amino acids 163-180 of the porcine AMTN protein, was synthesized and coupled separately to keyhole limpet hemocyanine (KLH) and bovine serum albumin (BSA) by Fisher Scientific (Ottawa, ON). The peptide-KLH conjugate was used for the production of polyclonal antisera in two white New Zealand rabbits according to a standard 120-day protocol (Fisher Scientific). The antisera were purified by antigen-specific affinity

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chromatography using a standard protocol (Fisher Scientific), eluted with 50 mM glycine-HCl/0.5 M NaCl (pH 2.3), neutralized with NaOH, aliquoted and stored at -80 °C. 4.2.3. Tandem Mass Spectrometry (TMS)

TMS was used to identify, quantify and compare protein composition in the different batches of Emdogain®. For the first TMS analysis (Run #1), 3 of the batches had samples at two temperature and time extremes analyzed (i.e. 3 hours at 4°C and 168 hours at 37°C). Using the same TMS protocol for the second analysis (Run # 2), 1 sample from each of the 5 batches (3hours at 4°C) was used.

10 µL samples of Emdogain® (1:4 dilution in TBS) were processed using the “2-D clean-up kit” according to the manufacturer’s protocol (GE healthcare Life Sciences) in order to precipitate EMD proteins into pellets for TMS analysis. Analysis of the precipitated samples was conducted at the SPARC BioCentre at Sick Kids Hospital according to their protocol:

Samples were reduced, alkylated, digested, and TMT labelled according to manufacturer’s directions (Thermo Fisher TMT 10 Plex, Product 90110). Labelled peptides from all samples were combined and lyophilized. Peptides were then cleaned up using a homemade SCX column, and lyophilized.

Samples were analyzed on a Thermo Scientific Orbitrap Fusion-Lumos Tribid Mass Spectrometer (ThermoFisher, San Jose, CA) outfitted with a nanospray source and EASY-nLC 1200 nano-LC system (ThermoFisher, San Jose, CA). Lyophilized peptide mixtures were dissolved in 0.1% formic acid and loaded onto a 75µm x 50 cm PepMax RSLC EASY-Spray column filled with 2µM C18 beads (ThermoFisher San, Jose CA) at a pressure of 800 Bar and a temperature of 60C. Peptides were eluted over 120 min at a rate of 250nl/min using a gradient set up as 0%-40% gradient of Buffer A (0.1% Formic acid; and Buffer B, 0.1% Formic Acid in 80% acetonitrile).

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Peptides were introduced by nano-electrospray into the Fusion-Lumos mass spectrometer (Thermo-Fisher). Data was acquired using the MultiNotch MS3 acquisition with synchronous precursor selection (SPS). MS1 acquisition was performed with a scan range of 550m/z - 1800 m/z with resolution set to 120 000, maximum injection time of 50ms and AGC target set to 4e5. Isolation for MS2 scans was performed in the quadrupole, with an isolation window of 0.7. MS2 scans were done in the linear ion trap with a maximum injection time of 50ms and a normalized collision energy of 35%. For MS3 scans, HCD was used, with a collision energy of 30% and scans were measured in the orbitrap with a resolution of 50000, a scan range of 100m/z-500m/z, an AGC Target of 3e4, and a maximum injection time of 50ms.The dynamic exclusion was applied using a maximum exclusion list of 500 with one an exclusion duration of 25 s.

The Sus scrofa genome database was used to identify the peptide sequences. Reporter Ion intensities from the TMS analysis were used as a proxy for relative abundances of the different proteins and peptides to reveal the composition and relative abundance of Emdogain® proteins. Due to limited availability of samples and prohibitive cost, only one sample from each of the 15 time and temperature points was analyzed.

4.2.4. Sodium dodecyl sulphate – polyacrylamide gel electrophoresis (SDS-PAGE)

15% SDS-PAGE was used to separate proteins for each sample; two sets of gels were run for each batch: one set was silver-stained and the other set was used for Western Blotting. Samples for both gels were prepared identically. For each sample, 1 µL of Emdogain® (1:4 dilution in TBS) was combined with 10 µL of Bovine Serum Albumin (BSA; 1:32 dilution of a 2mg/mL BSA stock in 1xTBS) as an internal standard, 5 µL of 4x Running buffer and an additional 4 µL of TBS. Samples were denatured at 95°C for 5 minutes, after which they were loaded onto the gel, with the addition of the BenchMark TM pre- stained protein ladder by Life Technologies TM in the first lane of each gel. The only additional step taken for the gels that were going to be used for Western blotting was the addition of a positive control. One of the lanes of each gel was loaded with a solution of 5

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µL of porcine AMTN (4.37 µL/mL), 10 µL PBS and 5 µL of 4x running buffer. Using the Mini-Protean ® Electrophoresis cell (Bio-Rad), a total of 4 gels were run for each batch – 2 to be used for silver staining and 2 for Western blotting. A pair of gels was required for each experiment as each gel had 10 wells but each batch had a total of 15 samples.

Silver staining was performed using the ProteoSilver TM Silver Stain kit and protocol by Sigma-Aldrich ®. Silver-stained gels had each sample’s protein profile assessed by comparing band intensities (using Image J software 1.51s; National Institutes of Health, USA). A histogram of band intensities vs. molecular weights (distance travelled in the gel) was created i.e. a vertical cross-section of each lane. For a given batch, the histogram of band intensities of each of the three temperature treatments for one time point were superimposed, with the BSA band peak for each graph being made equal (serving as an internal control). A qualitative assessment of any patterns amongst the three temperatures as a function of time was then conducted.

A previous pilot experiment was used to test the sensitivity threshold of the primary antibody for porcine AMTN – a dilution series of the stock porcine AMTN was performed, and the resulting SDS-PAGE was Western blotted with anti-AMNT antibodies (See Appendix B). The detection threshold for the Western Blotting was determined to be between 1.165µg/mL to 4.659µg/mL.

Western blotting of the second set of gels for each batch was performed using anti- porcine AMTN antibodies. Specifically, the protein bands from the set of gels was transferred to PDVF membranes using a semi-dry Western blotting transfer apparatus (100V for approximately 1 hour). The membranes were then blocked overnight with blocking buffer (1.5mg milk protein powder + 50mL TBST at 4 oC). This was followed by a 24 hour incubation period with the primary antibody (polyclonal rabbit anti-porcine AMTN diluted 1:350 with blocking buffer). After the membranes were washed 5 times for 5 minutes each with TBST, the membranes were incubated with the secondary antibody (goat anti-rabbit 800nm in a 1:4000 dilution with TBST; LI-COR®). The fluorescence was then captured and an image produced using the LI-COR® Odyssey Imaging system.

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Differences in fluorescence intensities of AMTN bands were analyzed using Image J software. Specifically, the fluorescence images of each gel were inverted and converted to an 8-bit gray scale. The band intensities were then analyzed and normalized (dividing the intensity of the respective BSA band for each sample).

4.2.5. EnzChek® Protease Assay

A fluorescence-based assay “EnzChek® Protease Assay Kit” was used to identify presence of protease activity and any differences between 4 batches of Emdogain®. EnzChek® Protease Assay contains casein derivatives that are heavily labeled with the pH-insensitive green-fluorescent BODIPY® FL (E6638). Protease-catalyzed hydrolysis releases highly fluorescent BODIPY FL dye-labeled peptides. The accompanying increase in fluorescence is proportional to protease activity and was detected using the FLUOstar® Omega multi-mode microplate reader. The settings for the detection of the fluorescence were set at 485 µm (excitation) and 520µm (emission).

The 3 hours at 4°C treatment sample was utilized from each batch for this part of the experiment. Three repeat samples of each batch were used, with an additional three readings taken of each, to create replicate data points for statistical analysis.

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5. Results

5.1. TMS

TMS, performed at the SPARC BioCenter at Sick Kids Hospital, was used twice to analyze samples. In the first analysis, three batches of Emdogain® had two samples analyzed each (3x2 = 6 samples total). A 3 hours/ 4°C sample and a 168 hours/37°C sample were used from each batch (to compare effects of time and temperature extremes). A total of 528 proteins and peptides were identified. The TMS identified the presence of pig Enamelin, Amelogenin, MMP 20, Ameloblastin, AMTN, ODAM, KLK4, as well as factors like Bone morphogenetic protein 4 and 7 (BMP-4, BMP-7). An additional finding of interest was the presence of the tissue inhibitor of metalloproteinases 1 (TIMP-1) in all samples tested. There were minute differences in reporter ion intensities of enamel-related proteins between samples (Appendix C; figure 5), however, no definitive temperature or time dependent patterns were noted. The only difference noted was the overall smaller quantities of proteins in the Batch 1 sample (3 hours at 4oC) as compared to the 168 hours at 37oC Batch 1 sample.

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Figure 5: Bar graph of EMP abundances from TMS analysis for 2 samples per batch (3 batches total); one sample at 3 hours x 4°C and one at 168 hours x 37°C.

The second TMS analysis was performed on all five batches, with one sample (3 hours at 4°C) from each batch assessed. A total of 200 proteins and peptides were identified. It is important to note that the SPARC BioCenter had updated the TMS instruments and software, resulting in a more discriminatory and precise protein identification protocol. As in the previous analysis, porcine enamelin, amelogenin, ameloblastin, MMP20, KLK4, and BMP-4 were identified (see Appendix D; figure 6). Of note is the fact that both AMTN and ODAM were no longer detected by TMS.

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Figure 6: Bar graph of EMP abundances (reporter ion intensities) in each of the 5 Emdogain® batches; each sample at 3 hours and 4°C.

The data from the second TMS analysis allowed for a qualitative assessment of any inter- batch differences. TMS data showed that overall batch 1 contained substantially less of each EMP compared to the other batches. Batch 3 contained far less Amelogenin and Ameloblastin then batches 2,4 and 5. Furthermore, Batch 2 and 3 contained substantially less Enamelin detected then batch 4. Out of all the EMPs, the amelogenin family of proteins seemed to have the most variable abundance between batches.

5.2. Silver stained SDS-PAGE

The Emdogain® samples that were analyzed using the SDS-PAGE technique produced a unique and reproducible banding pattern (Figure7; Appendix E). It should be noted that gels for batches number 4 and 5 had a slightly different protein ladder used with unique molecular mass markers. As such, it was possible to combine the information provided by both protein ladder markers to estimate the molecular mass of all bands within the

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samples.

Figure 7: Representative section of silver-stained 15% SDS-PAGE with labeled bands. Left lane was loaded with the BenchMark TM pre-stained protein ladder; right lane contains Emdogain® sample from batch number 5 treated at 4oC for 3 hours.

There was a distinct intense band at about 60kDa (band 1), which represents the BSA control. However, all of the observable bands in the gel representing the EMD sample ranged in molecular mass within a span from 10kDa to about 37 kDa. Using the cross- section of band intensities plotted as gray scale intensities vs molecular mass, the most intense band of the EMD samples had a molecular mass slightly below the 19kDa mark (band 8). This band also was the broadest. A distinct, but less intense band was found at a higher mass of about 19kDa (band 7). A dark band was identified slightly below 37kDa (band 2), and a neighbouring band less intense then this had an approximate mass of 30kDa (band 3). Most gels also demonstrated a much fainter band right below the 30kDa mark (band 4). Two faint but discernable bands found in all SDS-PAGE samples were also located at about 20kDa (band 6) and slightly below the 26 kDa mark (band 5). A few smaller molecular mass bands were also identified, namely a distinct band slightly larger than 15kDa (band 9). There were also potentially two smaller bands below 15kDa but

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greater then 10kDa in molecular mass (band 10 and 11), however due to the staining sensitivity and gel resolution, the distinction between the two bands on the histogram is rarely captured, and as such either appears as a single peak or as minute twin peaks.

Figure 8: Representative histograms of band intensities vs Molecular weight for Emdogain® batch 4; Each graph represents the band intensity profiles of the three temperature treatments for a single time point.

With respect to the temperature and storage time treatments, the analysis of protein band profiles of each sample was unable to detect patterns in differences between time and temperature treatment (see figure 8). Though the overlapped histograms of band intensities (normalized by matching the peak of the first band to the right – the BSA control) for each of the three temperatures demonstrated differences, these differences did not display any trend as the storage time progressed. Indeed, the changes in profiles as treatment time progressed within a given batch appeared to be random in nature. This same random fluctuation in relative changes of band intensities was seen in all 5 batches (see Appendix F).

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5.3. Western Blotting

A

B

Figure 9. A: Representative Western blot membrane treated with anti-pAMTN of all 15 samples of batch 3. Lanes ordered in triplets according to time treatment (increasing order). Within each triplet, the temperature treatments increase sequentially (i.e. 4oC, 23oC and 37oC) B: Line graph of AMTN band intensities (normalized) vs time of storage. Lines grouped according to temperature treatment.

The Western blot of the SDS-PAGE gels brought forth interesting results. For one, the positive results of anti-pAMTN bands in each sample confirm the presence of this protein in each batch analysed. Furthermore, as a result of previous experiments using dilution series of pure pAMTN, we are confident that the concentration of pAMTN in Emdogain® is at least between 1.165µg/mL to 4.659µg/mL. After normalizing the

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intensity of the fluorescent pAMTN bands with the BSA control band, it appears that there is no correlation between pAMTN quantities and the temperature and time treatments. Indeed, there was no significant difference in band intensities between treatments within any of the 5 batches of Emdogain® (see Appendix G and H).

5.4. Fluorescence Enzyme assay

Figure 10: Data represents the mean ± standard deviation of the fluorescence for each control or batch; * represents a significant difference relative to controls (p <0.005) to batch 2 (p<0.0001); ** represents a significant difference relative to controls (p <0.0005), batch 2 (p<0.0001) and batch 4 (p<0.0001).

For the final part of this project, four of the Emdogain ® batches (batch 2,3,4 and 5) were analyzed for protease activity using the EnzChek® Protease Assay Kit. Three repeat samples of each batch were used and the fluorescence was read with FLUOstar® Omega multi-mode microplate reader. With respect to enzymatic activity through the evaluation of fluorescence as a direct proxy, it was observed that batch 3 had significantly greater fluorescence relative to the buffer and BODIPY FL substrate control (p <0.005) and to batch 2 (p<0.0001). Furthermore, Batch 5 had significantly greater fluorescence relative to the buffer and BODIPY FL substrate control (p <0.0005), batch 2 (p<0.0001) and batch 4 (p<0.0001). Mean fluorescence in Emdogain® batch 2 and 4 were not statistically significant from controls. It can thus be inferred that enzyme activity is

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present in certain Emdogain® batches. Moreover, there appears to be differences in protease activity among batches; indeed in some there appears to be no protease activity whatsoever (i.e. comparable to control).

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6. Discussion

TMS analysis identified all the anticipated EMPs within the EMD Emdogain®, and as expected, the most abundant protein in the enamel matrix was amelogenin. Unlike ameloblastin and enamelin, which have significant post-translational modifications, amelogenin has only one site that is phosphorylated (Bartlett 2013). The uncleaved secreted pig amelogenin protein is 173 residues in size (P173), with a molecular weight of about 26 kDa (Ryu et al., 1999). This is very similar to the full-length human amelogenin, which contains 175 amino acids (Salido et al., 1992). The majority of the secreted P173 is cleaved after Ser148 to generate P 148, a 20-kDa amelogenin (Met1- Ser148). Further post-secretory processing results in cleavage after Trp45 to yield the tyrosine-rich amelogenin polypeptide (TRAP; P45; Met1-Trp45) and the 13-kDa amelogenin (Leu46-Ser148) (Fincham et al., 1981).

P148 is the most abundant cleavage product in post-secretory stage pig enamel, followed by TRAP and the 13kDa amelogenin. By matching molecular weights, it appears that P148 is best represented by band #7 in the silver-stained SDS-PAGE gels (figure 7). Leucine-rich amelogenin (LRAP; P56) is another major amelogenin isoform with a mass of 5.4kDa, and is a product of amelogenin alternative splicing (Gibson et al., 1991). However, LRAP mRNA is about one tenth as abundant as the major amelogenin isoform (Yuan et al., 1996). Band # 10 (figure 7) is potentially representative of the position the 13kDa amelogenin fragment would travel in the SDS-PAGE. In developing enamel, MMP-20 is the main proteolytic enzyme present during the secretory stage of amelogenesis. It processes amelogenins into all major identified cleavage products that accumulate and are only slowly degraded further by MMP-20. TRAP and LRAP accumulate because MMP-20 has little activity to degrade them (Nagano et al., 2009). Due to the later appearance of KLK-4 in the transition stage, this enzyme does not contribute to the processing of amelogenin during the secretory stage, but does degrade it and all of its cleavage products (Nagano et al. 2009) during the maturation stage.

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Ameloblastin is the second most abundant EMP (Fukae and Tanabe, 1987). However, in vivo, intact ameloblastin (62 kDa) has never been isolated because it exists in only trace amounts (Chun et al., 2010). MMP-20 cleavage at the N-terminus (after Gln130, Arg170 or Ala222) accounts for the three initial ameloblastin products in the porcine enamel matrix, which include the 17 and 13kDa products originally observed on SDS-PAGE gels (Fukae and Tanabe, 1987) and also includes a 15 kDa cleavage product (Fukae et al., 2006). Band #9 in the silver-stained SDS PAGE (figure 7) appears to be at a favourable position to be matched as one of the ameloblastin cleavage products (~15kDa). The largest and most intense band in Emdogain® samples was band #8 (figure 7), which was located between 15 and 19kDa. It is possible that this wide band is actually a collection of multiple heavy stained bands, but due to the high protein concentrations and staining times (required to develop some of the less intense bands), these bands blended together to form a single band. It is thus possible that the 17kDa ameloblastin cleavage product is contained within the larger band #8.

The enamelin protein is initially secreted as a 186 kDa precursor phosphorylated glycoprotein. This protein is rapidly processed from its C-terminus to generate short-lived 155, 145, and 89 kDa N-terminal cleavage products found near the enamel surface (Hu et al., 2000). The 32 kDa enamelin proteolytic fragment (amino acids 136–241) is the only stable domain that accumulates in the deeper enamel, and is highly conserved among species, including that in porcine (Brookes et al., 2011). This 32kDa enamelin fragment seems to match the position of band #2 (figure 7) in the silver-stained gels.

With respect to the enzymes, previous studies have reported KLK-4 appearing at about 30kDa on an SDS-PAGE gel (Nagano et al., 2009), which correlates well with band #3 in this study’s silver-stained gels. MMP-20, on the other hand, could not be identified on SDS polyacrylamide gels, but it did appear as a 41 and 45kDa doublet on western blots (Nagano et al. 2009). In figure 7, there are two faint un-labeled bands below band #1 at a similar mass. Whether these bands contain MMP-20 is, at best, an assumption. Collectively, it is not possible to infer if any of the aforementioned SDS-PAGE band correlations made in this discussion are accurate without more specific analysis and identification of specific bands (e.g. immune-fluorescence or mass spectrometry).

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What can be ascertained from the combined silver-stained SDS-PAGE, Western blotting and TMS analysis is that neither temperature nor time of storage appeared to have an appreciable effect on EMP abundance. Indeed, as time progressed none of the silver- stained SDS-PAGE gels demonstrated any new band formations at lower molecular weights, which would signal the accumulation of degradation products. Though the silver-stained SDS-PAGE gels did provide a window to the protein mass distribution profile of Emdogain®, the method might have been too crude to detect appreciable changes in single protein quantities. Any differences that were observed on the gels may have been a result of error or artifacts from the image capturing. There was potential experimental error in the pipetting and handling/preparation of samples. Also, potential lack of homogeneity of the gels and/or the staining protocol might have created apparent differences in bands intensities that may not be real.

Though one of the TMS analyses did not identify either AMTN or ODAM, perhaps due to their minute quantities, the more specific and sensitive western blotting was able to positively identify AMTN within all samples and batches. Only a single fluorescent band was observed at the corresponding molecular mass of the positive porcine AMTN control (approximately 22kDa), which also seems to correlate with band #5 on the silver-stained gels (figure 7). Interestingly, no bands at lower molecular weights appeared on the western blot. This potentially indicates a lack of any degradation products or at least an absence of cleaved peptides with recognizable epitopes for antibody binding. AMTN is not post-translationally modified to any significant degree (Lacruz et al., 2012). The porcine AMTN sequence does indicate quite a few potential cleavage sites by either KLK-4 or MMP-20 (figure 11). In silico predicted MMP-20 cleavage sites (Turk et al., 2006) seem to be far more abundant than those for KLK-4 (Matsumura et al., 2005). It is interesting to note that the basal lamina re-establishes along the distal surface of the ameloblast membrane at the same time as KLK-4 is being secreted (Lu et al., 2008). The basal lamina proteins AMTN (and ODAM) both may be involved in mediating the adhesion of maturation stage ameloblasts to the enamel surface (Lu et al., 2008). The fact that AMTN is able to assemble and function in the presence of KLK-4 in the extracellular space suggests that it is not subject to degradation by KLK-4 (Lu et al., 2008). Moreover,

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it has been observed that AMTN is not degraded by either enamel proteases to any significant extent (Lacruz et al., 2012). Indeed, though all the EMPs get cleaved by MMP-20 during the secretory stage and further degraded and eliminated from the enamel matrix by KLK-4 during the transition/maturation stage, AMTN has very limited sequence similarity to the other EMPs (Moffatt et al., 2008). It is possible that this protein’s tertiary structure prevents access to the predicted cleavage sites. Alternatively, changes in ion concentrations during amelogenesis might alter enzyme activity or site recognition. MMP-20, for example, appears to be most active at high calcium concentrations (33.4 mM CaCl2) and least active at either high phosphate or both high calcium and phosphate concentrations (high calcium= 33.4 mM CaCl2, high phosphate=

22.9 mM KH2PO4). The processing of amelogenins and other EMPs by MMP-20 in a developing matrix is thus a dynamic process and can be controlled by changes in concentration of mineral ions (Khan et al., 2013)

Figure 11. Porcine AMTN amino acid sequence (Sus scrofa database www.uniprot.org). Protein ID: F1RUQ3. Downward facing arrows identify predicted KLK4 cleavage sites; upward facing arrows indicate predicted MMP-20 cleavage sites. Shorter arrows (half-arrows) indicate lesser likelihood.

AMTN’s and ODAM’s resistance to enzymatic degradation might also be related to the adhesion of these proteins to the enamel mineral. The adsorption of the proteins to the hydroxyapatite (HA) might serve as a protective feature against enzymatic proteolysis. It

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has been shown that certain proteins in the acquired enamel pellicle (AEP) are more resistant to enzyme degradation once they are bound to the mineral surface versus when they are free-floating in the enzymatic environment of the saliva. Incubation of histatin-1 in saliva (a constituent of the AEP) in the absence of HA resulted in about 28% of histatin-1 being degraded at 30 min and 83% by 120 min (McDonald et al, 2011). In contrast, incubation of histain-1 adsorbed onto HA in that same saliva resulted in only a minimal degradation of 3% at 30 min and 20% at 120 min (McDonald et al., 2011).

All evidence above indicates that Emdogain shows remarkable stability under the tested conditions. The reasons for this may include the following: Perhaps the formulation of the PGA carrier, within which the EMD of Emdogain® is contained, lends a resiliency to the tested storage conditions. PGA acts as a carrier for the deposition of EMD onto the affected tissue. Amelogenins are insoluble at physiological pH but can be dissolved at either low or high pH. Specifically, the PGA vehicle is an acidic vehicle, with a pH below 4.5 (Wennstrom and Lindhe, 2002). The EMD precipitates from the carrier once it reaches physiological pH and temperature and adheres to the root surface to form a compact layer that is ultimately believed to promote selective cell proliferation and periodontal tissue regeneration (Wennstrom and Lindhe, 2002). It is important for EMD to remain stable against denaturation and irreversible aggregation during the various stages of manufacturing, storage and delivery, if its regenerative capacity is to be maintained (Apicella et al, 2015). There have been a few investigations into the effects of additives on the stability of EMD. For example, presence of arginine reproduces the benefits of a reduction in pH and/or an increase in the ionic strength of the buffer for the stability of EMD. This lessens the need to increase the phosphoric acid content (for pH control), and it had no adverse consequences for precipitation of EMD at physiological pH (Apicella et al., 2015). Interestingly, TMS analysis did positively identify minute amounts of Metalloproteinase inhibitor 1 (TIMP-1) within all the tested batches. Whether this enzyme inhibitor, other components (salt or ion concentrations), or perhaps the pH of the formulation stabilize the proteins and inhibit enzymatic activity is yet to be determined. The fact that significant enzymatic activity was detected in the protease

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assays after the samples were treated and diluted in buffers prior to substrate addition, might further support the positive role of the carrier in EMD stability.

A few additional points with respect to the TMS analysis findings should be briefly mentioned. One notable difference between the first and second TMS analysis was the substantial decrease in amount of proteins identified, and in particular the fact that both AMTN and ODAM were no longer detected by TMS in the second analysis. In part, this disparity might be explained by a change from MS-2 to MS-3 analysis that was undergone at the SPARK Bio-centre prior to the second TMS analysis. This upgrade in TMS might have led to more selective peptide identification of only parent proteins with higher abundances. In addition, as mentioned with regards to the first TMS analysis, the only substantial difference noted was the smaller quantities of all proteins in the Batch 1 sample (3 hours at 4oC) as compared to the 168 hours at 37oC Batch 1 sample. This difference is most likely not due to treatment effects, but rather a result of an error either in the preparation of samples for TMS or the TMS analysis itself. Of course, multiple independent repeats of the analysis with more samples would be required to allow for more conclusive observations; a solitary test of a single sample has no statistical weight and brings in the possibility of error during the experimental procedure.

However, there does appear to be certain differences between batches of Emdogain® with respect to protein abundances and enzymatic activity. Indeed, TMS analysis did reveal some differences in EMP abundances between batches. Of note was that batch 1 had the smallest relative quantities of each EMP compared to the other four batches. Batch 1 had the earliest expiry date (however, at the time of analysis it was not yet expired), and as such was probably the oldest batch tested. One can speculate that because of its longer shelf time, more of the EMPs have been degraded over time. These inter-batch differences support the hypothesized heterogeneity that potentially exists with the harvesting of EMD from developing porcine enamel. Indeed, as the enamel develops, many changes occur both in the amounts and types of proteins being secreted, as well as the gradual degradation of the extracellular proteins as the crystals mature. There is currently no publicly accessible information with regards to the extraction processes and

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production protocols of Emdogain®, and thus potential culprits for a lack in product homogeneity cannot be ascertained. Irrespective of the cause of the observed inter-batch differences, the lack of homogeneity in this product can be extrapolated as a potential factor in the inconsistencies observed in clinical results. Indeed, multiple publications on root coverage have noted Emdogain®’s variability in clinical results when used in conjunction with a CTG (Henriques et al 2010; Rasperini et al, 2011). Furthermore, no additional benefits in treatment of intra-osseous defects were observed for the combined treatment of EMD and GTR, versus either alone. (Sculean et al., 2001b). Moreover, even though EMD can induce periodontal regeneration, its results were not consistent, whereas GTR was more reliable in this regard (Sculean et al., 1999). The observations attained in this project with regards to batch composition variability, in conjunction to conclusions derived from the literature, demonstrate a potential area of further investigation. Product uniformity also represents an arguably realistic goal, and thus there exists room for modification and improvement of the existing product.

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7. Conclusion and Future Directions

All the proposed biologically active agents in EMD, EMPs, were identified and present in the Emdogain® samples analyzed. The abundances of these EMPs appeared to be stable to all tested time and temperature conditions. However, there were notable differences in their quantities between different batches. Furthermore, there were statistically significant differences in enzymatic activity present within certain batches of the EMD. Such results indicate that there potentially exists heterogeneity in the contents of this product. Given the observed stability of the product to differing storage conditions, it can be speculated that such variability potentially lies in the harvesting of the developing enamel matrix, which it itself is a dynamic process and far from uniform.

The experiments and results from this project served as a pilot investigation into the potential batch-to-batch variability of protein components and their relative abundances in Emdogain®. Prudent follow-up work would involve gaining statistical power with adequate amounts of replicates to confirm the conclusions drawn from this project. Arising from the work presented here, one potential next step would be to correlate differences in EMP abundances with effects on different cell types in vitro and potentially clinical in vivo investigations. It would be of interest to determine which, if any, EMPs contribute to the variability in clinical outcomes through batch-to-batch differences in their abundances. Clinical outcomes that could be assessed would include PD reductions, CAL gains and radiographic bone levels. With respect to batch variability and wound healing, possible parameters that could be assessed include strength of flap attachment as a function of time, inflammatory biomarkers levels and patient-reported pain levels. Additionally, histological studies could be used to further investigate the effect of different EMD batches on periodontal regeneration, and whether the relative amounts of certain EMPs have an effect on tissue regeneration amount. Such studies would allow for deciphering which combinations of EMPs, and at what minimal concentrations, are needed to retain clinical efficacy for periodontal regeneration. These types of correlations, if present, would potentially open the door to test products

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consisting of recombinant EMPs, which would allow for greater production control, homogeneity and resulting clinical consistency then that seen in current EMD products.

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APPENDICES

Appendix A: Emdogain® batch LOT numbers and expiry dates

Batch LOT Expiry date 1 JJ366 2017-03 2 JR049 2017-05 3 LP632B 2018-06 4 LN602C 2018-05 5 LH966B 2018-04

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Appendix B: Western Blot of porcine AMTN dilution series

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Appendix C: EMP abundances (represented as reporter ion intensities) from TMS analysis for 2 samples per batch (3 batches total); one sample at 3 hours x 4°C and one at 168 hours x 37°C.

Abundance (Reporter Ion Intensity) Batch 1 Batch 2 Batch 3

Protein (Sus number of Predicted 3 hours at 168 hours 3 hours at 168 hours 3 hours at 168 hours Scrofa) AA MW (kDa) 4°C at 37°C 4°C at 37°C 4°C at 37°C

Enamelin 1142 128.3 5775.1 10697.5 9307.2 8658.7 10670.3 9393.8

MMP-20 483 54.1 4835.1 13364.6 10721.2 10481.9 8789.8 9460.8

Ameloblastin 421 44.9 1559.4 4749.5 3916.1 4259.6 3391.7 3886.6 Amelogenin (combined) 132-254 15-27.2 8582.7 23071.5 16893.1 16909.6 14228.8 17222.4

KLK-4 254 27.2 149 361.6 298.2 311.8 220.2 248

BMP-4 409 46.7 9.2 25.8 19.3 19.6 20.7 14.4

BMP-7 431 49.2 30.4 88.2 68.1 76.3 66.3 72.4

TIMP1 207 23.1 46.7 147.6 105.3 109.3 100.3 117.7

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Appendix D: EMP abundances (reporter ion intensities) in each of the 5 Emdogain® batches; each sample at 3 hours and 4°C.

Number of Predicted Abundance (Reporter ion intensity) Amonio MW Proteins (Sus Scrofa) acids (kDa) Batch 1 Batch 2 Batch 3 Batch 4 Batch 5

Enamelin 1142 128.3 3763.7 5811.8 6172.4 7663.8 8431.1

MMP-20 483 54.1 2185 5424.7 4798.4 5145.7 4766.9

Ameloblastin 421 44.9 576.4 1345.4 969.6 1278.9 1209.9

Amelogenin (combined) 132-254 15-27.2 17012.7 34553.3 17909.5 36734.3 27957.6

KLK-4 254 27.2 145.4 385.1 297.1 345.7 387.3

BMP-4 409 46.7 9.6 26.7 21.8 29.3 27.4

TIMP-1 207 23.1 45.4 101.5 94.1 82 81.2

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Appendix E: Silver-stained SDS-PAGE gels of Emdogain ® batch samples at difference time and temperature treatments.

Appendix E.1: Emdogain ® batch 1 with BSA band

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Appendix E.2: Emdogain ® batch 2 with BSA band

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Appendix E.3: Emdogain ® batch 3 with BSA band

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Appendix E.4: Emdogain ® batch 4 with BSA band

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Appendix E.5: Emdogain ® batch 5 with BSA band

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Appendix F: Histograms of band intensities vs Molecular weight for each Emdogain® batch; Each graph represents the band intensity profiles of the three temperature treatments for a single time point.

Appendix F.1: Histograms for Emdogain® batch 1

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Appendix F.2: Histograms for Emdogain® batch 2

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Appendix F.3: Histograms for Emdogain® batch 3

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Appendix F.4: Histograms for Emdogain® batch 5

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Appendix G: Western blot membranes treated with anti-pAMTN of all 15 samples for each Emdogain® batch. Lanes ordered in triplets according to time treatment (increasing order). Within each triplet, the temperature treatments increase sequentially (i.e. 4oC, 23oC and 37oC)

Appendix G.1: Western blot for Emdogain® batch 1

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Appendix G.2: Western blot for Emdogain® batch 2

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Appendix G.3: Western blot for Emdogain® batch 4

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Appendix G.4: Western blot for Emdogain® batch 5

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Appendix H: Line graphs of normalized AMTN band intensities (computed from the Western blots) vs time of storage. Lines grouped according to temperature treatment. Note: Error bars not included, as they were too large for the scale of the graphs. However, suffice it to say that no statistical significance was detected between points.

Appendix H.1: Line graph for Emdogain® batch 1

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Appendix H.2: Line graph for Emdogain® batch 2

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Appendix H.3: Line graph for Emdogain® batch 4

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Appendix H.4: Line graph for Emdogain® batch 5

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