Host-Pathogen Interactions of Porphyromonas Gingivalis

Host-Pathogen Interactions of Porphyromonas gingivalis

Jiamin Aw

ORCID 0000-0002-0973-9376

Submitted in total fulfilment of the requirement of the degree of Doctor of Philosophy

April 2018

Melbourne Dental School

The University of Melbourne

Declaration

This is to certify that:

(i) the thesis comprises only my original work towards the PhD, except when indicated in the Preface, (ii) due acknowledgment has been made in the text to all other material used, (iii) the thesis is fewer than 100,000 words in length, exclusive of tables, maps, bibliographies and appendices.

Jiamin Aw Melbourne Dental School The University of Melbourne April 2018

Preface

In accordance with the regulations of The University of Melbourne, I acknowledge that some of the work presented in this thesis was collaborative. Specifically:

(i) In Chapter 6, cryo-electron microscope images of P. gingivalis W50 and P. gingivalis ΔPG0382 were acquired by Dr. Yu-Yen Chen (The University of Melbourne).

The remainder of this thesis comprises only my original work.

Publications

The work presented in this thesis has given rise to the following publication:

Aw, J., Scholz, G.M., Huq, N.L., Huynh, J., O’Brien‐Simpson, N.M., Reynolds, E.C. (2018), “Interplay between Porphyromonas gingivalis and EGF signalling in the regulation of CXCL14”, Cellular Microbiology, e12837

The work done during my PhD also contributed to the following publications:

Scholz, G.M., Heath, J.E., Aw, J. and Reynolds, E.C. (2018), “ Regulation of the petidoglycan amidase PGLYRP2 in epithelial cells by IL-36”, Infection and Immunity, doi: 10.1128/IAI.00384-18

Huynh, J., Scholz, G.M., Aw, J. and Reynolds, E.C. (2017), “Interferon Regulatory Factor 6 Promotes Keratinocyte Differentiation in Response to Porphyromonas gingivalis”, Infection and Immunity, Vol. 85 No. 5, pp. 1-12.

Huynh, J., Scholz, G.M., Aw, J., Kwa, M.Q., Achuthan, A., Hamilton, J.A. and Reynolds, E.C. (2016), “IRF6 Regulates the Expression of IL-36 by Human Oral Epithelial Cells in Response to Porphyromonas gingivalis”, The Journal of Immunology, Vol. 196 No. 5, pp. 2230–2238.

Kwa, M.Q., Huynh, J., Aw, J., Zhang, L., Nguyen, T., Reynolds, E.C., Sweet, M.J., Hamilton, J.A., Scholz, G.M. (2014), “Receptor-interacting protein kinase 4 and interferon regulatory factor 6 function as a signalling axis to regulate keratinocyte differentiation”, Journal of Biological Chemistry, Vol. 289 No. 45, pp. 31077-31087.

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Abstract

Periodontal health is supported by various host immune defence mechanisms, which act in concert to maintain host-microbe homeostasis. However, breakdown of homeostasis can lead to the development of chronic periodontitis, an inflammatory disease that causes the destruction of periodontal tissues. Pattern recognition receptors, including Toll-like receptors (TLRs), enable the detection of microorganisms and subsequent activation of the host immune response. The modular intracellular Toll/Interleukin-1 receptor (TIR) domain of TLRs forms heterotypic interactions with intracellular adaptor proteins, such as MAL and MYD88, to activate downstream signalling pathways to regulate the transcription of inflammatory genes (e.g. cytokines and chemokines). P. gingivalis is a major periodontal pathogen and can disrupt homeostasis between the host and tooth-accreted subgingival biofilm (plaque) by dysregulating the immune response. The extracellular gingipain proteases (Kgp and RgpA/B) produced by P. gingivalis are potent virulence factors, which can stimulate as well as proteolytically degrade host immunomodulatory factors. Consequently, an otherwise host-protective immune response can become destructive and cause tissue pathology when dysregulated by P. gingivalis.

This thesis has characterised two facets of the interaction between the host and P. gingivalis. The first part of this thesis investigated the molecular regulation of the orphan chemokine, CXCL14, by P. gingivalis in oral epithelial cells (e.g. OKF6 cells). By using an isogenic P. gingivalis gingipain (Kgp/Rgp) protease-deficient mutant and a cysteine protease inhibitor, the stimulation of CXCL14 expression was shown to be mediated by the gingipain proteases. Given this finding, a role for protease-activated receptors (PARs) in the regulation of CXCL14 expression was investigated. Gene silencing experiments revealed that P. gingivalis-stimulated CXCL14 expression occurs in a PAR-3-dependent manner. Notably, CXCL14 expression was found to be transcriptionally repressed in response to epidermal growth factor-induced activation of the MEK-ERK1/2 pathway. However, P. gingivalis can overcome the repression of CXCL14 via the gingipain protease-mediated degradation of EGF. Therefore, P. gingivalis not only directly stimulates CXCL14 expression via PAR-3, but also promotes its expression by antagonising EGF signalling.

The functions of CXCL14, including its ability to regulate inflammatory gene expression and oral epithelial cell migration, were also investigated. No evidence was obtained to indicate that CXCL14 regulates inflammatory gene expression in oral epithelial cells (e.g. OKF6 cells) or macrophages (e.g. RAW 264.7 cells), or oral epithelial cell migration. Furthermore, CXCL14 did not induce changes in the expression levels of inflammatory genes when injected into the mouse gingiva. However, CXCL14 was shown in vitro to potently kill oral Streptococcus species.

Significantly, P. gingivalis was resistant to CXCL14 killing, most likely because the gingipain proteases can degrade CXCL14. Therefore, the dysregulation of CXCL14 by P. gingivalis may potentially destabilise the proportions of bacterial species in the tooth-accreted biofilm and thereby promote biofilm dysbiosis.

In the second part of this thesis, the expression and function of TIR domain-containing proteins (Tcps) by P. gingivalis was investigated. Other bacterial pathogens have been shown to express Tcps as a mechanism of molecular mimicry to subvert TLR-mediated immune responses. Bioinformatics analysis identified eleven putative Tcps (e.g. PG0382) in nine different strains. Further analyses revealed that the putative TIR domain in the C-terminal half of PG0382 contains sequence features similar to TLR adaptor proteins and bacterial Tcps. Homology modelling of PG0382 suggests that the domain may adopt a TIR-like structure. Like other bacterial Tcps, PG0382 is predicted to also contain a coiled-coil motif in the N-terminal half of the protein, which may facilitate homodimerisation. Functional characterisation of PG0382 by transient expression in human HEK293T cells and analysis by Western blotting and immunofluorescence confocal microscopy revealed that the N-terminal half of PG0382 appears to largely dictate its subcellular localisation. Moreover, co-expression of PG0382 strongly reduced MAL and MYD88 protein levels. Therefore, P. gingivalis PG0382 may have potential immunomodulatory functions.

An isogenic P. gingivalis PG0382-deficient mutant (P. gingivalis ΔPG0382) was created to establish a role for PG0382 in modulating the host immune response to P. gingivalis. Phenotypic characterisation of the mutant indicates that PG0382 is not important for P. gingivalis growth, formation of an electron-dense surface layer, or gingipain protease activity. The absence of PG0382 did not affect the inflammatory gene response of oral epithelial cells (e.g. OKF6 cells) towards P. gingivalis. However, P. gingivalis ΔPG0382 stimulated a weaker inflammatory response in macrophages (e.g. RAW 264.7 cells). A peritoneal infection model in mice was used to further investigate a role for PG0382 in modulating the host immune response to P. gingivalis. However, the absence of PGO382 did not affect the recruitment of neutrophils and inflammatory monocytes in response to P gingivalis infection. Further studies will be required to determine whether PG0382 has a role in P. gingivalis immune subversion.

This thesis has identified and defined novel interactions between host-derived and P. gingivalis- derived factors in modulating the immune response. Moreover, it provides a molecular basis for exploring potential roles for CXCL14 and P. gingivalis PG0382 in the development and progression of chronic periodontitis.

Acknowledgements This thesis is the culmination of my PhD journey, which would not have been possible or as enjoyable without the support and encouragement of numerous people. I would like to start by expressing my deepest gratitude to my supervisor Associate Professor Glen Scholz, whose enthusiasm for scientific research knows no bounds. His unwavering belief in my abilities encouraged me to challenge myself as a researcher. Without his patience and knowledge, I would not have been able to achieve all that I have over the past few years. I would also like to extend my thanks to my co-supervisor Professor Neil O’ Brien-Simpson for his thought-provoking discussions. A special thanks to Professor Eric Reynolds for giving me the opportunity to undertake my PhD under the Oral Health CRC.

My sincerest thanks go to my colleagues at the Melbourne Dental School who have offered their valued technical support and advice. I have greatly benefited the support of Christine Seers, who helped me devise a strategy for creating the P. gingivalis PG0382-deficient mutant. I would like to thank Yu-Yen Chen for acquiring the cryo-EM images of P. gingivalis. I would also like to express my gratitude towards Katrina Walsh and Alexis Gonzalez who have shared their insightful knowledge about flow cytometry. Thank you to Dhana Gorasia, Michelle Glew and Dina Chen for their technical advice in the lab, and also to Laila Huq, who have brightened up the student room with her cheeriness. A special thanks to Jacqueline Heath, for the time we have spent at the Bio21 BRF developing a mouse model, which seemed impossible at the time. And also, for your insightful knowledge into life and research; your passion and determination is truly inspirational.

I am also thankful to have met some wonderful friends over the past few years. I am especially grateful to Mei Qi Kwa and Jennifer Huynh, who not only shown me the ropes in the lab when I first started in my Honours, but also shared their contagious love of research (and cute animals) with me. I would also like to thank Li-Ming Bhutta and Ben Huang for their support and encouragement throughout this journey.

Importantly, this work would not have been possible without the support of my family. Mummy and Baba, I wish to thank you for supporting my pursuit in this quest. You have both given me the courage and belief to conquer the impossible. Last but not least, to my best friend and partner, John Huang. Thank you for being extraordinarily tolerant and supportive. You were my pillar of strength when I faltered; this journey would not have been completed without you.

Table of Contents

Declaration ...... i Preface ...... ii Publications ...... iii Abstract ...... iv Acknowledgements ...... vi List of Figures ...... xii List of Tables ...... xiv Abbreviations ...... xv Chapter 1 Introduction ...... 1 1.1 Overview of the Oral Microbial Habitat ...... 2 Formation of the Oral Biofilm...... 2 1.2 Periodontal Disease ...... 5 Overview ...... 5 Bacterial Aetiology of Chronic Periodontitis ...... 5 Periodontal Disease and Systemic Health ...... 7 1.3 Host Immune Response in Periodontitis ...... 8 Innate Immune Response in Periodontitis ...... 8 1.3.1.1 Oral Epithelial Cells ...... 9 1.3.1.2 Neutrophils ...... 10 1.3.1.3 Macrophages ...... 12 1.3.1.4 Dendritic Cells ...... 13 Adaptive Immune Response in Periodontitis ...... 14 1.3.2.1 T Lymphocytes ...... 14 1.3.2.2 B Lymphocytes ...... 15 Molecular Mediators of Host Immunity ...... 16 1.3.3.1 Complement System ...... 16 1.3.3.2 Antimicrobial Mediators ...... 17 1.3.3.3 Cytokines ...... 19 1.3.3.4 Chemokines ...... 21 1.4 Pattern Recognition Receptors ...... 23 Toll-like Receptors ...... 24 1.4.1.1 TLR Distribution and Subcellular Localisation ...... 24 1.4.1.2 TLR Structural Organisation ...... 25

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Leucine-Rich Repeat Motif ...... 25 Toll/Interleukin-1 Receptor (TIR) Domain ...... 27 1.4.1.3 Toll-like Receptor Signal Transduction...... 28 1.4.1.4 TIR domain-containing Adaptor Proteins ...... 31 Myeloid Differentiation Primary Response 88 (MYD88) ...... 31 MYD88 Adaptor-Like (MAL) ...... 31 TIR-domain-containing Adaptor-inducing Interferon- (TRIF) ...... 32 TRIF-related Adaptor Molecule (TRAM) ...... 32 Sterile Alpha and Armadillo-motif-containing (SARM) ...... 32 NOD-like Receptors ...... 33 Protease-activated Receptors ...... 37 1.5 Microbial Subversion of TLR Signalling ...... 38 PAMP Modification ...... 38 Targeting TLR Signalling Proteins ...... 39 1.6 Bacterial TIR Domain-containing Proteins ...... 40 Structural Properties of Bacterial Tcps ...... 41 Interference with TLR signalling by Bacterial Tcps ...... 41 Functional Consequences of Bacterial Tcps ...... 42 1.7 Immune Subversion by Porphyromonas gingivalis ...... 42 P. gingivalis Gingipain Proteases ...... 43 1.7.1.1 Gingipain Proteases and Immune Subversion ...... 43 1.7.1.2 Gingipain Proteases and Tissue Destruction ...... 44 Fimbriae ...... 45 Atypical Lipopolysaccharides ...... 46 SerB Phosphatase ...... 47 1.8 Research Objectives ...... 47 Chapter 2 Materials and Methods ...... 48 2.1 Materials ...... 49 Tissue culture reagents ...... 49 Bacterial culture reagents ...... 49 General reagents and chemicals ...... 49 Molecular biology reagents ...... 49 Molecular cloning ...... 49 Quantitative real-time PCR probes...... 50 SDS-PAGE and Western Blotting ...... 50

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Antibodies ...... 50 2.2 In vitro methods ...... 51 Cell culture...... 51 2.2.1.1 OKF6/TERT-2 cells ...... 51 2.2.1.2 HEK293T cells...... 51 2.2.1.3 RAW 264.7 cells ...... 51 Bacterial strains and culture conditions ...... 51 2.2.2.1 P. gingivalis ...... 51 2.2.2.2 Streptococcus strains ...... 52 2.2.2.3 E. coli ...... 52 Challenging of OKF6 cells and RAW264.7 cells with P. gingivalis ...... 52 RNA interference-mediated gene silencing ...... 52 RNA purification ...... 52 Reverse transcription ...... 53 Quantitative real-time PCR ...... 53 CXCL14 Enzyme-linked immunosorbent assays (ELISA) ...... 53 In vitro proteolytic degradation of EGF and CXCL14...... 54 LTQ Orbitrap Elite mass spectrometry ...... 54 Antibacterial assays ...... 55 Wound healing assay ...... 55 Transfection of HEK293T cells ...... 55 Cell lysis ...... 55 Co-immunoprecipitation Assay ...... 55 SDS-PAGE ...... 56 Western Blotting ...... 56 Immunofluorescence staining and confocal microscopy ...... 56 Construction of PG0382 mammalian expression vectors...... 57 2.2.19.1 Polymerase Chain Reaction ...... 57 2.2.19.2 Bacterial transformation ...... 57 Generation of P. gingivalis PG0382-deficient mutant (ΔPG0382) ...... 58 2.2.20.1 Splice Overlap Extension Polymerase Chain Reaction (SOE PCR) ...... 58 2.2.20.2 Electroporation of P. gingivalis ...... 58 Gingipain proteinase assay ...... 59 2.3 Bioinformatics methods ...... 59 Sequence alignments...... 59

Structural modelling...... 59 2.4 Mouse studies ...... 59 Mice ...... 59 Intragingival injection ...... 60 Mouse gingival RNA extraction ...... 60 Intraperitoneal infection ...... 60 Flow cytometric analysis ...... 61 2.5 Statistical analysis ...... 61 Chapter 3 The regulation of CXCL14 in oral epithelial cells by Porphyromonas gingivalis...... 68 3.1 Introduction ...... 69 3.2 Results...... 70 P. gingivalis stimulates CXCL14 expression in human oral epithelial cells in a TLR2-independent manner ...... 70 Gingipain proteases mediate the stimulation of CXCL14 expression by P. gingivalis ...... 73 PAR-3-dependent regulation of P. gingivalis-stimulated CXCL14 gene expression ...... 75 EGFR signalling negatively regulates CXCL14 transcription in a MEK-dependent manner ...... 78 P. gingivalis gingipain proteases antagonise the negative regulation of CXCL14 gene expression by EGF ...... 82 3.3 Discussion ...... 84 Chapter 4 Function of CXCL14 ...... 90 4.1 Introduction ...... 91 4.2 Results...... 92 Effects of CXCL14 on inflammatory gene expression in macrophages ...... 92 Effects of CXCL14 on inflammatory gene expression in oral epithelial cells ...... 93 Effects of CXCL14 on inflammatory gene expression in the mouse gingiva ...... 95 Effect of CXCL14 on oral epithelial cell migration ...... 97 Bactericidal activity of CXCL14 against oral bacteria ...... 99 Gingipain protease-dependent degradation of CXCL14 by P. gingivalis ...... 101 Identification of CXCL14 peptides resulting from Kgp digestion ...... 102 4.3 Discussion ...... 103 Chapter 5 Identification and characterisation of P. gingivalis TIR domain-containing proteins ...... 109 5.1 Introduction ...... 110 5.2 Results...... 110

Identification of putative P. gingivalis TIR domain-containing proteins ...... 110 PG0382 is predicted to contain a coiled-coil motif ...... 116 Comparison of PG0382 with mammalian TLRs and adaptor proteins ...... 117 Comparison of PG0382 with bacterial Tcps ...... 121 Structural prediction of PG0382 ...... 123 Expression of PG0382 in HEK293T cells ...... 125 PG0382 expression causes the loss of MAL and MYD88 in HEK293T cells ...... 130 Lack of complex formation between PG0382TD and MAL...... 134 Subcellular localisation of PG0382TD, MAL and MYD88 ...... 135 5.3 Discussion ...... 137 Chapter 6 Investigation of PG0382 and host inflammation ...... 141 6.1 Introduction ...... 142 6.2 Results...... 142 Generation of an isogenic P. gingivalis PG0382-deficient mutant ...... 142 Phenotypic characterisation of P. gingivalis ΔPG0382...... 146 Gingipain protease activity of P. gingivalis ΔPG0382 ...... 148 Stimulation of inflammatory gene responses in oral epithelial cells by P. gingivalis ΔPG0382 ...... 149 Stimulation of inflammatory gene responses in macrophages by P. gingivalis ΔPG0382 ...... 150 Innate immune response to P. gingivalis ΔPG0382 in mice ...... 151 6.3 Discussion ...... 159 Chapter 7 General Discussion ...... 163 7.1 Summary ...... 164 7.2 Implications of a dysregulated CXCL14 response for chronic inflammation and microbial dysbiosis ...... 164 7.3 CXCL14 in tissue regeneration ...... 167 7.4 A potential role for PG0382 in immune subversion ...... 167 7.5 The TIR domain as a primordial microbial signalling module ...... 169 7.6 Bacterial Tcps as therapeutic agents for inflammation...... 170 7.7 Conclusion ...... 171 Bibliography ...... 172 Appendix ...... 204

List of Figures Figure 1.1 The effects of subgingival plaque on chronic periodontitis...... 4 Figure 1.2 Structural arrangement of Toll-like receptors...... 26 Figure 1.3 Crystal structure of the TLR3 LRR motif...... 27 Figure 1.4 Crystal structure of the TLR1 and TLR2 TIR domains...... 28 Figure 1.5 TLR signalling pathways...... 30 Figure 1.6 Structural arrangement of TIR domain-containing adaptor proteins...... 33 Figure 1.7 Structural arrangement of NOD-like receptors...... 35 Figure 1.8 NOD-like receptor signalling pathways...... 36 Figure 3.1 P. gingivalis stimulates CXCL14 gene expression in a TLR2-independent manner. .... 71 Figure 3.2 P. gingivalis stimulates CXCL14 gene expression in an IRAK-1 and IRF6-independent manner...... 72 Figure 3.3 P. gingivalis gingipain proteases stimulate CXCL14 expression...... 74 Figure 3.4 P. gingivalis stimulates PAR-2 gene expression in oral epithelial cells...... 75 Figure 3.5 PAR-3-dependent regulation of P. gingivalis-inducible CXCL14 expression...... 77 Figure 3.6 EGF differentially regulates cytokine expression...... 79 Figure 3.7 EGF suppresses CXCL14 transcription via MEK signalling...... 81 Figure 3.8 P. gingivalis antagonises the regulation of CXCL14 by EGF...... 83 Figure 3.9 A proposed model for the regulation of CXCL14 gene expression in oral epithelial cells by P. gingivalis...... 88 Figure 4.1 CXCL14 stimulation of mouse macrophage RAW 264.7 cells...... 93 Figure 4.2 CXCL14 stimulation of OKF6 cells...... 94 Figure 4.3 Effects of CXCL14 on inflammatory gene expression in the mouse gingiva...... 96 Figure 4.4 Effect of CXCL14 on OKF6 cell migration...... 98 Figure 4.5 Bactericidal activity of CXCL14...... 100 Figure 4.6 Gingipain protease-mediated degradation of CXCL14...... 101 Figure 5.1 Phylogenetic tree of P. gingivalis TIR domain-containing proteins...... 112 Figure 5.2 Predicted protein domains of annotated P. gingivalis TIR domain-containing proteins...... 115 Figure 5.3 COILS analysis output for PG0382 ...... 116 Figure 5.4 Amino acid sequence of PG0382...... 117 Figure 5.5 Multiple sequence alignment of PG0382 TIR domain with TLR TIR domains...... 119 Figure 5.6 Multiple sequence alignment of PG0382 TIR domain with TLR adaptor proteins TIR domains...... 120 Figure 5.7 Multiple sequence alignment of PG0382 TIR domain with TIR domains of known bacterial TIR domain-containing proteins...... 122

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Figure 5.8 PG0382 structural prediction...... 124 Figure 5.9 Construction of pEF-V5-PG0382 and pEF-V5-PG0382ΔTD expression vectors...... 126 Figure 5.10 Ectopic expression of V5-PG0382, V5-PG0382ΔTD, and V5-PG0382TD in HEK293T cells...... 128 Figure 5.11 Subcellular localisation of ectopically expressed PG0382, PG0382ΔTD, and PG0382TD in HEK293T cells...... 129 Figure 5.12 Effects of PG0382 on MAL and MYD88 expression...... 131 Figure 5.13 Effects of PG0382 on MAL and MYD88 mRNA levels in HEK293T cells...... 132 Figure 5.14 Concentration-dependent effects of PG0382 on MAL and MYD88 protein expression...... 133 Figure 5.15 Analysis of PG0382TD and MAL interaction by co-immunoprecipitation...... 134 Figure 5.16 Co-localisation of PG0382TD with MAL and MYD88 by immunofluorescence...... 136 Figure 6.1 Strategy for the generation of an isogenic P. gingivalis PG0382-deficient mutant. ... 144 Figure 6.2 Gel electrophoresis analysis of PCR products...... 145 Figure 6.3 Growth rates of wildtype P. gingivalis and P. gingivalis ΔPG0382...... 146 Figure 6.4 Phenotypic characterisation of P. gingivalis ΔPG0382...... 147 Figure 6.5 P. gingivalis gingipain protease activity...... 148 Figure 6.6 Effects of P. gingivalis ΔPG0382 on inflammatory responses of oral epithelial cells.149 Figure 6.7 Effects of P. gingivalis ΔPG0382 on inflammatory cytokine responses of macrophages...... 151 Figure 6.8 FACs Gating strategy for identifying innate immune cells...... 152 Figure 6.9 Activation of macrophages in response to P. gingivalis...... 154 Figure 6.10 Recruitment of inflammatory monocytes in response to P. gingivalis...... 156 Figure 6.11 Recruitment of neutrophils in response to P. gingivalis...... 158

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List of Tables Table 1.1 Human antimicrobial mediators of host defence...... 19 Table 1.2 Roles of major groups of cytokines in host defence...... 21 Table 1.3 Roles of chemokines in host defence...... 23 Table 1.4 Toll-like receptor subcellular localisation and ligands...... 24 Table 1.5 PARs activating proteases, cellular expression and key functions...... 38 Table 1.6 Bacterial Tcps and immune subversion...... 40 Table 2.1 List of plasmids used in this study...... 62 Table 2.2 PCR primers used for constructing PG0382 mammalian expression vectors...... 63 Table 2.3 PCR primers used to generate PCR products for constructing ermF cassette...... 64 Table 2.4 SOE PCR reaction thermal cycling conditions...... 66 Table 2.5 Flow cytometric analysis antibody cocktails...... 67 Table 4.1 CXCL14 peptides identified by MS from Kgp digestion...... 102 Table 5.1 Annotated TIR domain-containing proteins of P. gingivalis strains...... 111 Table 5.2 Amino acid sequence identity (%) between P. gingivalis TIR domain-containing proteins...... 113 Table 5.3 Amino acid sequence identity (%) between P. gingivalis TIR domain-containing proteins TIR domains...... 114 Table 5.4 Amino acid sequence identity (%) between PG0382 TIR domain and TLR TIR domains...... 118 Table 5.5 Amino acid sequence identity (%) between PG0382 TIR domain and TIR domains of TLR protein adaptors...... 120 Table 5.6 Amino acid sequence identity (%) between PG0382 TIR domain and bacterial Tcp TIR domains...... 121 Table 5.7 Identification of PG0382 TIR domain structural homologs using FUGUE...... 123

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Abbreviations

AP-1 Activator protein 1

BCR B cell receptor

BLAST Basic Local Alignment Search Tool bFGF Basic fibroblast growth factor

BPE Bovine pituitary extract cAMP Cyclic adenosine monophosphate

CCL C-C motif ligand

CCR C-C motif receptor

CRP C-reactive protein

Cryo-EM Cryo-electron microscopy

CXCL C-X-C motif ligand

ECL Enhanced chemiluminescence

EDSL Electron surface dense layer

EGF Epidermal growth factor

EGFR Epidermal growth factor receptor

FBS Foetal bovine serum

FcR Fc receptor

G-CSF Granulocyte-colony stimulating factor

HSP Heat shock protein

ICAM Intracellular adhesion molecule

IFN Interferon

IGF-1 Insulin-like growth factor

IKK IκB kinase

IAPs Inhibitor of apoptosis

IL Interleukin

IL-1Ra IL-1 receptor antagonist

IL-1RacP IL-1 receptor accessory protein

IRAK Interleukin-1 receptor-associated kinase

IRF Interferon regulatory factor

KGF Keratinocyte growth factor

Kgp Lysine-specific gingipain

LPS Lipopolysaccharide

LRR Leucine-rich repeat

MAPK Mitogen-activated protein kinase

MCP-1 Monocyte chemotactic protein-1

MMP Matrix metalloproteinase

MAL Myeloid differentiation primary response adaptor-like

MYD88 Myeloid differentiation primary response 88

NETs Neutrophil extracellular traps

NF-κB Nuclear factor kappa B

NK Natural killer

NLR NOD-like receptor

NO Nitric oxide

NDP Nucleotide

NTPase Nucleoside-triphosphatase

NTP Nucleoside triphosphate

PAD Peptidyl-arginine deiminase

PAMP Pathogen-associated molecular pattern

PDGF Platelet-derived growth factor

PI3K Phosphoinositide 3-kinase

PKA Protein kinase A

Poly(I:C) Polyinosinic:polycytidylic acid

PRR Pattern recognition receptor

RANKL Nuclear factor κ-B ligand

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RANTES Regulated on activation, normal T cell expressed and secreted

Rgp Arginine-specific gingipain

RIP Receptor-interacting serine/threonine-protein kinase

RLR RIG-I-like receptors

SARM Sterile α and TIR motif containing protein

SFD-1 Stromal-derived factor-1

SMURF1 SMAD specific E3 ubiquitin protein ligase 1

STAT Signal transducer and activator of transcription

TAB TAK-1 associated binding protein

TAK Transforming growth factor-β activated kinase

TBK TANK-binding kinase

Tcp TIR domain-containing protein

TCR T cell receptor

TERT-2 Telomerase reverse transcriptase 2

TGF Transforming growth factor

TIR Toll/Interleukin-1 receptor domain

TIRAP Toll-interleukin 1 receptor domain-containing adaptor protein

TLCK N--Toysl-L-Lysine chloromethyl ketone hydrochloride

TLR Toll-like receptor

TNF Tumour necrosis factor

TNFR Tumour necrosis factor receptor

TRAF TNF receptor-associated factor

TRAM TRIF-related adaptor molecule

TRIF TIR-domain-containing adaptor-inducing interferon-β

VCAM Vascular cell adhesion molecule

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Introduction

1.1 Overview of the Oral Microbial Habitat The mucosal surfaces of the human body are inhabited by a myriad of microorganisms, and the oral cavity provides an excellent niche for the establishment of unique microbial communities. The mucosal surfaces of the oral cavity (e.g. gingiva, tongue, palate and cheek) and the non-shedding surface of the teeth provide distinct habitats to sustain the growth of different microbial communities that form biofilms. A combination of environmental (e.g. nutrient availability, pH and temperature) and host-derived (e.g. antimicrobial peptides) factors create an ecological niche that dictates the distribution of bacterial species at various sites in the oral cavity (Carlsson, 1997). The ecological properties of the oral cavity are complex, dynamic and highly variable in individuals. Normal physiology, genetics and lifestyle factors (e.g. diet) can influence the composition of these microbial ecosystems. In health, a symbiotic relationship is maintained between the host and oral commensal species; microbial inhabitants partake an active role in shaping and maintaining health by acting as antigenic stimulants to facilitate homeostatic immunity (Marsh and Devine, 2011). However, changes in the microenvironment can lead to the proliferation of pathogens or potential pathogens (pathobionts) within the biofilm and cause disease (Hajishengallis and Lamont, 2012).

Formation of the Oral Biofilm As indicated above, microorganisms can form organised and highly dynamic communities on biotic and abiotic substrates, known as biofilms (O’Toole et al., 2000). The mucosal and hard surfaces of the oral cavity are favourable sites for biofilm formation. Microorganisms within oral biofilms are constantly exposed to environmental changes, such as pH fluctuation, nutrient availability and host-produced enzymes (e.g. lysozyme) (Carlsson, 1997; O’Toole et al., 2000). The formation of a biofilm allows participating microorganisms to be more resistant to physical and biochemical forces within the oral cavity. Over time, the biofilm develops into a coordinated microbial community, which during health shares a homeostatic relationship with the host.

The oral microbiome is a consortium of over 700 microbial species that make up different dynamic polymicrobial communities (Aas et al., 2005). The microenvironment dictates the composition of microbial species within biofilms on the teeth, tongue, and the buccal, upper and lower mucosal surfaces within the oral cavity (Aas et al., 2005). Supragingival plaque occupies the smooth surface of the teeth, whilst subgingival plaque is localised to the gingival crevice along the tooth enamel and the sulcular epithelium (Fig. 1.1). The oxygenated environment around the teeth benefits the growth of aerobic bacterial species within supragingival plaque (e.g. Lactobacillus and Actinomyces species). Contrastingly, subgingival plaque is predominantly comprised of facultative and strict anaerobic bacterial species (e.g. Fusobacterium nucleatum, Porphyromonas gingivalis and Prevotella intermedia) (Zijnge et al., 2010).

The formation of the tooth-accreted biofilm is a sequential and organised process involving various microbial species (Kolenbrander et al., 2010; O’Toole et al., 2000). The initial primary colonisers are the pioneering bacterial species that interact with host proteins to build a microbial monolayer. Gram-positive bacterial species, including Streptococci and Actinomyces species (e.g. Streptococcus gordonii and Actinomyces naeslundii), form the oral microbiota of subgingival plaque (Yao et al., 2003; Palmer et al., 2003; Li et al., 2004; Dige et al., 2009). The early colonisers express adhesins (e.g. antigen I/II polypeptides and amylase-binding protein produced by oral Streptococci) to form attachments to the gingival epithelium (O’Toole et al., 2000; Rogers et al., 2001). Co-aggregation between primary colonisers and other bacterial species that cannot directly colonise the gingival surface is essential for biofilm formation. F. nucleatum acts as a central player in facilitating biofilm formation by promoting co-aggregation (Al-Ahmad et al., 2007). The F. nucleatum radD adhesin can interact with primary colonising species, such as Streptococcus sanguinis and A. naeslundii, to facilitate mixed- species biofilm formation (Kaplan et al., 2009). In addition, co-aggregation-mediated interactions facilitated by F. nucleatum promotes the survival of late colonising obligate anaerobes in oxygenated environments (Bradshaw et al., 1998). P. gingivalis, Treponema denticola and Tanerella forsythia are late colonising species that are commonly associated with chronic periodontitis (Socransky et al., 1998). In collaboration, bacterial species within oral biofilms therefore form highly organised and structured communities.

Figure 1.1 The effects of subgingival plaque on chronic periodontitis. Subgingival plaque is localised to the gingival crevice along the tooth enamel and the sulcular epithelium. The junctional epithelium is the region where the epithelium connects to the tooth and is highly susceptible to bacterial invasion, therefore it is maintained by a high level of neutrophil infiltration. A normal balance between the oral commensal and host immune response is required to maintain host-biofilm homeostasis in the healthy periodontium (left). The accumulation of subgingival plaque (right) can induce chronic periodontitis and results in the destruction of the epithelial tissue and alveolar bone resorption. Figure taken from (Darveau, 2010).

1.2 Periodontal Disease Overview Periodontal disease is a spectrum of inflammatory conditions that are associated with the tissues supporting the teeth, otherwise known as the periodontium. Periodontal conditions can be categorised as: (i) gingivitis, (ii) chronic periodontitis, and (iii) aggressive periodontitis (Highfield, 2009). Gingivitis is associated with mild inflammation of the periodontium without the loss of connective tissue. Gingivitis can transit to chronic periodontitis, where there is a breakdown of gingival tissue, periodontal ligament detachment, and in severe cases, alveolar bone resorption. Aggressive periodontitis shares similar characteristics with chronic periodontitis, with the exception of a faster rate of disease progression and occurs more frequently in juveniles (Highfield, 2009).

Epidemiological studies suggest that there are potential links between chronic periodontitis and various systemic conditions, such as cardiovascular disease and rheumatoid arthritis (Seymour et al., 2007). Although the nature of the association of chronic periodontitis with these other important diseases are yet to be fully determined, it is thought that the continual activation of the immune system, instigated by subgingival biofilm, compromises the immune system and leads to the destruction of host tissues.

Bacterial Aetiology of Chronic Periodontitis Chronic periodontitis is a complex, multifactorial disease that can be attributed to lifestyle (e.g. diet and smoking) and genetic factors (e.g. familial neutropenia) (Pihlstrom et al., 2005). The oral biofilm is also a predominant driver in the development and progression of chronic periodontitis. Numerous studies have documented qualitative and quantitative changes in oral biofilm (plaque) composition in the transition from health to disease (Hong et al., 2015; Marsh, 1994; Paster et al., 2001; Socransky et al., 1998). The conception of different hypotheses for the development of periodontitis has evolved over the years. However, each hypothesis highlights the importance of host-biofilm homeostasis to maintain periodontal health.

The Non-Specific Plaque Hypothesis was developed to describe a correlation between plaque accumulation and disease development. It was believed that an increase in bacterial load in plaque exceeded a containment threshold, and thus initiated disease onset (Theilade, 1986). Therefore, disease prevention was focused on maintenance of oral hygiene and the physical removal of plaque (e.g. tooth brushing and scaling). The subsequent isolation and identification of disease-associated microorganisms led to the Specific Plaque Hypothesis, in which specific bacterial species in plaque were associated with either health or disease (Loesche, 1979; Loesche et al., 1977). A landmark study by Socransky et al. showed that specific bacterial

5 species could be grouped into six clusters based on their association with health and disease. Of the six clusters, the “red complex”, comprised of P. gingivalis, T. forsythia and T. denticola, was most frequently associated with disease (Socransky et al., 1998). A number of recent studies have also demonstrated, in a more comprehensive and detailed manner, significant differences in oral microbiota composition between periodontal health and disease (Griffen et al. 2012; Abusleme et al., 2013; Hong et al., 2015). Indeed, advances in DNA sequencing and bioinformatics technologies have provided greater insight into the bacterial aetiology of chronic periodontitis by enabling analysis of microbial community composition. In addition to the “red complex” bacterial species, these more recent DNA-based sequencing studies suggest that bacterial species from the Spirochaetes and Synergistetes taxa are also likely to be associated with the pathogenesis of chronic periodontitis (Griffen et al., 2012; Abusleme et al., 2013). As such, it has been proposed that antibiotic treatment against specific bacterial species could be employed to treat chronic periodontitis (Slots & Rams, 1990; Cionca et al., 2009). However, a conservative and highly selective approach would need to be undertaken to avoid the possible selection and overgrowth of antibiotic-resistant pathogens (Slots and Ting, 2002).

The Ecological Plaque Hypothesis, proposed by Marsh and colleagues, emphasised the interplay between the oral microenvironment and microflora in disease development and progression (Marsh, 1994, 2003). Changes in ecological factors (e.g. pH and nutrients) can select for the bacterial species composition of the oral biofilm. For instance, the consumption of high sugar content foods increases the growth of acid-tolerant bacterial species, such as Streptococcus mutans, which can readily metabolise dietary sugars to acid (Becker et al., 2002). Similarly, metabolic activity from plaque microbiota can also alter the environment and contribute to host-biofilm destabilisation. Facultative anaerobes, such as F. nucleatum, can metabolise oxygen to create a reducing microenvironment that assists the colonisation and growth of strict anaerobes, including P. gingivalis (Diaz et al., 2002). Thus, the environment and microbial growth are co-dependent, and disease can ensue when one or the other is disrupted.

The Polymicrobial Synergy and Dysbiosis hypothesis is an extension of the Ecological Plaque Hypothesis, and emphasises the synergistic contribution of microorganisms in dysbiosis to disease development. The model describes a role for keystone pathogens, such as P. gingivalis, in initiating environmental changes that disrupts microbial homeostasis. As a low abundance species, P. gingivalis appears to have a disproportionate effect on plaque microbiota virulence (Hajishengallis et al., 2011). The pathogenicity of P. gingivalis comes from its ability to manipulate the host immune response to subvert defence mechanisms (refer to Section 1.7 for further details). P. gingivalis creates an inflammatory state that facilitates nutrient acquisition, whilst inhibiting host bacterial killing mechanisms to ensure its survival (Hajishengallis et al.,

2012). By impairing the host immune response to the biofilm, P. gingivalis causes an imbalance in microbial distribution and shifts microbiota homeostasis to dysbiosis. As such, microorganisms that are otherwise commensals can act in a synergistic manner to stimulate chronic inflammation and promote disease. The importance of a polymicrobial biofilm in the development of chronic periodontitis was demonstrated in germ-free mice, whereby P. gingivalis failed to promote disease in the absence of other bacterial species (Hajishengallis et al., 2011). Accordingly, the Polymicrobial Synergy and Dysbiosis model highlights the participation of dysbiotic microbial communities in disease pathogenesis and illustrates keystone pathogens as a trigger for dysbiosis. Therefore, therapeutic intervention targeting specific keystone pathogens is a potential avenue to prevent disease development.

Periodontal Disease and Systemic Health Epidemiological studies suggest that there are potential links between chronic periodontitis and various systemic conditions (Genco and Van Dyke, 2010; de Pablo et al., 2009; Seymour et al., 2007). Bacterial and inflammatory factors derived from the oral cavity appear to influence the progression of systemic diseases (Hajishengallis, 2015; Seymour et al., 2007), whereby periodontal pathogens or inflammatory by-products can enter the systemic circulation and act as stimulatory factors that contribute to the development of systemic disease.

Atherosclerosis results from the accumulation of fatty deposits on the vasculature and can lead to various systemic complications (e.g. aneurysms and myocardial infarction). C-reactive protein (CRP), an acute-phase protein produced by the liver, is a classical indicator of systemic inflammation. Increased CRP levels have been detected in patients with chronic periodontitis and ongoing periodontal therapy can successfully return CRP to baseline levels (D’Aiuto et al., 2005; Seinost et al., 2005). Periodontal pathogens can stimulate inflammation in the vasculature to promote atheroma formation. P. gingivalis, which has been detected in atherosclerotic plaques, can cause increased expression of vascular adhesion molecules (e.g. intracellular adhesion molecule-1 (ICAM-1) and vascular cell-adhesion molecule-1 (VCAM-1)) by endothelial cells to promote leukocyte (e.g. monocytes) recruitment and inflammation in the vasculature (Haraszthy et al., 2000; Khlgatian et al., 2002). Significantly, P. gingivalis can stimulate the uptake of low-density lipoprotein by macrophages and foam cell formation, a classical hallmark of atherosclerotic lesions (Qi et al., 2003). Elevated levels of inflammatory mediators (e.g. Interleukin-1 (IL-1, IL-6 and tumour necrosis factor (TNF)), secreted as a result of periodontitis, are also correlated with increased CRP production by the liver (Loos et al., 2000; Noack et al., 2001). Thus, the combination of bacterial dissemination and increased levels of inflammatory cytokines in the systemic circulation may contribute to the risk of exacerbating atherosclerosis.

Periodontitis may also be associated with exacerbating rheumatoid arthritis, an inflammatory condition affecting the underlying structures of bone in synovial tissues. DNA from various oral bacterial species (e.g. P. gingivalis and T. denticola) have been detected in synovial fluid from patients with rheumatoid arthritis (Moen et al., 2006), and some patients suffering from rheumatoid arthritis have been successfully treated with antibiotics (Ogrendik, 2009). The peptidyl-arginine deiminase (PAD) secreted by P. gingivalis can citrullinate host proteins, which may lead to the generation of autoantibodies and hence contribute to the development of rheumatoid arthritis (Koziel et al., 2014; Wegner et al., 2010). In addition, P. gingivalis proteases have also been suggested to enhance tissue breakdown in arthritis by degrading host extracellular matrix proteins, including laminin, fibronectin and collagen (Imamura et al., 2003; Ruggiero et al., 2013).

Associations between poor maternal periodontal health and increased risk of adverse pregnancy outcomes have also been reported (López et al., 2002; Mitchell-Lewis et al., 2001). Inflammatory mediators (e.g. IL-1 and TNF) and/or bacterial endotoxins (e.g. bacterial lipopolysaccharide (LPS)) arising from periodontal infection may enter the systemic circulation and transverse the placental barrier to trigger increased prostaglandin and TNF levels, which might then promote preterm birth (Gibbs, 2001). Women with pregnancy-associated gingivitis who received periodontal treatment during pregnancy had significant reductions in rates of preterm birth and/or low birth weight (Jeffcoat et al., 2001; López et al., 2005). Chronic periodontitis may therefore also contribute to the pathogenesis of systemic disease; however, further studies will be required to confirm causality links, as there are possibly alternate underlying mechanisms that may cause predisposition to some systemic conditions.

1.3 Host Immune Response in Periodontitis Although a controlled immune response is essential to prevent infection, dysbiosis in the oral biofilm can hinder immune-mediated microbial clearance. Consequently, the persistence of immune cells and continual production of inflammatory mediators can contribute to tissue destruction and delay the resolution of inflammation.

Innate Immune Response in Periodontitis The innate immune system is comprised of physical, cellular and bioactive components, which defend the body upon exposure to potential pathogens (e.g. microbes and fungi). As the first point of contact, the oral epithelium acts as a barrier to prevent the breach of microbes. Additionally, oral fluids, such as saliva and gingival crevicular fluid, contain antimicrobial proteins (e.g. lysozyme) that can inhibit the overgrowth of the biofilm. At the molecular level, innate immune cells patrol tissues within the human body to promote the rapid clearance of

8 potential pathogens. Importantly, innate immune cells also initiate the activation of the adaptive immune system through antigen presentation and other mechanisms. The immune system is reinforced by immune mediators, including complement proteins, cytokines, chemokines and antimicrobial proteins, which facilitate the clearance of pathogens. Together, the innate immune system is essential for initiating and establishing host defence against infection.

1.3.1.1 Oral Epithelial Cells The myriad of microorganisms occupying the oral cavity makes it particularly susceptible to infection. The oral cavity, which is covered by keratinised and non-keratinised epithelial cells, depending on anatomical location, acts as a mechanical barrier to bacterial infection (Squier and Kremer, 2001). The regions that are exposed to greatest physical forces (e.g. gingiva and hard palate), such as during mastication, are covered by keratinised epithelium that is tightly attached to the underlying connective tissue. Contrastingly, the soft palate and buccal regions of the oral cavity are covered with non-keratinising epithelium for greater flexibility, which is required for chewing and speech.

The oral mucosa has a stratified architecture characterised by an innermost basal layer that is in direct contact with the underlying basement membrane, comprised of extracellular matrix proteins (e.g. collagen and fibronectin) and growth factors (e.g. transforming growth factor (TGF-). The basal layer is followed by the spinous and granular layers. Keratinised epithelium, such as that covering the hard palate, has an additional keratin layer, which resembles the epidermis of the skin. Cells at the surface of the oral mucosa are maintained through constant replacement from a reservoir of stem cells in the basal layer, which undergo a specialised differentiation program as they migrate towards the superficial layer of the epithelium (Potten and Morris, 1988). Differentiating cells entering the spinous layer undergo a change in the types of keratins expressed. For instance, keratin 5 and 14 are downregulated and keratin 1 and keratin 10 are expressed to form tonofibrils (Fuchs and Green, 1980; Roop et al., 1987). Cells from the spinous layer undergo further differentiation and express filaggrin to facilitate the bundling of keratins into macrofibrils for the formation of a protein scaffold (Dale et al., 1997). Additionally, transglutaminases catalyse the crosslinking of structural envelope proteins (e.g. involucrin and loricrin) to further reinforce the cornified structure (Thacher and Rice, 1985). Thus, a mechanically robust scaffold is established within the cornified cell envelope to provide vital barrier function.

The epithelium is further strengthened by the intercellular linking of proteins at tight junctions and adherens junctions. Tight junctions localised towards the apical surface of the epithelium are maintained by a number of proteins, including claudin, occludin and zonula occludens. They

9 maintain the apico-basal polarity of the epithelial layer and regulate the paracellular passage of ions and water molecules (Kirschner et al., 2013; Yuki et al., 2007). Adherens junctions localised basal to tight junctions provide strong mechanical links between adjacent cells to reinforce epithelial cohesion. Adherens proteins, such as E-cadherin, are anchored to intracellular actin filaments to protect the epithelia from shear forces (Niessen, 2007). Thus, the presence of junctional proteins fortifies the epithelium and limits the transmigration of pathogens. However, the breakdown of proteins in tight and adherens junctions, for example by P. gingivalis and T. denticola proteases, may provide a gateway for bacterial spread deeper into the underlying tissues (Chi et al., 2003; Katz et al., 2002).

The junctional epithelium localised at the bottom of the gingival sulcus is comprised of non-keratinising squamous epithelium consisting of basal and suprabasal layers. Junctional epithelial cells express lower levels of E-cadherin (Hatakeyama et al., 2006), which increases the intercellular space and enhances tissue permeability. Importantly, the more permeable nature of the junctional epithelium permits greater occupation by immune cells, including neutrophils, and thus contributes to a reservoir of neutrophil-associated antimicrobial peptides to promote tissue homeostasis (refer to Section 1.3.1.2 and 1.3.3 for further details).

In addition to acting as a mechanical barrier against microbes, the oral epithelium also forms an immunological front. Antimicrobial molecules (e.g. defensins) are secreted constitutively, and in an inducible manner, as part of the host-protective mechanism against microorganisms in the oral cavity (Dale and Fredericks, 2005) (refer to Section 1.3.3.2 for further details). Importantly, oral epithelial cells also functionally connect the innate immune response to the adaptive immune response during infection. In particular, they express Pattern Recognition Receptors (PRR) that stimulate an immune response upon the recognition of specific microbial patterns (refer to Section 1.4 for further details). PRR activation leads to intracellular signalling cascades that promote the expression of pro-inflammatory genes (e.g. IL-8 and IL-6) to alert and direct the cells of the innate and adaptive immune system to the site of infection. Our laboratory recently undertook a preliminary transcriptomics-based experiment to identify novel P. gingivalis-inducible genes in oral epithelial cells, and one of the genes identified encodes the orphan chemokine, CXCL14. In summary, the oral epithelium not only provides barrier protection, it also produces immunomodulatory factors that recruit and activate immune cells.

1.3.1.2 Neutrophils The junctional epithelium adjacent to the tooth is highly susceptible to microbial invasion (Fig. 1.1), and therefore it is maintained by a high level of neutrophil infiltration (Tsukamoto et

10 al., 2012). The fundamental role of neutrophils is to prevent the establishment of infection by releasing cytotoxic granules to eliminate pathogens. The granules can contain antimicrobial peptides (e.g. -defensins and lipocalin) and/or reactive oxygen metabolites (e.g. nitric oxide and superoxide) (Faurschou and Borregaard, 2003; Rice et al., 1987). In addition, neutrophil extracellular traps (NETs) are comprised of chromatin fibres and bactericidal components (e.g. elastase and myeloperoxidase), which impede bacterial spread and limit secondary damage to surrounding tissues (Cooper et al., 2013).

The close association of chronic periodontitis with defective neutrophil recruitment and function reflects the importance of neutrophils in maintaining periodontal health. Individuals with neutropenia typically have a greater incidence of periodontitis, and treatment with colony-stimulating factor-3 (CSF-3; otherwise known as G-CSF) can rescue neutrophil counts to facilitate bacterial clearance in the periodontium (Deasy et al., 1980; Hastürk et al., 1998). The constitutive transepithelial migration of neutrophils across the junctional epithelium acts as a unified antimicrobial front (Darveau, 2010; Faurschou and Borregaard, 2003). An increasing gradient of ICAM-1 and IL-8 expression from the basal to superficial layers of the junctional epithelium ensures a constant influx of neutrophils at the epithelium-plaque interface (Tonetti et al., 1998). Neutrophils are also actively recruited from the blood to the site of infection during inflammation by the chemokines CXCL2 and CXCL5, as part of the immune defence mechanism (Kolaczkowska and Kubes, 2013; Phillipson et al., 2006). In addition, the distribution of NETs on gingival surfaces and in gingival crevicular exudates also provides host protection by limiting bacterial adherence and invasion of gingival tissues (Krautgartner and Vitkov, 2008; Vitkov et al., 2009). In summary, neutrophils not only provide host defence upon infection, they also play an integral protective role at steady state to maintain periodontal health.

The inflammatory nature of the neutrophil response to the oral biofilm can also destabilise tissue homeostasis and thereby contribute to the onset of periodontitis. Bacterial persistence in the gingival sulcus can lead to the inappropriate recruitment and activation of neutrophils. As a result, neutrophils respond to microbes in the biofilm by releasing tissue-degrading proteins, such as matrix metalloprotease (MMP)-8, MMP-9 and elastase (Shin et al., 2008). The excessive production of these proteins can cause tissue destruction and delay the wound healing process. For instance, the cleavage of platelet derived growth factor (PDGF) by elastase can compromise growth signals in periodontal ligament cells (Nemoto et al., 2005). Consistently, individuals displaying hyperactive neutrophil function, for example, as a result of a gene polymorphism in the neutrophil Fc gammareceptor IIa (FcRIIa), are predisposed to developing severe periodontitis (Nicu et al., 2007). Therefore, the dysregulation of neutrophil responses can contribute to the pathogenesis of periodontitis.

1.3.1.3 Macrophages Macrophages facilitate the inflammatory response by producing inflammatory mediators, and phagocytosing microbes to clear cellular debris to promote the wound healing response. Macrophages can be broadly classified into two major phenotypes: (i) classically-activated M1 macrophages, and (ii) alternatively-activated M2 macrophages. M1 macrophages are involved in inflammation and are activated by bacterial components (e.g. LPS) to stimulate a pro-inflammatory response, whilst M2 macrophages have anti-inflammatory characteristics, and are typically activated by IL-4, IL-10 and IL-13, and produce IL-10, TGF- and IL-1 receptor antagonist (IL-1Ra) (Gordon, 2003; Stein et al., 1992).

Once activated, M1 macrophages express cytokines (e.g. TNF, IL-12 and IL-6) and chemokines (e.g. CCL2, CXCL10 and CXCL11) that activate and recruit additional immune cells to aid in the clearance of pathogens (Murray and Wynn, 2012). Another major role M1 macrophages has during infection is phagocytosis, to facilitate bacterial clearance (Huynh and Grinstein, 2007). Contrastingly, M2 macrophages are involved in suppressing inflammation, whereby their secretion of IL-10 antagonises M1 macrophages (Murray and Wynn, 2012). In addition, M2 macrophages produce growth factors, including PDGF and TGF-, which increase the expression of tissue inhibitors of metalloproteinases (TIMPs), as well as the activation of extracellular matrix producing myofibroblasts, to promote wound healing (Barron and Wynn, 2011; Murray and Wynn, 2012).

Increased numbers of pro-inflammatory macrophages in gingival tissue from periodontitis patients suggests that macrophages play a role in the pathogenesis of periodontitis (Lappin et al., 2000). An excessive bacterial load (plaque) in periodontitis can cause the chronic recruitment and activation of macrophages. Furthermore, the pro-inflammatory mediators produced by macrophages contribute to the excessive recruitment of other immune cells (e.g. neutrophils and T lymphocytes), and hence promote chronic inflammation. Periodontal infection of mice with P. gingivalis induces macrophage secretion of TNF and IL-1 classical pro-inflammatory cytokines and whose levels correlate with periodontal disease progression (Graves and Cochran, 2003). Together, TNF and IL-1 stimulate a broad range of immunological effects, including orchestrating the migration of leukocytes as part of the physiological response to infection (refer to Section 1.3.3.3 for further details). In periodontitis, a physiological response can transit to disease because the constant recruitment of leukocytes induces chronic inflammation. In addition, the cytokines secreted into the inflammatory milieu by macrophages can contribute to alveolar bone resorption. For example, IL- and TNF can stimulate receptor activator of nuclear factor kappa-B ligand (RANKL) gene expression in osteoblasts, which

12 promotes the development of bone-resorbing osteoclasts (i.e. osteoclastogenesis) (Hofbauer et al., 1999). Importantly, mice in which macrophages had been depleted using clodronate-loaded liposomes were protected from P. gingivalis-induced alveolar bone resorption (Lam et al., 2014). Thus, the excessive production of cytokines arising from the chronic recruitment of macrophages can sustain inflammation that is detrimental to the host.

1.3.1.4 Dendritic Cells Dendritic cells are professional antigen-presenting cells that engage with naïve T lymphocytes to stimulate their activation (Banchereau and Steinman, 1998; Croft et al., 1992). Dendritic cells can also mediate B lymphocyte antibody responses through cytokine production (de Saint-Vis et al., 1998). At steady-state, dendritic cells maintain tissue homeostasis by inducing immune tolerance towards commensal species, whilst still ensuring effective defence against pathogens (Steinman et al., 2003). When activated by microbial components (e.g. LPS and bacterial DNA), dendritic cells release an array of chemokines (e.g. CCL5 and CCL2) to attract additional immune cells (De Smedt et al. 1996; Sparwasser et al. 1998; Sallusto et al. 1999). The chemokine receptor CCR7 is upregulated upon dendritic cell activation and responds to CCL19 and CCL21 produced by stromal cells in secondary lymphoid organs (e.g. spleen and lymph nodes) (Kellermann et al., 1999; Lin et al., 1998). Consequently, activated dendritic cells migrate to lymphoid tissues to activate and communicate antigenic information to lymphocytes. Furthermore, dendritic cells provide immunoregulation to prevent the excessive activation of lymphocytes during infection. Mice depleted of Langerhan cells (using a diphtheria toxin receptor-based system) were used to study the role of dendritic cells in the P. gingivalis-induced mouse model of periodontitis. The mice exhibited enhanced tooth loss, which was associated with increased T lymphocyte-dependent interferon-(IFN-) production and reduced T regulatory cell numbers (Arizon et al., 2012). Therefore, dendritic cells provide host protection by regulating the adaptive immune response to prevent chronic inflammation.

The dysbiotic tooth-accreted biofilm in periodontitis can dysregulate dendritic cell activity. Periodontal pathogens, such as P. gingivalis, can modulate dendritic cell function, and thereby cause a weakened immunostimulatory response (Kanaya et al., 2004, 2009). P. gingivalis does so, at least in part, by stimulating IL-10 secretion from dendritic cells to suppress inflammation (Jotwani et al., 2003; Pulendran et al., 2001). Interestingly, increased IL-10 expression can also cause the chronic recruitment of dendritic cells by enhancing the expression of chemokine receptor, CCR6, by immature dendritic cells, and thus promote their recruitment in a CXCL12-dependent manner into gingival tissues during periodontitis (Dieu-Nosjean et al., 2001). Consequently, this feedback loop may lead to the chronic recruitment of dendritic cells. Dendritic cells also contribute to periodontal tissue destruction by producing increased levels of

MMPs (e.g. MMP-9) in response to P. gingivalis (Jotwani et al., 2010). Consequently, oral pathogens can compromise dendritic cell function and skew dendritic cells towards a host- destructive response.

Adaptive Immune Response in Periodontitis The adaptive immune system generates a highly specialised response tailored to specific pathogens. Cells of the innate immune system (e.g. dendritic cells) engage with naïve T and B lymphocytes to stimulate their proliferation and differentiation into effector and memory cells. Effector T and B lymphocytes recognise target antigens and can induce cell-mediated and humoral immunity, respectively. Both T and B lymphocytes also differentiate into memory cells, which are retained to induce a faster response upon a subsequent encounter with the same antigen. Although the adaptive immune system is important for host protection, the presence of excessive numbers of T and B lymphocytes in periodontal lesions strongly suggest that they also contribute to the pathogenesis of chronic periodontitis (Teng, 2003).

1.3.2.1 T Lymphocytes T lymphocytes recognise processed peptide antigens, presented by cells of the innate immune system (e.g. dendritic cells), via the T cell receptor (TCR). T lymphocytes can differentiate into CD8+ cytotoxic or CD4+ helper T lymphocytes. Cytotoxic T lymphocytes execute immunity by triggering apoptosis of infected cells, and secreting granules containing cytotoxic components (e.g. perforin, granzymes and granulysin) and inflammatory cytokines (e.g. IFN- and TNF) to mediate pathogen clearance (Henkart, 1994; Kägi et al., 1994). In contrast, T helper lymphocytes are involved in coordinating the immune response. Based on their cytokine profiles, T helper lymphocytes are divided into different subsets, including Th1, Th2 and Th17. Th1 cells produce IFN-, IL-2 and TNF to promote cell-mediated immunity and phagocyte-mediated inflammation, whilst Th2 cells produce IL-4, IL-6 and IL-10 to drive an antibody-mediated humoral immune response (Kidd, 2003; Mosmann and Coffman, 1989). Th17 cells produce various cytokines, including IL-17 and IL-22, which primarily act on epithelial cells to induce the expression of antimicrobial proteins (e.g. -defensins and S100 proteins) (Harrington et al., 2005; Liang et al., 2006). Regulatory T lymphocytes are a T helper subset that exert regulatory activity by secreting anti-inflammatory cytokines (e.g. IL-10 and TGF-) to dampen/inhibit immune responses. Together, the proportions of T lymphocyte subsets can dictate the type of adaptive immune response that is mounted, and an imbalance can contribute to inflammatory and autoimmune diseases.

Increased numbers of T lymphocytes in periodontal lesions is correlated with the progression of gingivitis to periodontitis (Liu et al., 2010; Yamazaki et al., 1995). Early periodontal lesions

14 exhibit a typical Th1 cytokine profile characterised by high IFN- levels, which promotes phagocytic activity and cell-mediated immunity (Dutzan et al., 2009). However, the detection of increased IL-4 and IL-6 levels in advanced periodontal lesions suggested that there is a shift from a Th1 to Th2 response in chronic periodontitis (Lappin et al., 2001). Ex vivo analysis of T lymphocytes from patients with chronic periodontitis displayed reduced cell-mediated immune function (Ivanyi and Lehner, 1970). Therefore, it was proposed that Th1 cytokines were important for establishing a stable lesion, whereas the persistence of the tooth-accreted biofilm (plaque) led to a Th2 dominant response. However, the Th1/Th2 paradigm in periodontitis was challenged when several studies demonstrated contradictory levels of Th1 and Th2 cytokines in periodontal lesions (Berglundh, Liljenberg and Lindhe, 2002; Takeichi et al., 2000). The identification of the Th17 lineage led to the re-examination of the Th1/Th2 paradigm. The high levels of the Th17 cytokines IL-17 and IL-22 in inflamed periodontal tissues and gingival crevicular fluid suggested that Th17 cells might mediate host protection by promoting neutrophil chemotaxis and antimicrobial defence (Liang et al., 2006; Vernal et al., 2005; Yu et al., 2007). Indeed, IL-17 receptor-deficient mice showed enhanced alveolar bone loss in a P. gingivalis-induced mouse model of periodontitis (Yu et al., 2007). Although this suggests that Th17 cells have host-protective functions in periodontitis, IL-17-mediated neutrophil infiltration has been implicated in driving alveolar bone resorption in ageing mice (Eskan et al., 2012). Despite emerging studies demonstrating a role for Th17 cells in the pathogenesis of periodontitis, the roles of Th17 cells in host-protection and host-destruction need to be further examined.

1.3.2.2 B Lymphocytes B lymphocytes mediate humoral immunity by secreting antibodies. B lymphocytes are activated upon antigen engagement of the B cell receptor (BCR). In addition, cytokines produced by activated Th2 cells (e.g. IL-4) also promote B lymphocyte activation and differentiation. Once activated, B lymphocytes proliferate and differentiate into plasma cells, which produce antibodies with unique antigen-binding specificities. Antibodies can opsonise bacteria by binding to target antigens present on the surface of bacteria and thereby promote their clearance by phagocytes (e.g. macrophages).

B lymphocytes constitute a major inflammatory infiltrate in established periodontal lesions (Gemmell and Seymour, 1998). The importance of humoral immunity was demonstrated when B lymphocyte-deficient rats were shown to have increased alveolar bone resorption when infected with P. gingivalis (Hou et al., 2000; Klausen et al., 1989). Elevated levels of periodontal bacteria-specific antibodies in individuals affected by gingivitis suggest there is an antibody- driven response to plaque (Ebersole et al., 2001; Lamster et al., 1990). The ability of anti-sera

15 containing high titre, anti-P. gingivalis antibodies from patients with periodontitis to inhibit bone resorption in vitro suggested that the antibodies mediate a protective immune response (Meghji et al., 1993). Interestingly, a separate study found that anti-P. gingivalis antibodies exhibited reduced opsonisation capabilities (Cutler et al., 1991). Clinical studies indicate that high titres of antibodies do not necessarily confer protection, as anti-P. gingivalis antibodies from patient sera can display varying avidities (Lopatin and Blackburn, 1992). The degradation of opsonising antibodies by P. gingivalis proteases (e.g. gingipain proteases) may also influence the immunoreactivity of antibodies (Vincents et al., 2011).

Importantly, the persistence of a tooth-accreted biofilm (plaque) in chronic periodontitis can lead to the generation of auto-reactive antibodies. Increased levels of autoantibodies targeting components of the extracellular matrix (e.g. type I collagen and laminin) have been detected in periodontal lesions, and can contribute to localised tissue destruction in periodontitis (Berglundh et al. 2002; De-Gennaro et al. 2006). Antibodies specific for bacterial antigens produced during periodontitis that can cross-react with self-antigens might result in an autoimmune response. For example, antibodies against P. gingivalis GroEL, a homolog of the human heat shock protein 60 (HSP60), have been shown to cross-react with endothelial HSP60, resulting in endothelial dysfunction in atherosclerosis (Ford et al., 2005; Seymour et al., 2007; Tabeta et al., 2000). As such, humoral immunity can potentially be damaging when it exerts non-specific effects against host antigens.

Molecular Mediators of Host Immunity The host immune response is supported and regulated by an array of bioactive molecules that can activate and amplify host inflammation, promote the activation of immune cells and facilitate the clearance of microorganisms. Importantly, they also regulate the immune response to protect against the development of chronic inflammatory and autoimmune diseases. Therefore, alterations in their levels of expression and/or biological activity can have detrimental effects.

1.3.3.1 Complement System The complement system is comprised of serum proteins, which are activated by proteolytic cleavage to facilitate the clearance of microorganisms. A cascade of enzymatic reactions, involving the successive cleavage of zymogens, mediates the activation and amplification of complement. Complement proteins circulate the body in an inactive form and can be activated through multiple pathways, namely the classical, lectin and alternative pathways, which ultimately converge to generate the same effector molecules (e.g. C3a, C3b and C5a) (Zipfel and Skerka, 2009). When activated, the components of the complement system can exert multiple

16 immunomodulatory activities, including stimulating inflammation (C3a and C5a) and opsonisation of microorganisms (C3b) to promote their uptake by phagocytes (e.g. dendritic cells and macrophages). Other activated complement proteins (C5b, C6, C7, C8 and C9) form a membrane attack complex that can directly perforate the membranes of some microorganisms, causing cell lysis (Zipfel and Skerka, 2009).

Elevated levels of activated complement proteins have been detected in gingival crevicular fluid from patients with periodontitis (Attströum et al., 1975; Patters et al., 1989). Indeed, the continual activation of the complement system by oral pathogens might stimulate chronic inflammation by promoting vasodilation to enhance the flow of inflammatory exudate and chemotaxis of immune cells (Hajishengallis, 2010). Antagonistically, some bacterial proteases (e.g. gingipain proteases and interpain A) can degrade complement factor C3 to inhibit the activation of the complement system (Popadiak et al., 2007; Potempa et al., 2009). P. gingivalis gingipain proteases can also direct signalling crosstalk between complement receptors (e.g. C5aR) and immune receptors to stimulate an inflammatory response, whilst subverting host defence (refer to Section 1.7.1.1 for further details). The administration of a C5aR antagonist in mice was demonstrated to be an effective therapeutic in arresting periodontitis disease progression by reducing levels of pro-inflammatory and bone resorptive cytokines (e.g. TNF and IL-1) (Abe et al., 2012). Therefore, the dysregulation of the complement system can contribute to periodontal inflammation.

1.3.3.2 Antimicrobial Mediators Antimicrobial proteins provide host protection by killing microorganisms directly or indirectly, for example, through limiting nutrient acquisition (Table 1.1). Antimicrobial proteins, including defensins and cathelicidin, are cationic peptides that kill microorganisms directly by binding to negatively charged membrane components (e.g. LPS and lipoteichoic acids), and thus compromise bacterial membrane integrity, resulting in cell lysis (Greer et al., 2013). The human defensins are divided into two subclasses, namely -defensins and -defensins. The -defensins (e.g. human neutrophil peptide (HNP)-1, HNP-2 and HNP-3) are contained in neutrophilic granules, and therefore are largely localised to the junctional epithelium, where there is an influx of neutrophils (Dale et al., 2001). Contrastingly, -defensins (e.g. hBD-1, hBD-2 and hBD- 3) are expressed throughout the stratified oral epithelium to serve as an antimicrobial layer (Dale and Fredericks, 2005). hBD-1 and hBD 2 are expressed constitutively in the spinous, granular and cornified layers in the oral epithelium at steady state, however hBD-2 expression is also upregulated in the presence of oral commensals and pro-inflammatory stimuli (Greer et al., 2013; Krisanaprakornkit et al., 2000; Mathews et al., 1999). hBD-3 is expressed in the basal

17 layers during health and can extend towards the spinous layer in disease (Dale and Fredericks, 2005; Greer et al., 2013).

Human cathelicidin is a cationic, antimicrobial peptide that is expressed by leukocytes (e.g. neutrophils and monocytes) and various epithelia following pro-inflammatory stimulation (Dale et al., 2001). The cathelicidin pro-protein (hCAP18) is processed by host serine proteases (e.g. proteinase 3 and kallikerin) to produce the active cationic peptide, LL-37 (Sørensen et al., 2001; Yamasaki, 2006). Like defensins, LL-37 exhibits a broad spectrum of antimicrobial activity by disrupting microbial membranes. In addition, LL-37 is also chemotactic for monocytes (Yang et al., 2000). Although defensins and LL-37 are effective in killing various oral bacteria (e.g. F. nucleatum and P. intermedia) (Greer et al., 2013), bacterial species in the “red complex” have been found to be less susceptible to the bactericidal activity of -defensins and LL-37 (Bachrach et al., 2008; Joly et al., 2004; Ouhara et al., 2005). The resistance of T. denticola to killing by hBD-2 was found to be attributable to the ability of the bacterium to produce a unique outer membrane lipid, with a lower binding affinity for -defensins (Brissette and Lukehart, 2007; Schultz et al., 1998), whereas P. gingivalis gingipain proteases can degrade and thereby inactivate antimicrobial peptides (Devine et al., 1999; Maisetta et al., 2003; McCrudden et al., 2013). Therefore, the resistance of pathogenic oral microbes to antimicrobial peptides may contribute to microbial dysbiosis.

Antimicrobial proteins can also inhibit microbial growth by preventing the acquisition of essential metal ions by microorganisms. Calprotectin is a heterodimer comprised of S100A8 and S100A9 proteins, and exerts bacteriostatic activity by chelating essential divalent metal ions (e.g. zinc and manganese) (Corbin et al., 2008). During inflammation, calprotectin levels are upregulated in periodontal tissues and gingival crevicular fluid, where it has been shown to facilitate immunity by promoting epithelial barrier function, and thus inhibit P. gingivalis invasion of epithelial cells (Nakamura et al., 2000; Nisapakultorn et al., 2001). Lactoferrin, which is present in gingival crevicular fluid and saliva, inhibits bacterial growth by sequestering iron (Dale and Fredericks, 2005) and exerts growth-inhibitory activity on various oral bacteria (e.g. S. mutans and P. gingivalis) (Aguilera et al., 1998; Arnold et al., 1980). Additionally, lactoferrin was shown to inhibit multispecies biofilm formation in vitro (Arslan et al., 2009; Wakabayashi et al., 2009). Together, antimicrobial mediators prevent the establishment of bacterial infection by exerting bactericidal activity and inhibiting bacterial growth.

Table 1.1 Human antimicrobial mediators of host defence. Antimicrobial Cellular sources Key immune functions

-defensins: Neutrophils • Permeabilise bacterial membrane. HNP-1 Paneth cells • HNP1-3 stimulate migration of HNP-2 monocytes HNP-3 HD5 HD6

-defensins: Epithelial cells • Permeabilise bacterial membrane hBD-1 • hBD-2 stimulates migration of hBD-2 macrophages, neutrophils and mast hBD-3 cells via CCR6

Cathelicidin: Epithelial cells • Permeabilise bacterial membrane LL-37 Monocytes • Stimulate monocyte and neutrophil Neutrophils chemotaxis Calprotectin: Keratinocytes • Sequester divalent cations (e.g. zinc S100A8 Neutrophils and manganese) required for S100A9 Monocytes bacterial metabolism Macrophages • Stimulate neutrophil chemotaxis

Lactoferrin Neutrophils • Sequester iron required for bacterial metabolism Adapted from (Kolls et al. 2008).

1.3.3.3 Cytokines Cytokines are critical immune mediators that coordinate the host inflammatory response through pleiotropic paracrine and autocrine immunomodulatory effects (Table 1.2). Cytokines act through specific receptors, which are cell-surface glycoproteins that function as oligomeric complexes, comprised typically of two to four receptor chains. For instance, the type-1 IL-1 receptor (IL-1R), which recognises IL-1, consists of immunoglobulin-like domains and an intracellular Toll/Interleukin-1 receptor (TIR) domain. Upon IL-1 binding, IL-1R heterodimerises with the IL-1 receptor accessory protein (IL-1RAcP) for signal transduction to occur (Sims and Smith, 2010). Increased levels of TNF and IL-1 are associated with chronic periodontitis (Graves and Cochran, 2003). They are potent pro-inflammatory cytokines, which amplify the inflammatory response by stimulating the upregulation of additional pro- inflammatory genes (Graves and Cochran, 2003). Furthermore, they can enhance the expression of adhesion molecules, including ICAM-1 and VCAM-1, by endothelial cells to promote leukocyte recruitment into affected tissues (Moser et al., 1989; Pober et al., 1986). Pro-inflammatory cytokines (e.g. IL-12 and IL-18) produced by innate immune cells (e.g. macrophages) can also enhance the activation of T lymphocytes to provide specific immunity against infection (Trinchieri, 2003). Other cytokines, including IL-6 and IL-7, can stimulate the proliferation and

19 differentiation of B lymphocytes (Muraguchi et al., 1988; Takatsu, 1997). By contrast, IL-10 is an important anti-inflammatory cytokine, which regulates the immune response by dampening the production of pro-inflammatory molecules (Couper et al., 2008). In collaboration, the combination of cytokines in the inflammatory milieu facilitate and coordinate the host immune response.

Although cytokines are crucial for host defence, they also contribute to inflammatory disease pathology when dysregulated. A dysregulated immune response maintained by excessive levels of pro-inflammatory cytokines is a major contributor to the pathology of chronic periodontitis. Consistently, exogenous administration of TNF in rats was shown to exacerbate inflammation and increase alveolar bone loss (Gaspersic et al., 2003). The IL-1R and TNF receptor (TNFR) signalling in a primate model of experimental periodontitis was shown to significantly reduce alveolar bone loss, which correlated with reductions in the numbers of inflammatory immune cells and bone-resorbing osteoclasts (Assuma et al., 1998). Furthermore, P. gingivalis can destabilise the balance between IL-10 and IL-12 levels to promote inflammation. IL-10-deficient mice were demonstrated to be more susceptible to P. gingivalis-induced alveolar bone loss because of increased IL-12 production, which resulted in enhanced T lymphocyte-mediated activity (Sasaki et al., 2004). Thus, the immune response can be compromised when cytokine signalling networks are dysregulated in chronic periodontitis.

Table 1.2 Roles of major groups of cytokines in host defence.

Cytokine group Examples Key functions

Pro-inflammatory IL-1, IL-6, IL-12, • Promote primary innate immune cytokines TNF response and activation of the inflammatory response • Facilitate the activation of the adaptive immune response Anti-inflammatory IL-10, IL-13, TGF • Downregulate the immune response cytokines and inflammation

Colony-stimulating CSF-1, CSF-2, CSF-3 • Drive myelopoiesis by stimulating factors proliferation and differentiation of monocytes and granulocytes

Th1 regulatory IL-12, IL-18 • Promote Th1 differentiation and cytokines activation

Th2 regulatory IL-4, IL-5, IL-25 • Promote Th2 differentiation and cytokines activation

Th17 regulatory IL-17, IL-22, IL-23 • Promote Th17 differentiation and cytokines activation

B lymphocyte IL-4, IL-5, IL-6, IL-7 • Promote B lymphocyte proliferation regulatory cytokines and activation

Growth factors EGF, TGFPDGF • Regulate tissue repair and fibrosis by promoting proliferation, differentiation and migration of fibroblast, endothelial and epithelial cells. Adapted from (Taylor, 2010).

1.3.3.4 Chemokines Chemokines play an indispensable role in regulating the directed migration of immune cells (Table 1.3). Chemokines are typically characterised by the presence of three to four N-terminal cysteine residues, and further divided into subfamilies based on the position of the first two cysteine residues (Allen et al., 2007). The majority of chemokines belong to the CC and CXC subfamilies. The CXC family of chemokines is further divided based on the presence or absence of a glutamate, leucine and arginine (ELR) motif in the N-terminal region. Chemokines bind to seven transmembrane G-protein coupled receptors, which are named based on the chemokine type they bind (e.g. CC receptor (CCR) and CXC receptor (CXCR)). Most chemokine receptors can

21 be activated by multiple ligands, which enables significant functional redundancy between chemokines (Allen et al., 2007). For instance, CXCR2, which is highly expressed by neutrophils, is activated by multiple chemokines, including CXCL1, CXCL2 and CXCL3.

Chemokines possess an array of immunomodulatory functions. Homeostatic chemokines regulate the migration of immune cells as part of haematopoiesis, which is required for the formation, development and differentiation of leukocytes. For instance, CXCL12 is produced by bone marrow stromal cells and regulates the retention of haematopoietic stem cells in the bone marrow niche (Ara et al., 2003). As their name suggests, inflammatory chemokines are induced in response to inflammatory stimuli, such as microbial products as well as inflammatory cytokines. Classical inflammatory chemokines include IL-8 (CXCL8) and CXCL1, which regulate the recruitment of neutrophils during the early stages of infection. Chemokines also facilitate the adaptive immune response by directing the recruitment of lymphocytes. For example, CXCL9 and CXCL10 activate CXCR3 expressed on the cell-surface of naïve T lymphocytes to promote their migration (Groom and Luster, 2011; Kobayashi, 2006). In addition to regulating immune cell chemotaxis, some chemokines have also been shown to have bactericidal activity. These chemokines, including CCL20 and CCL28, contain positively-charged surface amino acids and appear to act in a similar manner as antimicrobial peptides (e.g. -defensins) to kill bacteria (Hoover et al., 2002; Yang et al., 2003).

By regulating the types of immune cells recruited and activated, chemokines can influence the polarisation of immune responses. The upregulation of CCL2 and its receptor, CCR4, in chronic periodontitis can cause the excessive recruitment of macrophages and exacerbate the inflammatory response (Garlet et al., 2003; Souto et al., 2014). Elevated chemokine expression can also contribute to tissue destruction and alveolar bone resorption. Increased levels of CXCL12 in the gingival crevicular fluid of patients with chronic periodontitis is associated with enhanced osteoclast bone-resorptive activity and MMP-9 production (Grassi et al., 2004; Havens et al., 2008). Moreover, the dysregulation of chemokine responses by bacterial pathogens can lead to a suboptimal immune response. P. gingivalis can inhibit the recruitment of immune cells by blocking nuclear factor-NF-) activation to suppress chemokine expression (e.g. IL-8) (Takeuchi et al., 2013). P. gingivalis gingipain proteases can also degrade chemokines, including IL-8, to inhibit neutrophil chemotaxis (Darveau et al., 1998; Zhang et al. 1999; Stathopoulou et al. 2009). In addition, the CXCL12 receptor, CXCR4, can also be “hijacked” by P. gingivalis to dampen the antimicrobial functions of macrophages (refer to Section 1.7.2 for further details). Consequently, dysregulated chemokine responses arising from microbial dysbiosis can perturb the balance between protective and destructive immunity.

Table 1.3 Roles of chemokines in host defence.

Chemokine Former name Receptor Key immune functions

CXCL1 Gro CXCR2 • Regulate neutrophil migration

CXCL8 IL-8 CXCR1 • Regulate neutrophil migration CXCR2

CXCL10 IP-10 CXCR3 • Regulate T lymphocyte migration. • Promote Th1 adaptive immunity by inducing IFN-expression CXCL12 SDF-1 CXCR4 • Regulate T lymphocyte migration • Maintenance of haematopoietic stem cells

CCL1 I-309 CCR8 • Regulate Th2 lymphocyte migration

CCL2 MCP-1 CCR2 • Regulate monocyte/macrophage migration

CCL5 RANTES CCR1 • Regulate innate immune cells (e.g. natural killer cells) migration • Regulate adaptive immune cells (e.g. T lymphocytes) migration

CCL20 MIP-3 CCR6 • Regulate Th17 lymphocyte migration • Antimicrobial activity

1.4 Pattern Recognition Receptors Pattern recognition receptors (PRRs) are germline-encoded receptors that are critical for the detection of conserved pathogen-associated molecular patterns (PAMPs) by the host. The major families of PRRs include transmembrane Toll-like receptors (TLRs), protease-activated receptors (PARs), cytosolic NOD-like receptors (NLRs), and RIG-I-like receptors (RLRs). PRRs differ in their subcellular localisation and ability to recognise specific PAMPs. PRR activation initiates intracellular signalling cascades, which classically lead to the activation of transcription factors, including NF-B and Interferon Regulatory Factors (e.g. IRF3), which regulate the expression of inflammatory genes (e.g. TNF, IL-6, and IFN-). Although not classical PRRs, protease-activated receptors (PARs) can also detect bacteria by their expression of proteases. Given that bacterial dysbiosis is a critical driver of chronic periodontitis, the following section will focus on receptors that are involved in recognising bacteria.

Toll-like Receptors The TLR family is comprised of ten members and are expressed in varying cell types (Table 1.4) (Takeda et al., 2003). The subcellular localisation of TLRs varies to allow optimal recognition of their cognate ligand. TLRs form homodimers or heterodimers upon activation, and the intracellular TIR domain facilitates protein-protein interactions with adaptor proteins to induce a series of signalling cascades. Ultimately, transcription factors (e.g. NF-B and IRFs) are activated and induce the expression of inflammatory genes to activate and coordinate the immune response. TLRs also mediate the development of adaptive immunity.

Table 1.4 Toll-like receptor subcellular localisation and ligands. Receptor Subcellular Ligand Synthetic agonist Localisation TLR1 Cell surface Triacyl lipopeptides Pam3CSK4

TLR2 Cell surface Lipoproteins Pam3CSK4 Diacyl lipopeptides FSL-1 Triacyl lipopeptides

TLR3 Endosomes Double-stranded RNA Poly I:C

TLR4 Cell surface Lipopolysaccharide TLR5 Cell surface Flagellin TLR6 Cell surface Diacyl lipopeptides TLR7 Endosomes Single-stranded RNA Imiquimod

TLR8 Endosomes Single-stranded RNA Imiquimod

TLR9 Endosomes Unmethylated CpG DNA CpG- oligonucleotides TLR10 Unknown Unknown

1.4.1.1 TLR Distribution and Subcellular Localisation TLRs are widely distributed in various tissues throughout the body. Consistent with their role in immune surveillance, TLRs are highly expressed in tissues that are exposed to the external environment (e.g. oral cavity and skin, and respiratory, gastrointestinal and urogenital tracts), and are therefore potentially more susceptible to infection (Zarember and Godowski, 2002). Epithelial cells at the apical surface of the oral mucosa express lower levels of TLR1 to TLR9 to avoid harmful activation of the immune system by constant exposure to commensal microbes

(Beklen et al., 2008). Peripheral blood leukocytes and spleen cells express the largest repertoires of TLRs (Zarember and Godowski, 2002). The expression of TLRs can vary between different cellular subsets, which can change over the course of leukocyte development. For instance, the expression levels of TLR1, 2, 4, and 5 by immature dendritic cells decrease following maturation (Visintin et al., 2001). Importantly, the selective expression and activation of TLRs on dendritic cells imparts functional specialisation and tolerance in immunity (Dalod et al., 2014).

The TLRs are localised to the cell-surface or intracellular membranes (e.g. endosomes) to facilitate PAMP recognition. TLRs that are localised to the plasma membrane include: TLR2, which can heterodimerise with TLR1 and TLR6 to recognise bacterial triacyl and diacyl lipoproteins, respectively (Takeuchi et al. 2002; Ozinsky et al. 2000); TLR4, which recognises LPS, and TLR5, which recognises flagellin (Hayashi et al., 2001; Hoshino et al., 1999). The other TLRs are localised to intracellular compartments (e.g. endosomal membranes) and include: TLR3, which recognises double-stranded viral RNA (Alexopoulou et al., 2001); TLR7 and TLR8, which recognise single-stranded RNA, and TLR9, which recognises unmethylated CpG- containing DNA (Hemmi et al., 2000). The complex regulation and trafficking of TLRs to the cell surface and endosomal compartments is mediated by specific chaperones (e.g. endoplasmin) (Gay et al., 2014). The subsequent degradation of activated TLRs is essential to regulate the host immune response. Ligand binding by cell-surface TLRs, such as LPS with TLR4, triggers clathrin-mediated endocytosis and subsequent lysosomal degradation (Husebye et al., 2006), whilst endosomal TLRs (e.g. TLR9) are targeted for proteasomal degradation (Chuang and Ulevitch, 2004). As such, the trafficking of TLRs to their correct subcellular localisation allows for optimal ligand recognition as well as the subsequent downregulation of signalling.

1.4.1.2 TLR Structural Organisation Leucine-Rich Repeat Motif TLRs are type I transmembrane receptors, and consist of an N-terminal leucine-rich repeat (LRR) domain, a transmembrane region, and a C-terminal TIR domain (Fig. 1.2). The LRR domain is comprised of multiple repeats of a conserved “LxxLxLxxN” motif, where “x” denotes a hydrophobic residue. The conserved leucine residues confer the domain with a horseshoe-like structure, as the side-chains point inward to form a hydrophobic core (Fig. 1.3). Each TLR has a distinct LRR domain, which is important for PAMP-binding specificity (Bella et al., 2008).

LRR motif

Transmembrane domain

Phospholipid bilayer

TIR domain

Figure 1.2 Structural arrangement of Toll-like receptors. TLRs are comprised of an extracellular LRR domain that forms a horseshoe-like structure, which binds a cognate ligand. The intracellular Toll-Interleukin-1 (TIR) domain is crucial for facilitating adaptor protein recruitment, through homotypic interactions, for subsequent intracellular signalling.

Figure 1.3 Crystal structure of the TLR3 LRR motif. The TLR3 ectodomain (PDB: 3CIG) forms a solenoid structure, comprised of twenty-three LRR repeats. The conserved hydrophobic residues of the LRRs form a tightly packed hydrophobic core, which provides lateral stability.

Toll/Interleukin-1 Receptor (TIR) Domain The intracellular TIR domain facilitates the binding of specific adaptor proteins (e.g. myeloid differentiation primary response 88 (MYD88) and MYD88 adaptor like (MAL)), which is required for subsequent downstream signalling. As its name suggests, the TIR domain shares a high degree of homology with the cytoplasmic domain of the type-1 IL-1 receptor. The TIR domain is composed of around 200 amino acid, and typically adopts a flavodoxin-like fold comprised of five central -sheets (A - E) surrounded by five -helices (A - E) (Fig. 1.4). The loops adjoining -sheets and -helices are named according to the secondary elements to which they are connected; for instance, the BB loop connects the B strand and B helix.

Mammalian TIR domains share 20-30% sequence conservation, which provides sufficient structural diversity to confer signalling specificity (Xu et al., 2000). The TIR domain of TLRs is conserved in three regions, denoted as box 1, box 2 and box 3 (Watters et al., 2007). Box 1 signifies the start of the TIR domain and is characterised by a conserved (F/Y)DAFISY motif. The box 2 motif in the BB loop, which has been studied most extensively, is an important structural determinant for TIR-TIR interactions (Xu et al. 2000). The BB loop is characterised by an

RDxɸ1ɸ2G motif, where “x” denotes any residue, and “ɸ” denotes a hydrophobic residue. The proline residue at the ɸ2 position has also been shown to be required for TLR signalling

(Poltorak et al., 1998; Tao et al., 2002). For example, the natural mutation of Pro712 to histidine in TLR4 renders CH3/HeJ mice unresponsive to LPS (Poltorak et al., 1998). Similarly, a Pro681 to histidine mutation in the BB loop of TLR2 impaired signalling in response to Mycobacterium tuberculosis (Underhill et al., 1999). Further studies revealed that the mutation disrupted the interaction between TLR2 and MYD88 (Xu et al., 2000). Box 3 is defined by a FW motif, which is surrounded by basic amino acids. The functional role of box 3 has yet to be fully elucidated; however, mutagenesis studies indicate that it is required for signalling, and, in the case of IL-1R, for correct receptor subcellular localisation (Slack et al., 2000). In summary, TIR domains share conserved features to enable specific TIR-TIR interactions and downstream signalling.

A B

Figure 1.4 Crystal structure of the TLR1 and TLR2 TIR domains. The TIR domain of (A) TLR1 and (B) TLR2 contain a characteristic flavodoxin fold comprised of -sheets surrounded by helices. The box 1, box 2 and box 3 are as indicated by the yellow, green and pink colouring, respectively. 1.4.1.3 Toll-like Receptor Signal Transduction When activated, TLRs can either homo or heterodimerise through TIR-TIR domain interactions. Furthermore, the interacting TIR domains provide a platform for subsequent binding by TIR domain-containing adaptor proteins to initiate downstream signalling. Depending on the adaptor protein(s) that is recruited, TLRs can signal either through the MYD88-dependent pathway or TRIF-dependent pathway (Fig. 1.5). The MYD88-dependent pathway is utilised by all TLRs, with the exception of TLR3, to stimulate downstream signalling. In the case of TLR2 and TLR4, MAL acts as a bridging adaptor to facilitate the recruitment of MYD88 (Fitzgerald et al., 2001). The protein kinases Interleukin-1 receptor-associated kinase 4 (IRAK-4) and IRAK-1 subsequently interact with MYD88 through their N-terminal death domain (Wesche et al., 1997). Activated IRAK-1 phosphorylates and thereby activates the E3 ubiquitin ligase, Tumour

28 necrosis factor receptor-associated factor-6 (TRAF6). TRAF6, in conjunction with the ubiquitin conjugating enzymes, UEV1A and UBC13, catalyses the Lys-63 polyubiquitination of TAB2 and TAB3, which are regulatory components of the protein kinase, TGF--activated kinase (TAK1). This causes the activation of TAK1, and brings it into close proximity with the inhibitor nuclear factor-B (IB) kinase (IKK) complex. The IKK complex regulates the activity of IB, which sequesters NF-B in the cytoplasm to prevent its activation and translocation into the nucleus. TAK1-mediated phosphorylation of the IKK complex leads to NF-B activation via the phosphorylation and subsequent proteasomal degradation of IB. This releases NF-B, allowing it to translocate into the nucleus to activate inflammatory gene expression (e.g. TNF and IL-6).

TLR3, as well as TLR4, signals through the TIR domain-containing adaptor-inducing Interferon- (TRIF)-dependent pathway (Yamamoto et al., 2003a); TLR4 specifically requires the protein adaptor TRIF-related adaptor molecule (TRAM) for the recruitment of TRIF (Jenkins et al., 2010). TRIF signalling leads to the activation of NF-B and IRF3. TRIF associates with TRAF6 and induces NF-B activation in a similar manner to the MYD88-dependent pathway. In addition, TRIF can also stimulate IRF3 activation by forming a complex with TANK-binding protein kinase-1 (TBK1) and inducible B kinase IKKi, which phosphorylate and thereby activates IRF3 (Sato et al., 2003). Once activated, IRF3 can then translocate into the nucleus to transcribe type I interferon and interferon-stimulated genes (e.g. 2'-5'-oligoadenylate synthetases).

Figure 1.5 TLR signalling pathways. All TLRs, except for TLR3, signal via the MYD88- dependent pathway. TLR4 utilises both the MYD88 and TRIF pathways to activate the inflammatory response.

1.4.1.4 TIR domain-containing Adaptor Proteins As indicated above, TIR domain-containing adaptor proteins play a central and critical role in TLR signalling. The subcellular localisation of TIR domain-containing adaptor proteins is essential in facilitating and ensuring efficient signal transduction. The initial binding of TIR domain-containing adaptor proteins (e.g. MYD88 and TRIF) upon TLR activation promotes the assembly of intracellular signalling platforms for the subsequent recruitment of downstream adaptor proteins to confer TLR signalling specificity. Post-translational modifications to adaptor proteins also serve as an additional level of regulation for TLR signalling.

Myeloid Differentiation Primary Response 88 (MYD88) The crucial role of MYD88 in TLR signalling was demonstrated when MYD88-deficient mice were shown to be unresponsive to LPS (Kawai et al., 1999). The MYD88 protein has an N-terminal death domain and a C-terminal TIR domain, which are separated by an intermediate domain (Fig. 1.6). At steady state, MYD88 is localised throughout the cytosol; however, MYD88 has also been shown to form punctate inclusions when ectopically overexpressed in cells (e.g. RAW 264.7 mouse macrophages) (Into et al., 2010; Nishiya et al., 2007). When activated, the N-terminal death domain of MYD88 is essential for its correct subcellular localisation to facilitate TIR domain interaction with TLRs, and to recruit IRAK family members (e.g. IRAK-4 and IRAK-1) for downstream signalling (Dunne et al., 2003; Nishiya et al., 2007).

MYD88 activity can be modulated to regulate the inflammatory response. An alternative splice variant of MYD88, lacking the intermediate domain, is expressed in monocytes upon LPS stimulation (Janssens et al., 2002). This splice variant of MYD88 inhibits LPS-induced NF-B signalling because it cannot recruit IRAK-4, which is necessary for IRAK-1 phosphorylation and activation (Burns et al., 2003). MYD88-mediated signalling is also regulated by TGF-, whereby MYD88 is ubiquitinated and targeted for proteasomal degradation (Lee et al., 2011).

MYD88 Adaptor-Like (MAL) The importance of MAL for TLR2 and TLR4 signalling was established with MAL-deficient mice (Horng et al., 2002; Yamamoto et al., 2002). MAL is comprised of an N-terminal phosphoinositide-binding domain, which can bind phosphatidylinositol-4, 5-biphosphate

(PtdIns(4,5)P2) and thereby localise MAL to the plasma membrane (Kagan et al., 2006), and a C-terminal TIR domain (Fig. 1.6). The physical association of MAL with the plasma membrane facilitates its role as a bridging adaptor for MYD88 with TLR2 and TLR4. The electro-negative charge on the surface of the TIR domain of MAL allows it to interact with the electro-positive surfaces of the corresponding TIR domains of TLR4 and MYD88 (Dunne et al., 2003). In addition

31 to acting as a bridging adaptor, MAL can direct TLR signalling in a MYD88-independent manner by recruiting TRAF6 into a signalling complex through its TRAF6-binding domain (Mansell et al., 2004). MAL is also involved in the negative regulation of TLR2 and TLR4 signalling. The phosphorylation of MAL by Bruton’s tyrosine kinase in response to TLR2 and TLR4 activation triggers its interaction with the suppressor of cytokine signalling 1 (SOCS-1) to promote the subsequent degradation of MAL (Mansell et al., 2006). Thus, not only is MAL involved in signal transduction required to initiate inflammation, its degradation also forms part of the negative regulation of TLR responses to limit inflammation.

TIR-domain-containing Adaptor-inducing Interferon- (TRIF) TRIF is essential for mediating TLR3 and TLR4-activated type I interferon induction (Yamamoto et al. 2003b). TRIF is directly recruited to TLR3, whilst TRAM is required to facilitate TRIF recruitment to TLR4 (Yamamoto et al. 2003c). TRIF is characterised by an N-terminal TRAF6-binding domain, and a C-terminal TIR domain (Fig. 1.6). Mutations in the TRAF6-binding domain compromises NF-B but not IRF3 activation (Jiang et al., 2004; Sato et al., 2003). TRIF forms a complex with TBK1 through its N-terminus to mediate IRF3 activation (Sato et al., 2003). Moreover, studies indicate that NF-B-activating kinase (NAK)-associated protein (NAP1) is required for TRIF to form a signalling complex with TBK1 (Sasai et al. 2005). TRIF is also subject to negative regulation, whereby the C-terminal domain of TRAF1 can interact with the TIR domain of TRIF to inhibit signalling (Su et al., 2006).

TRIF-related Adaptor Molecule (TRAM) TRAM is similar to MAL, in that it acts as a bridging adaptor for TRIF to interact with TLR4 (Yamamoto et al. 2003c). The TIR domain is localised at the C-terminus of TRAM (Fig. 1.6). Myristoylation of the second glycine residue of TRAM allows its hydrophobic interaction with the lipid bilayer to stabilise its association with the plasma membrane (Rowe et al., 2006). TLR4-mediated TRAM-TRIF signalling occurs following endocytosis of the TLR4 complex (Kagan et al., 2008). It was proposed that the rearrangement of the TLR4 TIR domain in the acidic environment of endosomes enables optimal interaction with TRAM (Gangloff, 2012). The protein kinase C (PKC)-mediated phosphorylation of TRAM at serine-16 is essential for TRAM signalling (McGettrick et al., 2006). However, the mechanistic involvement of PKC-mediated phosphorylation of TRAM has yet to be elucidated.

Sterile Alpha and Armadillo-motif-containing (SARM) SARM was the last TIR domain adaptor protein to be identified, and acts as a negative regulator of TRIF signalling (Carty et al., 2006). It is comprised of two consecutive sterile alpha motif

(SAM) domains, and a TIR domain at the C-terminus (Fig. 1.6). The expression of SARM is upregulated in response to TLR4 activation and SARM acts as an inhibitor of NF-B and IRF3 activation (Carty et al. 2006). It was proposed that SARM associates with TRIF transiently in the resting state, and the activation of TLR3 or TLR4 stabilises this interaction to block the recruitment of TIR domain adaptor proteins to inhibit downstream signalling (O’Neill, 2006). Although SARM does not affect MYD88-dependent NF-B signalling, it was recently found to inhibit the MYD88-dependent activation of the transcription factor activator-protein-1 (AP-1) (Peng et al., 2010). Further studies will be required to clarify the mechanism of SARM inhibition of TRIF, and to elucidate whether SARM has additional functions in TLR signalling.

MYD88 Key

Death domain MAL Intermediate domain

TRIF TIR domain Phosphoinositide- TRAM binding motif

TRAF6-binding domain SARM SAM motif

Figure 1.6 Structural arrangement of TIR domain-containing adaptor proteins. NOD-like Receptors NOD-like receptors (NLRs) are a family cytosolic PRRs that can detect invading pathogens and their products. The NLR family consists of 23 members, which are expressed in various cell types, including epithelial cells and immune cells (Franchi et al., 2009). Structurally, NLRs are characterised by an N-terminal effector domain (e.g. BIR, CARD or Pyrin domain), which mediates its interaction with other proteins, a central NACHT domain, which mediates homo-oligomerisation, and a C-terminal LRR domain for bacterial-sensing. Based on their N-terminal domain, NLRs can be classified into four subfamilies: (i) NLRA, (ii) NLRB, (iii) NLRC and (iv) NALP (Fig. 1.7). The activation of NLRs is triggered by the LRR binding a PAMP, which initiates the oligomerisation of the receptor through the NACHT domain for subsequent recruitment of downstream signalling proteins.

NOD1 and NOD2 are two of the most widely studied members of the NLR family and recognise bacterial cell wall peptidoglycan fragments, namely, muropeptide (iE-DAP) and muramyl dipeptide (MDP), respectively (Chamaillard et al., 2003; Inohara et al., 2003). Upon ligand binding, NOD1 and NOD2 undergo self-oligomerisation and recruit receptor-interacting serine-

33 threonine kinase 2 (RIP2) via a CARD-CARD interaction (Abbott et al., 2004). Activated RIP2 promotes TRAF-mediated (TRAF2 and/or TRAF5 in the case of NOD1 and TRAF6 in the case of NOD2) K63-linked ubiquitination TAK1 and the IKK complex. The ensuing co-localisation of TAK1 and IKK complex leads to the phosphorylation and subsequent activation of the IKK complex for the degradation of IB, resulting in NF-B activation and nuclear translocation (Fig. 1.8). RIP2 can also signal with inhibitor of apoptosis (IAPs) to induce NOD-mediated NF-B activation (Krieg et al., 2009). Notably, NOD2 has also been found to be involved in the negative regulation of TLRs. For example, co-stimulation of TLR2 and NOD2 with peptidoglycan and MDP, respectively, inhibited TLR2-induced IL-12 production in splenocytes (Watanabe et al., 2004).

NLRs also regulate inflammation by forming inflammasomes, multimeric complexes that regulate the activation of inflammatory caspases, which mediate IL-1 maturation. Pro-IL-1 expression is typically maintained at the transcriptional level by NF-B, whilst the processing and secretion of mature IL-1is regulated by inflammasome-mediated caspase-1 activation (Fitzgerald, 2010). NLRP1, NLRP3 and NLRC4 act as cellular sensors for the assembly of canonical inflammasomes. When activated, NLRP1 and NLRP3 interact with the adaptor protein apoptosis-associated speck-like protein containing a CARD domain (ASC), which in turn binds the CARD domain of caspase-1 to facilitate inflammasome assembly (Schroder and Tschopp, 2010). Contrastingly, homo-oligomers of NLRC4 can interact directly with caspase-1 independently of ASC (Schroder and Tschopp, 2010). Consequently, a regulated and controlled inflammatory response is achieved through both the transcriptional control and post- translational processing of IL-1.

Key NLRA CIITA CARD NLRB NAIP NACHT

LRR NOD1 and BIR NLRC4 Pyrin NOD2 NLRC Undefined domain

NLRC3 and NLRC5

NLRP1 NLRP2 NLRP and NLRP9

NLRP10

Figure 1.7 Structural arrangement of NOD-like receptors.

Figure 1.8 NOD-like receptor signalling pathways. NOD2 homo-oligomerises upon binding of muramyl dipeptide (MDP) fragments derived from microbes, and recruit downstream signalling adaptor proteins to stimulate a pro-inflammatory response.

Protease-activated Receptors The innate immune system can also sense proteolytic enzymes during infection through protease-activated receptors (PARs). The family of four PARs are transmembrane G-protein coupled receptors that regulate an array of responses (Table 1.5). Host- and bacterial-derived serine proteases cleave PARs to expose an N-terminal sequence, which acts as a tethered ligand and interacts with the second extracellular loop of the receptor to initiate intracellular signal transduction (Soh et al., 2010). Synthetic, activating peptides have been designed to mimic the tethered ligand and stimulate receptor activation. Ligand activation of PARs induces a conformational change to stabilise the binding of heterotrimeric G proteins, comprised of ,  and  subunits. Once activated, the G proteins act as guanine nucleotide exchange factors, whereby GDP bound by the  subunit is converted to GTP. This also results in the dissociation of the  and  subunits to induce diverse cellular responses, including inflammation, haemostasis and thrombosis (Soh et al., 2010). Although PARs are activated by similar mechanisms, they can regulate different biological outcomes depending on their tissue distribution. Furthermore, the association of PARs with different G subunits also confer signalling specificity. For instance,

PAR-1 can interact with Gq11, G12/13 and Gi, whilst PAR-2 is coupled to Gq. Following activation, PARs (with the exception of PAR-1) are subsequently phosphorylated and recognised by -arrestins to facilitate their internalisation via clathrin-coated pits to dampen signalling (DeFea et al., 2000; Paing et al., 2002).

PARs have been extensively studied for their role in the coagulation cascade; however, they have also been shown to be activated by bacterial proteases and activate host inflammatory responses. The gingipain proteases expressed by P. gingivalis were shown to activate PAR-2 in gingival epithelial cells and enhance the production of IL-6 (Lourbakos et al., 2001). Moreover, PARs have also been shown to contribute to the inflammatory response by promoting immune cell recruitment by stimulating the upregulation of endothelial VCAM-1 and ICAM-1 expression (Coughlin and Camerer, 2003; Rahman et al., 2002). Consistently, alveolar bone resorption in a P. gingivalis-induced model of periodontitis was reduced in PAR-2-deficient mice (Holzhausen et al., 2006). Thus, PARs can serve as a defence mechanism to trigger host inflammation by sensing bacterial proteases.

Table 1.5 PARs activating proteases, cellular expression and key functions.

Activating Activating Receptor Expression Key function(s) Protease peptides

PAR-1 Platelets Thrombin SFLLR-NH2 • Platelet aggregation Epithelia • Vascular smooth muscle Endothelia cell proliferation Fibroblasts • Cytokine secretion Neurons

PAR-2 Epithelium Trypsin SLIGKV-NH2 • Cell migration Endothelium Tryptase • Cell proliferation Fibroblasts Factor VIIa Neurons Factor Xa Elastase

PAR-3 Endothelium Thrombin • Cytokine secretion

PAR-4 Platelets Thrombin GYPGQV-NH2 • Platelet activation Endothelium Trypsin AYPGKF-NH • Neutrophil recruitment Plasmin 2

Adapted from (Ossovskaya and Bunnett 2004). 1.5 Microbial Subversion of TLR Signalling Many bacterial and viral pathogens have evolved mechanisms to subvert and thereby modulate the host immune response to their benefit. The innate immune system is particularly attractive as a target for subversion, as suppressing innate immunity may also compromise adaptive immunity. Pathogens can subvert the host immune responses through multiple mechanisms, including PAMP modification and targeting proteins critical for PRR signalling.

PAMP Modification Structural modifications of membrane-surface PAMPs can mask PRR recognition sites to avoid immune detection. LPS is a major component of the outer membrane of Gram-negative bacteria. It is an amphipathic molecule comprised of a hydrophilic polysaccharide chain, denoted as the O-antigen, a core region, and a hydrophobic lipid A moiety. TLR4 recognises the lipid A component of LPS, which is conserved in Gram-negative bacteria (Poltorak et al., 1998; Shimazu et al., 1999). Interestingly, the microenvironment to which Gram-negative bacteria are exposed may generate LPS species with different inflammatory capacities due to differential modification of lipid A (e.g. acylation) (Dixon and Darveau, 2005). Highly-acylated lipid A (e.g. hexa-acylated lipid A of Escherichia coli) is a potent inflammatory stimulus, whereas less- acylated lipid A (e.g. tetra-acylated lipid A of P. gingivalis) is a weaker inflammatory stimulus (Barksby et al., 2009). Salmonella typhimurium LPS is subjected to 3-O-deacylation and

38 palmitoylation by the lipid modifying enzymes 3-O-deacylase (PagL) and lipid A palmitoyltransferase (PagP), respectively (Bishop et al., 2000; Trent et al., 2001). These modifications were demonstrated to prevent TLR4, and hence NF-B, activation (Kawasaki et al., 2004). Similarly, Pseudomonas aeruginosa lipid A containing a 3-O-deacylation modification was also shown to stimulate weaker TLR4-mediated signalling (Stöver et al., 2004).

TLR5 recognises the conserved N-terminal region of flagellin, the main structural component of flagella. Polymorphisms in the N-terminal region of flagellin can induce weaker TLR5 activation. For example, the absence of the conserved 89-96 amino acid sequence towards the N-terminus of flagellin from alpha proteobacteria and epsilon proteobacteria (e.g. Helicobacter pylori and Campylobacter jejuni, respectively) results in it no longer being detected by TLR5 (Andersen- Nissen et al., 2005). Bacteria can also avoid TLR5 recognition by modulating flagellin expression; Listeria monocytogenes downregulates flagellin at physiological temperature (37 °C) (Kathariou et al., 1995). Thus, modifications to bacterial cell-surface components can significantly reduce the magnitude of the host immune response elicited.

Targeting TLR Signalling Proteins As discussed above, TLR adapter proteins are essential for signal transduction by TLRs. Therefore, pathogens have evolved mechanisms to target TLR adaptor proteins as a strategy to suppress host immunity. Bacterial pathogens can modulate MAPK signalling by expressing bacterial phosphatases. Shigella flexneri expresses a phosphatase, OspF, which is delivered into the host cell cytoplasm via a type III secretion system, where it translocates into the nucleus and dephosphorylates and inactivates ERK1/2 and p38 MAPK (Arbibe et al., 2007; Zurawski et al., 2006). Transcriptional analyses indicate that OspF suppresses the expression of early inflammatory genes, including IL-8 and CCL20 (Arbibe et al., 2007). Consequently, mice infected with an OspF-deficient S. flexneri mutant developed more severe mucosal lesions associated with a greater neutrophil influx (Arbibe et al., 2007). Viruses also target components downstream of TIR-domain-containing adaptors. The vaccinia viral protein, A52R, was demonstrated to inhibit NF-B activation by interacting with TRAF6, and thereby suppress TLR3-mediated IL-8 and CCL5 expression (Bowie et al., 2000; Maloney et al., 2005). In addition, A52R can also stimulate the activation of p38 mitogen-associated protein kinase (MAPK) to induce the expression of the anti-inflammatory cytokine, IL-10 (Maloney et al., 2005). The upregulation of IL-10 expression dampens inflammation and cell-mediated immune response, thus reducing virus elimination by the host. Taken together, the inhibition of TLR signalling is a key strategy used by pathogens to suppress host defence.

1.6 Bacterial TIR Domain-containing Proteins Some pathogens have evolved TIR domain-containing proteins that can disrupt host TIR-TIR interactions and thus interfere with TLR signalling (Table 1.6). Bacteria have been found to express TIR domain-containing proteins (Tcps) that inhibit TLR-induced responses. Using bioinformatics approaches, Newman et al. identified over 200 bacterial proteins containing a putative TIR domain (Newman et al., 2006). These bacterial Tcps are found in various bacterial phyla, including Chlorobi, Proteobacteria and Bacteroidetes (Spear et al., 2009). Phylogenetic analysis suggests that bacterial Tcps may have evolved through multiple horizontal gene transfer events (Zhang et al., 2011). Several bacterial Tcps have been characterised and studied for their ability to inhibit TLR signalling. This section will highlight the involvement of functionally characterised bacterial Tcps in suppressing TLR signalling.

Table 1.6 Bacterial Tcps and immune subversion. Protein Bacterium Mode of antagonisation Functional in vivo studies TcpB Brucella • Inhibit TLR4-mediated • Facilitate systemic spread melitensis signalling of bacteria • Interact with MAL, MD88 and TLR4 • Downregulate MAL expression

TcpC Escherichia • Inhibit TLR2 and • Required for virulence in coli TLR4-mediated signalling mouse model of urinary • Interact with MYD88 tract infection • Facilitate bacterial colonisation PdTIR Paracoccus • Interact with TLR4 and denitrifcans MYD88

YpTdp Yersinia pestis • Interact with MYD88 • Not required for bacterial • Interfere with colonisation TLR2-dependent signalling

TIRs Staphylococcus • Inhibit MAL and MYD88- aureus dependent immune response

TlpA Salmonella • Inhibit TLR signalling • Mice infected with enterica • Induce caspase-1- TlpA-deficient mutant dependent IL-1 secretion exhibited reduced lethality

Structural Properties of Bacterial Tcps Bacterial Tcps are typically 200-300 amino acids in length, and have a C-terminal TIR domain that is comprised of approximately 150-200 amino acids. Initial structural homology modelling of TcpC of E. coli CFT073, a uropathogenic strain of E. coli and TcpB of Brucella melitensis suggested that their TIR domains may adopt a fold similar to mammalian TIR proteins (Cirl et al., 2008). Indeed, the crystal structure of TcpB from B. melitensis and PdTIR from Paracoccus dentrificans revealed that their TIR domains form a flavodoxin-like fold, characteristic of human TIR domains (Chan et al., 2009; Snyder et al., 2014). The TIR domains of TcpB and PdTIR were isolated as monomers; however, their crystal lattice structures revealed potential dimer interfaces (Chan et al., 2009; Snyder et al., 2014). Contrastingly, the TIR domain of YpTdp from Yersinia pestis formed dimers in solution, which was mediated by the formation of two disulphide bonds (Rana et al., 2011). Additionally, the N-terminus of TcpB and PdTIR, and TlpA from Salmonella enterica, contain an -helical coil-coiled domain, which is proposed to facilitate protein dimerisation and stabilise the TIR-TIR interaction interface, and may thereby enhance the inhibition of TLR signalling (Fekonja et al., 2012).

Interference with TLR signalling by Bacterial Tcps Given the structural similarities between bacterial Tcps and human TIR domains, functional studies have largely focused on their role as potential virulence factors in suppressing TLR responses (Table 1.6). Newman et al. provided evidence for the ability of TlpA from S. enterica serovar Enteritidis to inhibit TLR and IL-1R-mediated NF-B activation (Newman et al., 2006). Further studies revealed that TcpC and TcpB were also capable of blocking TLR2- and TLR4-induced NF-B activation (Cirl et al., 2008). The molecular mechanism of TcpB-mediated inhibition of TLR2 and TLR4 is unclear because different studies identified different binding partners for TcpB. It was initially proposed that TcpB mimicked MAL function, by co-localising to the plasma membrane through its phosphoinositide-binding motif, and thereby blocked the formation of the signalling platform required for MYD88 recruitment (Radhakrishnan et al., 2009). TcpB has also been shown to interact with MAL, but not MYD88, to promote the ubiquitination and subsequent degradation of MAL (Sengupta et al., 2010). However, a more recent study suggested that TcpB can bind to MAL, MYD88 and TLR4, and thus may interfere with the activation of NF-B (Alaidarous et al., 2014). Like other bacterial Tcps, YpTdp can inhibit NF-B activation by TLR4 and IL-1R signalling (Spear et al., 2012). The interaction of YpTdp with MYD88 was dependent on the conserved proline residue (Pro173) in the BB loop of YpTdp (Spear et al., 2012). It is clear from these studies that bacterial Tcps can interact differently with mammalian TIR domain proteins. Thus, further studies are required to define

41 the precise mechanism(s) of interactions between bacterial Tcps and mammalian TIR domain proteins.

Functional Consequences of Bacterial Tcps Intuitively, the inhibition of TLR signalling by bacterial Tcps would dampen the host inflammatory response and thus might be advantageous to microbial growth. Indeed, in vitro studies revealed that TlpA-deficient S. entrica was no longer able to survive and/or replicate in human THP-1 monocytes. Moreover, mice inoculated with the TlpA-deficient S. entrica mutant had prolonged survival and reduced bacterial burden, compared to mice infected with the wildtype strain (Newman et al., 2006). Similarly, in vitro studies also demonstrated the importance of TcpC for the intracellular survival of E. coli CFT073 in RAW264.7 macrophage- like cells. Consistently, TcpC-deficient E. coli CFT073 mutant displayed reduced bacterial colonisation and tissue damage in a mouse model of urinary tract infection (Cirl et al., 2008). A B. melitensis TcpB-deficient mutant also displayed a reduction in dissemination during the early stages of infection in mice (Radhakrishnan et al., 2009). In contrast to these observations, host cytokine responses and bacterial colonisation were comparable for wildtype Y. pestis and a YpTdp-deficient Y. pestis mutant (Spear et al., 2012). However, Y. pestis growth characteristics in vitro were affected by YpTdp deficiency, as the YpTdp-deficient mutant exhibited spontaneous aggregation in broth culture, and was also intolerant of high salinity (Spear et al., 2012). In summary, bacterial Tcps have been shown to be involved in subverting immune detection and inhibiting the host inflammatory response to enhance survival and/or replication.

1.7 Immune Subversion by Porphyromonas gingivalis P. gingivalis is one of the core species of the “red complex” that is closely associated with driving the development of chronic periodontitis (Hajishengallis et al., 2011, 2012; Socransky et al., 1998). Subgingival levels of P. gingivalis have been shown to correlate with the clinical symptoms of chronic periodontitis, including bleeding on probing, periodontal pocket depth, attachment loss, and alveolar bone resorption (Byrne et al., 2009; Griffen et al., 1998; Socransky et al., 1998). Effective periodontal treatment is associated with reduced P. gingivalis levels in subgingival plaque (Haffajee et al., 1997; Kawada et al., 2004). Despite being a low abundance species, P. gingivalis is capable of dysregulating the host immune response to promote biofilm dysbiosis and chronic inflammation, which is achieved by the pathogen expressing various virulence factors (Hajishengallis et al., 2011). Thus, P. gingivalis has been proposed to be a “keystone pathogen” in chronic periodontitis (Hajishengallis et al., 2012). The following Section will discuss P. gingivalis virulence factors, and how they impact the host immune response and thereby promote disease development.

P. gingivalis Gingipain Proteases The extracellular gingipain proteases expressed by P. gingivalis are critical virulence factors. They are cysteine proteases, and encoded by distinct but related genes to produce the lysine- specific gingipain, Kgp, and the arginine-specific gingipains, RgpA and RgpB (Potempa et al., 1995). Kgp and RgpA contain a C-terminal haemagglutinin domain, followed by a catalytic protease domain. While its protease domain is identical to RgpA, RgpB lacks a haemagglutinin domain. The gingipain proteases are primarily attached to the outer membrane of P. gingivalis through LPS-associated modifications; however, they can also be found on outer membrane vesicles (OMVs) or secreted in a soluble form (Potempa et al., 2003; Veith et al., 2014). Consequently, the gingipain proteases can exert stimulatory activity on host tissues locally, as well as at distant sites.

1.7.1.1 Gingipain Proteases and Immune Subversion The gingipain protease-mediated dysregulation of immune signalling is important for the ability of P. gingivalis to counter many host defence mechanisms. For example, the gingipain proteases can proteolytically degrade the TLR4 co-receptor CD14 on monocytes and fibroblasts, thus making the cells hyporesponsive to LPS (Sugawara et al., 2000; Tada et al., 2002). In addition, the degradation of RIP2 by the gingipain proteases has also been shown to inhibit NOD1 and NOD2 signalling (Madrigal et al., 2012). The proteolytic degradation of inflammatory cytokines, such as TNF and IL-6, by the gingipain proteases may also dampen the host inflammatory response (Calkins et al., 1998; Banbula et al., 1999). Notably, the degradation of IL-8 by the gingipain proteases is proposed to cause “local chemokine paralysis” by impairing neutrophil recruitment (Darveau et al. 1998; Zhang et al. 1999; Stathopoulou et al. 2009). Interestingly, complement activity may be modulated based on the levels of the gingipain proteases produced during different stages of P. gingivalis infection. It has been proposed that during the early stages of infection, P. gingivalis is present in lower numbers and thus the levels of gingipain proteases are low. This may result in the proteolytic processing of complement C3, C4 and C5 into active fragments that stimulate inflammation, and hence increase nutrient availability to support P. gingivalis growth (Popadiak et al., 2007). Once P. gingivalis has established an ecological niche, the increased numbers of P. gingivalis present correlated with the higher levels of gingipain proteases produced, which may degrade and inactivate complement activity, and thus help to suppress the immune response (Popadiak et al., 2007). Therefore, the proteolytic activity of the gingipain proteases enables P. gingivalis to subvert the host immune response by degrading an array of host immunomodulatory factors.

The gingipain proteases can also manipulate the host immune response by directing immune signalling crosstalk. For instance, the gingipain proteases can proteolytically process C5 to

43 generate C5a, and thereby activate C5aR on neutrophils and macrophages (Maekawa et al., 2014; Popadiak et al., 2007; Wang et al., 2010). In neutrophils, signalling crosstalk between C5aR and TLR2 results in the production of TGF-, which triggers the activation of the E3 ubiquitin ligase SMURF1, and promotes proteasome-mediated degradation of MYD88 (Maekawa et al., 2014). Loss of MYD88 signalling thus suppresses the production of nitric oxide, which is important for neutrophil-mediated bacterial killing. Concomitantly, the synergistic activation of MAL-phosphoinositide 3-kinase (PI3K) signalling by C5aR-TLR2 crosstalk stimulates the expression of pro-inflammatory genes, whilst inhibiting actin polymerisation to impair phagocytosis. As such, the MYD88-dependent antibacterial response is redirected to a TLR2-MAL-PI3K signalling response to maintain inflammation. In macrophages, C5aR-TLR2

2+ crosstalk stimulates Gi-dependent intracellular Ca signalling to enhance P. gingivalis-stimulated cyclic adenosine monophosphate (cAMP)-protein kinase A (PKA) activity (Wang et al., 2010). This results in the inactivation of glycogen synthase kinase-3 (GSK3) and impairs nitric oxide production required for intracellular P. gingivalis killing. Additionally, P. gingivalis has been demonstrated to regulate the expression of selective pro-inflammatory cytokines by macrophages to manipulate the host immune response (Liang et al., 2011). C5aR signalling can synergise with TLR2 to stimulate the expression of TNF, IL-1 and IL-6, and thus stimulate periodontal inflammation and potentially generate nutrients from tissue breakdown. Concomitantly, C5aR-TLR2 crosstalk leads to ERK1/2-mediated IRF1 suppression, which selectively inhibits the expression of IL-12 and suppresses cell-mediated immunity. As such, P. gingivalis exploits C5aR-TLR2 crosstalk to subvert host defence, while the dysregulation of the host immune response via C5aR may generate a microenvironment that favours microbial dysbiosis. Consistently, C5aR-deficient mice were protected from P. gingivalis- induced alveolar bone resorption and dysbiosis of the commensal microbiota. Furthermore, C5aR-deficient mice were able to clear P. gingivalis more efficiently than wildtype mice (Wang et al., 2010). Therefore, the manipulation of C5aR not only promotes P. gingivalis survival, but has been proposed to also generate a “bystander” effect that protects otherwise susceptible bacteria from the host immune system, and thereby contribute to the development of dysbiosis (Hajishengallis et al., 2011; Liang et al., 2011; Maekawa et al., 2014).

1.7.1.2 Gingipain Proteases and Tissue Destruction In addition to targeting the immune system, P. gingivalis gingipain proteases can also directly cause tissue destruction by degrading extracellular matrix proteins, including fibronectin and laminin (Andrian et al. 2004; Ruggiero et al. 2013). The gingipain proteases also stimulate the upregulation of host-derived MMPs (e.g. MMP-2), which enhances the degradation of the extracellular matrix and promotes periodontal ligament detachment (Andrian et al., 2007;

Grayson et al., 2003). Moreover, the wound healing response is also compromised as a result of the gingipain protease-mediated disruption of the coagulation cascade. The gingipain proteases can proteolytically process and activate kallikrein and bradykinin, which increases vascular permeability (Hinode et al., 1992; Imamura et al., 1994). Taken together, the effects exerted by the gingipain proteases allow P. gingivalis to subvert host defences and inflict host tissue destruction, which ultimately contribute to the pathogenesis of chronic periodontitis.

Fimbriae P. gingivalis produces two types of fimbriae: FimA (major fimbriae) and Mfa1 (minor fimbriae), which extend as filamentous protrusions from the surface of the bacterium (Amano et al., 2004). Genetic variations of FimA are classified as type I, Ib, II, III, IV and V. Patients with chronic periodontitis have been found to have higher levels of P. gingivalis strains that express type II FimA, which has been characterised as having the highest adhesive and invasive capacity for human epithelial cells (Amano et al., 2004; Nakagawa et al., 2002). The fimbriae facilitate P. gingivalis colonisation by enhancing co-adhesion with other bacteria (e.g. S. gordonii and T. denticola) (Hashimoto et al., 2003; Park et al., 2005). The adhesive properties of P. gingivalis fimbriae has also been shown to facilitate host cell invasion of human epithelial cells to subvert immune detection (Nakagawa et al., 2002; Njoroge et al., 1997). Fimbriae have been shown to interact with 1 integrins on gingival epithelial cells to promote the initial adherence and subsequent invasion of P. gingivalis (Yilmaz et al., 2002). This interaction is associated with the formation of focal adhesions and paxillin phosphorylation, which facilitates cytoskeletal rearrangement necessary for P. gingivalis invasion (Yilmaz et al., 2002, 2003).

Fimbriae also contribute to the manipulation of the host immune response by inducing signalling crosstalk between innate immune receptors. For example, P. gingivalis can redirect TLR2 and CD11b/CD18 receptor signalling in macrophages as a means of immune subversion (Hajishengallis and Lambris, 2010). Fimbriae-mediated interaction with TLR2 stimulates MYD88-mediated NF-B activation, enhancing pro-inflammatory gene expression (Hajishengallis et al., 2009). Concurrently, TLR2 initiates “inside-out” signalling through PI3K to promote CD11b/CD18 activation (Hajishengallis et al., 2009; Harokopakis and Hajishengallis, 2005). Activated CD11b/CD18 can signal via ERK1/2 to downregulate IL-12 p35/p40 expression, and thus suppress IFN- expression required for the cell-mediated immune clearance of P. gingivalis (Hajishengallis et al., 2007). Consistently, CD11b/CD18-deficient mice were demonstrated to clear P. gingivalis infection more efficiently than wildtype mice (Hajishengallis et al., 2007). TLR2 and CXCR4 signalling in macrophages is also exploited by P. gingivalis to suppress host defence. P. gingivalis fimbriae engage with CXCR4, which signals via cAMP-dependent protein kinase A (PKA) to suppress TLR2-mediated NF B activation,

45 resulting in reduced TNF and nitric oxide production (Hajishengallis et al., 2008). Mice treated with the CXCR4 antagonist, AMD3100, exhibited increased levels of nitric oxide and constrained P. gingivalis systemic infection more efficiently than untreated mice (Hajishengallis et al., 2008). As such, P. gingivalis fimbriae not only promote bacterial invasion, but are also involved in manipulating host immune signalling to impede bacterial clearance.

Atypical Lipopolysaccharides P. gingivalis has also been shown to antagonise the host immune response by producing atypical and heterogeneous forms of LPS containing different lipid A moieties (Dixon and Darveau, 2005; Reife et al., 2006). The post-translational modification to the lipid A of P. gingivalis LPS confers its ability to act as either a weak TLR4 agonist (in comparison to other Gram-negative species, such as E. coli) or antagonist to suppress TLR4-mediated immune responses (Domon et al., 2008; Reife et al., 1995). The acylation and phosphorylation pattern of the lipid A moiety of P. gingivalis LPS exhibits different PRR-activating capacities, whereby mono/di-phosphorylated, penta-acylated LPS weakly activates TLR4; non-phosphorylated, tetra-acylated LPS does not activate TLR4, and mono-phosphorylated, tetra-acylated LPS antagonises TLR4 activation. The modifications of P. gingivalis LPS is attributable to lipid A 1’-phosphatase (PG1773) and lipid a 4’-phosphatase (PG1587) activity under different haemin conditions (Al-Qutub et al., 2006; Coats et al., 2009). Under haem-limiting conditions, P. gingivalis utilises lipid A 1’ and 4’- phosphatase to generate lipid A species that weakly activate TLR4. Conversely, lipid A 1’-phosphatase activity is suppressed under conditions of high haem availability and leads to the production of TLR4-antagonistic lipid A species. In vitro studies with a P. gingivalis phosphatase mutant that only express the antagonistic lipid A species revealed that the mutant inhibited the production of pro-inflammatory mediators (e.g. TNF and IL-6) and suppressed caspase-11-dependent non-canonical inflammasome activation (Slocum et al., 2014). The mutant also exhibited increased survival in macrophages. Interestingly, apolipoprotein-deficient mice orally infected with the same P. gingivalis phosphatase mutant had no effect on oral inflammation or alveolar bone loss. Instead, the mice infected with the P. gingivalis phosphatase mutant were found to be more prone to developing vasculature inflammation associated with increased macrophage infiltration. The phosphatases have also been shown to be important for bacterial colonisation, in a ligature-induced rabbit model of periodontitis (Zenobia et al., 2014). Taken together, these studies suggest that the suppression of TLR4 by P. gingivalis may cause dysbiosis and contribute to the induction of inflammation at sites distant from the initial infection, and thus potentiate the progression of systemic disease (e.g. atherosclerosis).

SerB Phosphatase P. gingivalis expresses SerB, a phosphoserine phosphatase that is part of the haloacid dehydrogenase family of hydrolytic dehydrogenases. P. gingivalis SerB has been shown to be involved in mediating host cell invasion and immune subversion. Gene expression studies (e.g. microarray analysis) with gingival epithelial cells suggested that SerB is involved in regulating the expression of cytoskeletal-associated proteins (e.g. vinculin and paxillin) as well as MAPK-related proteins (e.g. Cdc42) (Hasegawa et al., 2008). Functional studies revealed that recombinant SerB can cause the disassembly of the actin by dephosphorylating cofilin, a regulator of actin stability (Moffatt et al., 2012). The reorganisation of host cytoskeletal proteins by SerB appears to be important for cellular invasion, as SerB-deficient P. gingivalis exhibited impaired invasive capabilities and reduced survival in gingival keratinocytes (Hasegawa et al., 2008; Tribble et al., 2006). In addition, SerB is also involved in modulating the host immune response. SerB can inhibit IL-8 gene expression in human gingival epithelial cells by dephosphorylating Ser536 of the p65 subunit of NF-B (Takeuchi et al., 2013). Rats infected with a SerB-deficient P. gingivalis displayed reduced alveolar bone loss, although the inflammatory responses were similar between rats infected with SerB-deficient and wildtype P. gingivalis (Bainbridge et al. 2010). However, greater neutrophil infiltration into the junctional epithelium in rats infected with SerB-deficient P. gingivalis suggests that SerB impairs innate immune defences by inhibiting the recruitment of neutrophils (Bainbridge et al. 2010). As such, SerB supports P. gingivalis immune evasion by mediating cellular invasion and suppressing NF-B activation.

1.8 Research Objectives The breakdown of host-microbe (plaque) homeostasis is central to the pathogenesis of chronic periodontitis. Recent research strongly suggests that P. gingivalis plays a critical role in promoting the breakdown of homeostasis. Therefore, defining the interactions between host- and P. gingivalis-derived immunomodulatory factors is crucial for understanding the possible causes of dysbiosis and inflammation in chronic periodontitis. The overall objective of this project was to identify and characterise novel host-P. gingivalis interactions. The specific aims of this project were to:

1. Study the regulation of the chemokine CXCL14 in response to P. gingivalis.

2. Investigate the functions of CXCL14.

3. Identify and characterise potential P. gingivalis TIR domain-containing proteins (Tcp).

4. Study the ability of the P. gingivalis Tcp, PG0382, to modulate the host immune response.

Materials and Methods

2.1 Materials Tissue culture reagents All plasticware was obtained from Falcon (USA). Keratinocyte serum-free medium (K-SFM), Dulbecco’s Modified Eagle Medium (DMEM), Glutamax-1™, penicillin/streptomycin, bovine pituitary extract (BPE), human epidermal growth factor (EGF), Gibco® 10X phosphate buffered saline (PBS), 0.05% Trypsin-EDTA, and foetal bovine serum (FBS) were purchased from Thermo Fisher (USA).

Bacterial culture reagents Bacto™ brain heart infusion (BHI) was purchased from BD Biosciences (USA). Blood Agar Base No. 2 and Agar Bacteriological Agar No. 1 were purchased from Oxoid (United Kingdom). Defibrinated horse blood was purchased from Equicell (Australia). Menadione, haemin, and L-cysteine hydrochloride were purchased from Sigma-Aldrich (USA).

General reagents and chemicals FSL-1 was purchased from Invivogen (USA). ProLongTM Gold Antifade reagent containing 4’6’-diamidino-2-phenylindole (DAPI) stain was purchased from Life Technologies (USA). Recombinant human and mouse CXCL14 were purchased from R&D Systems (USA). Human CXCL14 ELISA kit was purchased from Abcam (UK). The following chemicals were purchased from Sigma-Aldrich (USA): β-mercaptoethanol, isopropanol, calcium chloride, and goat serum.

Molecular biology reagents The following siRNAs were purchased from Dharmacon (USA): ON-TARGETplus non-targeting control siRNA, ON-TARGETplus IRF6, ON-TARGET plus IRAK-1, ON-TARGETplus PAR-1, ON-TARGETplus PAR-2, ON-TARGETplus PAR-3, ON-TARGETplus PAR-4, and ON-TARGETplus TLR2. Opti-MEM I reduced serum medium, and Lipofectamine RNAiMAX were purchased from Life Technologies (USA). The Reliaprep™ RNA miniprep system, Reliaprep™ RNA Tissue Miniprep system, GoScript™ reaction buffer, random primers, nucleotide mix, recombinant RNasin® ribonuclease inhibitor, GoScript™ reverse transcriptase, GoTaq® probe qPCR master mix, CXR reference dye, and FuGENE® 6 transfection reagent were purchased from Promega (USA).

Molecular cloning PCR primers, SYBR safe DNA gel stain and AxyPrep™ plasmid miniprep kit, and chemically competent E. coli DH5were purchased from Life Technologies (USA). Pfu DNA polymerase, and 1 kb DNA ladder were purchased from Promega (USA). The 100 bp DNA ladder was purchased from Bioline (USA). Restriction enzymes, calf intestinal phosphatase (CIP), and T4

DNA Ligase were purchased from New England Biolabs (USA). The DNeasey blood & tissue kit, MinElute PCR purification kit, and EndoFree® plasmid maxi kit were purchased from Qiagen (Germany). The UltraClean™ tissue & cells DNA isolation kit and UltraClean™15 DNA purification kit were purchased from MO BIO laboratories (USA). Ampicillin and erythromycin were purchased from Sigma-Aldrich (USA).

Quantitative real-time PCR probes Pre-developed TaqMan assays were purchased from Life Technologies (USA). The following probe sets for human genes were used: CCL20 (Hs01011368_m1), CXCL14 (Hs01557413_m1), IL-8 (Hs00174103_m1), IL-36G (Hs00219742_m1), IRAK-1 (Hs01018347_m1), MYD88 (Hs01573837) IRF6 (Hs00196213_m1), PAR-1 (Hs00169258_m1), PAR-2 (Hs00608346_m1), PAR-3 (Hs00187982_m1), PAR-4 (Hs01006385_g1), TATA box binding protein (TBP) (Hs00427620_m1), TNF (Hs0113624_g1), and TLR2 (Hs00152932_m1). The following probe sets for mouse genes were used: CCL2 (Mm00441242_m1), CXCL1 (Mm04207460_m1), IL-6 (Mm00446190_m1), IL-10 (Mm01288386_m1), HPRT (Mm00446968_m1), MAL (Mm00446502_m1), and TNF (Mm00443258_m1).

SDS-PAGE and Western Blotting The Bio-Rad protein assay dye reagent was purchased from Bio-Rad (USA). Complete™ EDTA- free protease inhibitors were purchased from Roche (Switzerland). Bovine serum albumin (BSA) was purchased from Sigma Aldrich (USA). NuPAGE™ Novex® Bis-Tris precast gels, NuPAGE™ LDS sample buffer, NuPAGE™ MES buffer, NuPAGE™ MOPS buffer, and Novex® Sharp pre-stained protein standards were purchased from Life Technologies (USA). Immobilon-P PVDF Transfer Membrane and Whatman® filter paper were purchased from Millipore (USA). Enhanced chemiluminescence (ECL) reagent was obtained from GE Healthcare (USA).

Antibodies The AlexaFluor®-488 conjugated goat anti-rabbit, and AlexaFluor®-594 conjugated goat anti-mouse antibodies were purchased from Life Technologies (USA). The rabbit monoclonal anti-FLAG antibody and anti-FLAG M2 affinity gel were from Sigma-Aldrich (USA). The mouse monoclonal anti-V5 antibody was purchased from Invitrogen (USA). The mouse monoclonal anti-phospho-ERK1/2, rabbit polyclonal anti-p38, and rabbit polyclonal anti-phospho-p38 antibodies were purchased from Cell Signalling Technology (USA). The rabbit polyclonal anti-ERK2 antibody was purchased from Santa Cruz (USA). The HRP-conjugated swine polyclonal anti-rabbit and rabbit polyclonal anti-mouse antibodies were purchased from Dako (Denmark). The following antibodies were purchased from BD biosciences (USA): mouse monoclonal anti-HSP90, FITC-conjugated rat monoclonal anti-mouse CD45, PE-conjugated rat

50 monoclonal anti-mouse F4/80, FITC-conjugated rat monoclonal anti-mouse Ly6G, PE-Cy7- conjugated rat monoclonal anti-mouse CD86, and rat monoclonal anti-mouse CD16/CD32. The Per-CP-Cy5.5-conjugated rat anti-mouse Ly6C was purchased from Biolegend (USA).

2.2 In vitro methods Cell culture 2.2.1.1 OKF6/TERT-2 cells Human telomerase-immortalised OKF6/TERT-2 oral epithelial cells, (were generously provided by Professor James Rheinwald (Harvard Medical School, Cambridge, MA), and hereafter referred to as OKF6 cells) (Dickson et al., 2000) were cultured in keratinocyte serum-free medium

(K-SFM) supplemented with 0.4 mM CaCl2, 2 mM GlutaMax, 50 U/ml penicilllin, 50 µg/ml streptomycin, 25 µg/ml bovine pituitary extract, and 0.2 ng/ml EGF. Cells were cultured at 37 °C in a humidified atmosphere of 5% CO2 and passaged every 2-3 days, as required.

2.2.1.2 HEK293T cells HEK293T cells (Graham et al., 1977) were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% (v/v) FBS, 2 mM GlutaMax, and 50 U/ml penicillin and 50

µg/ml streptomycin. The cells were cultured at 37 ˚C in a humidified atmosphere of 5% CO2 and passaged 2-3 days, as required.

2.2.1.3 RAW 264.7 cells RAW 264.7 mouse macrophages (Raschke et al., 1978) were cultured in DMEM supplemented with 10% (v/v) FBS, 2 mM GlutaMax, and 50 U/ml penicillin and 50 µg/ml streptomycin. The cells were cultured at 37 ˚C in a humidified atmosphere of 5% CO2 and passaged every 2-3 days, as required.

Bacterial strains and culture conditions 2.2.2.1 P. gingivalis Freeze-dried cultures of P. gingivalis ATCC 33277 and P. gingivalis W50 were obtained from the culture collection of the Melbourne Dental School, University of Melbourne. P. gingivalis KDP136 was kindly gifted by Professor Koji Nakayama (Nagasaki University, Graduate School of Biomedical Sciences). P. gingivalis strains were maintained on 10% defibrinated horse blood agar base (HBA) supplemented with 5 µg/ml menadione at 37 ˚C in anaerobic conditions (15%

CO2, 5% H2, and 80% N2). The gingipain protease-deficient mutant (KDP136) was created by homologous recombination, whereby the rgpA and rgpB gene in the P. gingivalis Kgp-deficient mutant (KDP129) were replaced by a tetracycline-resistance cassette (Shi et al., 1999). Hence, tetracycline was used to maintain phenotype of the bacterial mutant. Bacteria were passaged

51 weekly on agar plates for a maximum of eight passages, before returning to frozen stocks. Bacterial colonies were grown in batch culture in Brain Heart Infusion (BHI) broth supplemented with 5 mg/ml cysteine, 5 µg/ml haemin, and 5 µg/ml menadione.

2.2.2.2 Streptococcus strains Streptococcus gordonii ATCC 35105 and Streptococcus sp. OT058 were obtained from the culture collection of the Melbourne Dental School, University of Melbourne. S. gordonii and S. sp OT058 were cultured on 10% defibrinated HBA, and maintained at 37 ˚C in aerobic and anaerobic conditions, respectively. Batch cultures were grown in BHI broth supplemented with 5 µg/ml haemin.

2.2.2.3 E. coli E. coli was cultured on Luria-Bertani (LB) agar, and maintained at 37 °C in aerobic conditions. Batch cultures were grown in LB broth at 37 ˚C with agitation (200 rpm) on an orbital shaker.

Challenging of OKF6 cells and RAW264.7 cells with P. gingivalis P. gingivalis cultures were grown to mid to late exponential phase (corresponding to an optical density of 0.6-0.8 measured at 650 nm) and harvested by centrifugation at 8,000 g for 20 min at 4 °C. The pelleted bacteria were suspended in antibiotic-free K-SFM. The bacteria were diluted accordingly in K-SFM or DMEM, and added to OKF6 cells or RAW264.7 cells, respectively, to achieve a bacterium-to-cell ratio of 100:1. Cells challenged with P. gingivalis were incubated in a humidified atmosphere of 5% CO2 at 37 ˚C for up to 24 h.

RNA interference-mediated gene silencing A reverse-transfection protocol was used for siRNA transfections. siRNAs were diluted to 120 nM with 100 µl Opti-MEM I reduced serum medium (Life Technologies). The diluted siRNA was mixed with 100 µl Opti-MEM I reduced serum medium containing 1 µl Lipofectamine RNAiMAX transfection reagent, and incubated at room temperature for 15 min. The transfection cocktail was placed in 12-well plates, followed by the addition of 2×105 OKF6 cells. The medium was replaced 24 h later, and the cells were challenged with P. gingivalis 48 h post-transfection.

RNA purification Total RNA was purified using the ReliaPrep RNA cell miniprep system. Cells seeded in a 12-well plates were lysed with 250 μl of BL lysis buffer containing 1-thioglycerol. Genomic DNA was subsequently sheared by pipetting approximately 10 times. Subsequently, 85 μl of isopropanol was added into the cell lysate. The cell lysate was applied to the RNA purification column and centrifuged at 12,000 rpm for 30 sec at room temperature. The column was washed with 500 μl of RNA Wash solution at 12,000 rpm for 30 sec at room temperature. Next, 30 μl of DNase I

Incubation Mix was added to the column for 15 min at room temperature to degrade contaminating genomic DNA. Subsequently, the column was washed with 200 μl of Column Wash solution, followed by 500 μl of RNA Wash solution in succession at 12,000 rpm for 30 sec. Finally, the RNA purification column was washed with 300 μl of RNA Wash solution at 12,000 rpm for 2 min. The RNA was eluted with 20 μl of nuclease-free water by centrifugation at 12,000 rpm for 2 min. RNA concentration and purity were measured with a NanoDrop Lite Spectrophotometer (Thermo Fisher). The ratio of absorbance at 260 nm and 280 nm was used to assess the RNA purity, and a ratio of >2.05 was considered acceptable.

Reverse transcription

Total RNA (typically 500 ng per reaction) was incubated with 0.5 µg Oligo(dT)15 and 0.5 µg random primers at 70 ˚C for 5 min, and then chilled on ice for 5 min. The RNA was reverse transcribed into cDNA using GoScript Reverse Transcriptase with a BioRad T100™ Thermal Cycler (BioRad, USA) under the following cycling conditions: annealing at 25 ˚C for 5 min, extension at 42 ˚C for 45 min, and inactivation at 70 ˚C for 15 min.

Quantitative real-time PCR Quantitative real-time PCR was performed in duplicate using a QuantStudioTM 7 Flex Real-Time PCR system under the following cycling conditions: 50 ˚C for 2 min, 95 ˚C for 10 min, followed by 40 cycles of 95 ˚C for 15 s and 60 ˚C for 1 min. Ten ng of cDNA was used per 10 μl reaction. The data were normalised against the TBP or HPRT gene, and changes in gene expression were calculated using the ΔΔCt method (Pfaffl, 2001). When appropriate, absolute quantification of target gene mRNA levels were obtained as 1/(2^ΔCT).

CXCL14 Enzyme-linked immunosorbent assays (ELISA) CXCL14 protein standards and experimental samples were added to a 96-well micro plate with anti-CXCL14 capture antibody and incubated at 37 ˚C for 1.5 h. The contents of the wells were discarded and a biotinylated anti-CXCL14 antibody was added and incubated at 37 ˚C for 1 h. Thereafter, the wells were washed thrice with PBS, and an avidin-biotin-peroxidase complex added and the plate was incubated at 37 ˚C for 30 min. The plates were again washed with PBS, and 3,3’,5,5’-Tetramethylbenzidine (TMD) substrate solution was subsequently added and incubated in the dark at 37 ˚C for 30 min. Colour development was inhibited by the addition of the stop solution. Absorbance was measured at 450 nm with a PerkinElmer 1420 Multilabel Counter VICTOR3™. Background correction was applied by subtracting blank absorbance from sample readings.

In vitro proteolytic degradation of EGF and CXCL14 Purified lysine-specific P. gingivalis gingipain proteinase Kgp was generously provided by Dr. Laila Huq (Huq et al., 2013). Digestion of EGF and CXCL14 was performed at an enzyme: substrate ratio of 1:25. Purified Kgp was first activated in 200 mM HEPES (pH 7.6), 4 mM CaCl2, and 10 mM cysteine for 5 min at 37 ˚C. The substrate (EGF or CXCL14) was then added and incubated at 37 ˚C. Aliquots of the reaction mixture were removed and Kgp activity was inhibited by the addition of 2 mM N-Tosyl-L-lysine chlormethyl ketone hydrochloride (TLCK). The aliquots were then subjected to SDS-PAGE and/or analysis by mass spectrometry.

LTQ Orbitrap Elite mass spectrometry Kgp-digested recombinant CXCL14 was desalted using C18 zip-tips (Millipore). Prior to sample loading, the zip-tip was hydrated with 50% (v/v) acetonitrile and 0.1% (v/v) trifluoroacetic acid, and then thrice washed with 0.1% (v/v) trifluoroacetic acid. The total volume of the samples was passed through the zip-tips five times, and the zip-tips were then washed five times with 0.1% (v/v) trifluoracetic acid. Peptides were eluted with 30 % (v/v) acetonitrile containing 0.1% (v/v) trifluoracetic acid. The eluted samples were frozen with liquid nitrogen and subsequently vacuum dried in a Digital Series SpeedVac System™ (Thermo Fisher). Samples were analysed by an LTQ Orbitrap Elite mass spectrometer (Thermo Scientific) with a nanoESI source interfaced with an Ultimate 3000 RSLC nano-HPLC (Thermo Scientific). The peptides were loaded onto the enrichment column at an isocratic flow of 5 μl/min in 3% (v/v) acetonitrile containing 0.1% (v/v) formic acid for 5 min before the enrichment column was switched in-line with the analytical column. The eluents were used for the LC were 0.1% (v/v) formic acid (Solvent A) and 100% (v/v) acetonitrile in 0.1% (v/v) formic acid (Solvent B). Separation was performed with a gradient of 6–80% Solvent B for 53 min. The mass spectrometer was operated in the data-dependent mode with a nano-ESI spray voltage of 2.0 kV, capillary temperature of 250 ˚C, and S-lens RF value of 55%. All spectra were acquired in positive mode with full scan MS spectra scanning from m/z 300 – 1650 in the FT mode at 240,000 resolution after accumulating to a target value of 1.0e6. The top 10 most intense precursor ions were subjected to high energy collision induced dissociation (HCD) with a normalised collision energy of 35, and activation time of 0.1 ms. Dynamic exclusion with 2 repeat counts over 30 sec, and exclusion for 70 sec was applied. The Mascot MS/MS ions search was performed using the following settings: Uniprot database, Lys-C, 1 missed cleavage, 10 ppm peptide tolerance, and 0.2 Da MS/MS tolerance.

Antibacterial assays Mid to late exponential phase bacteria were harvested by centrifugation and washed with PBS. The bacteria (5×104 ) were suspended in incubation buffer (10 mM Tris-HCl, 5 mM glucose, pH 7.4) and incubated with CXCL14 for 1 h at 37 ˚C. Serial dilutions of P. gingivalis, S. gordonii, and S. sp. OT058 were plated on HBA, while E. coli was plated on LB agar plates. Bacterial cell numbers were enumerated and % killing was expressed as: [1 - (number of colonies with CXCL14 incubation)/ (number of colonies incubated with vehicle control)] × 100.

Wound healing assay OKF6 cells were seeded into 12-well plates and cultured to 90% confluency. A sterile pipette tip was used to generate wounds across the cell monolayer. The cells were washed with PBS to remove detached cells, and replaced with K-SFM containing stimulants. Images were acquired using an Axio Scope phase contrast microscope. Percentage gap closure was expressed as [1 - (area of gap measured at specific time)/ (area of gap measured at 0 h)] × 100.

Transfection of HEK293T cells HEK293T cells were seeded at a density of 1×105 cells/cm2 of culture vessel. The cells were transfected the next day using FuGENE 6 transfection reagent. Briefly, 1 µg of plasmid DNA was diluted in 50 µl serum-free DMEM containing 3 µl FuGENE 6 transfection reagent. The transfection cocktail was incubated at room temperature for 15 min, and then added dropwise to the cell monolayer. The total amount of plasmid in each transfection was kept constant using parental empty plasmid. The list of plasmids used are listed in Table 2.1.

Cell lysis OKF6 and HEK293T cells were washed twice with ice-cold PBS, and then lysed with NP-40 lysis buffer (20 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1% (v/v) Nonident P-40, 10% (v/v) glycerol, 1 mM sodium orthovanadate, 10 mM β-glycerol phosphate, and 10 mM sodium fluoride) supplemented with protease inhibitors on ice for 30 min. The lysates were clarified by centrifugation at 12,000 rpm for 10 min at 4 ˚C. The protein concentration of the lysates were determined by the Bradford assay (Bradford, 1976) using the Bio-Rad protein assay dye reagent. Thereafter, the lysates were either used for co-immunoprecipitation assays, or stored at -80 ˚C for subsequent analysis by SDS-PAGE.

Co-immunoprecipitation Assay Lysates were adjusted to a protein concentration of 1 mg/ml in a total volume of 1 ml ice-cold lysis buffer, and incubated with 10 µl of anti-FLAG beads at 4 ˚C overnight with constant mixing. The beads were washed four times with ice-cold lysis buffer. Following the final wash, the beads

55 were eluted by heating at 70 ˚C for 10 min in 30 µl of NuPAGE LDS sample buffer containing 50 mM DTT. The eluates were then subjected to SDS-PAGE and Western blotting.

SDS-PAGE Samples were prepared in NuPAGE LDS sample buffer and heated at 70 ˚C for 10 min. Approximately 20 µg total protein was loaded per well onto 10% NuPAGE Bis-Tris gels. Electrophoresis was performed at 100 V for 2 h in 2-(N-morpholino) propanesulfonic acid (MOPS) (50 mM MOPS, 50 mM Tris (pH 7.7), 0.1 % (w/v) SDS, 1 mM EDTA) or 2-(N-morpholino) ethanesulfonic acid (MES) (50 mM MES, 50 mM Tris (pH 7.3), 0.1% (w/v) SDS, 1 mM EDTA) running buffer.

Western Blotting Proteins were transferred to polyvinylidene fluoride (PVDF) membranes at 500 mA for 90 min in transfer buffer (25 mM Tris-HCl, 200 mM glycine, 20% v/v methanol). Membranes were rinsed in Tris-buffered saline/Tween-20 (TBST: 20 mM Tris-HCl (pH 7.4), 150 mM NaCl, 0.1% (v/v) Tween-20), and then blocked with 3% (w/v) BSA in TBST for 1 h at room temperature. Membranes were incubated with primary antibodies (diluted in 1% (w/v) BSA in TBST) at 4 ˚C overnight. The membranes were washed with TBST over 1 h with several changes of buffer, and then incubated with the appropriate secondary antibody (diluted in 1% (w/v) BSA in TBST) for 1 h at room temperature. The membranes were again washed with TBST. Immunoreactive protein bands were visualised with ECL reagent and imaged with a LAS3000 imager (Fujifilm). Before re-probing membranes with another antibody, bound antibodies were removed by incubating the membrane in stripping buffer (50 mM Tris-HCl (pH 7.4), 2% (w/v) SDS, 100 mM β-mercaptoethanol) for 15 min at 55 ˚C. The membrane was washed for 1 h with TBST, and then blocked with 3% (w/v) BSA in TBST. Densitometric analysis was conducted using the ImageQuant software.

Immunofluorescence staining and confocal microscopy Glass coverslips were sterilised with 80% (v/v) ethanol for 10 min. The coverslips were then transferred to 24-well plates and allowed to air-dry. HEK293T cells were seeded onto the coverslips at a density of 5×104 cells/well and transfected using FuGENE6 (refer to Section 2.2.13 for further details). Twenty-four h later, the cells were washed with PBS, and fixed with 4% (w/v) paraformaldehyde (PFA) in PBS (pH 7.4) at room temperature for 30 min. The cells were permeabilised with 0.1% (v/v) Triton X-100 in PBS for 5 min, followed by blocking with 5% (v/v) goat serum in PBS for 1 h at room temperature. Thereafter, the cells were incubated in the appropriate primary antibody (diluted in 5% (v/v) goat serum in PBS) for 1 h at room temperature. Following three washes with PBS, the cells were incubated with either an

AlexaFluor®-488 conjugated goat anti-rabbit antibody or an AlexaFluor®-594 conjugated goat anti-mouse antibody for 1 h at room temperature in the dark. The cells were again washed thrice with PBS, and then mounted onto microscope slides using ProLong™ Gold antifade reagent with 4’,6-diamidino-2-phenylindole (DAPI) stain. Coverslips were cured overnight in the dark. Confocal images were acquired with a Leica SP5 microscope (Leica Microsystems).

Construction of PG0382 mammalian expression vectors 2.2.19.1 Polymerase Chain Reaction P. gingivalis W50 genomic DNA was isolated using a Qiagen DNeasy blood and tissue kit according to the manufacturer’s instructions. The PG0382 gene was amplified by Polymerase Chain Reaction (PCR) using Pfu DNA polymerase and custom-designed primers listed in Table 2.2. The thermal cycling conditions were as follow: initial denaturation at 95 ˚C for 4 min, followed by 35 cycles of denaturation at 95 ˚C for 30 s, annealing at 62 ˚C for 1 min, elongation at 72 ˚C for 2 min, and a final elongation at 72 ˚C for 10 min. The annealing temperature was calculated as: Tm – 10 ˚C, where Tm refers to the melting temperature of the primers. PCR products were purified using a MinElute PCR kit according to the manufacturer’s instructions. Thereafter, the purified PCR products and the mammalian expression vectors pEF-FLAG and pEF-V5, were digested with MluI. The linearised vector was dephosphorylated by incubation with 10 U of calf intestinal phosphatase per µg DNA for 1 h at 37 ˚C. The digested PCR products and vectors were separated on a 1% agarose gel, excised from the gel, and purified using the Ultraclean DNA purification kit according to the manufacturer’s instructions. The PCR product was ligated into the linearised vectors overnight at a 3:1 molar ratio (insert:vector) using T4 DNA ligase.

2.2.19.2 Bacterial transformation Ten ng of plasmid DNA was first gently mixed with chemically-competent E. coli DH5α (50 μl) on ice. The bacteria were then subjected to heat-shock at 42 ˚C for 20 sec, followed by incubation on ice for 2 min. The bacteria were inoculated into pre-warmed LB broth and incubated at 37 ˚C for 1 h with constant shaking (200 rpm). The bacteria were then spread onto LB agar plates supplemented with 100 µg/ml ampicillin and incubated overnight at 37 ˚C. Single colonies were randomly selected and grown in 2 ml LB broth supplemented with 100 µg/ml ampicillin overnight at 37 ˚C. Miniprep plasmid DNA was purified using the Axygen Plasmid DNA Purification Miniprep kit. Positive clones were identified by restriction digests of the purified DNA. Large-scale (maxi prep) plasmid preparations were performed by inoculating 100 µl of the bacterial culture into 200 ml of LB broth supplemented with 100 µg/ml ampicillin and incubating overnight at 37 ˚C with constant shaking (200 rpm). Plasmid DNA was purified using

57 an EndoFree Maxi kit. DNA sequencing of plasmids was performed at the Centre for Translational Pathology (The University of Melbourne).

Generation of P. gingivalis PG0382-deficient mutant (ΔPG0382) 2.2.20.1 Splice Overlap Extension Polymerase Chain Reaction (SOE PCR) A gene deletion cassette, comprised of the erythromycin-resistance gene ermF flanked with 5’ and 3’ sequences from PG0382, was constructed by SOE PCR. Three PCR products, namely, PG0382_IG5, PG0382_IG3, and ermF_PG0382 were generated using the indicated primer pairs and thermal cycling conditions (Table 2.3). The PCR products were run on a 1% agarose gel, excised from the gel, and then purified using the Ultraclean DNA purification kit according to the manufacturer’s instructions. The first-stage SOE PCR reaction (Table 2.4 - SOE PCR reaction 1), was undertaken to recombine PG0382_IG5 and ermF_PG0382. Primers PG0382_IG5 Fw and ermF Rv were added to the reaction at cycle 3 to amplify the recombined PCR products. The SOE PCR product generated was run on a 1% agarose gel; excised from the gel and purified using the Ultraclean DNA purification kit. The purified PCR product was used for a subsequent SOE PCR reaction with PG0382_IG3 (Table 2.4 - SOE PCR reaction 2). Primers PG0382_IG5 Fw and PG0382_IG3 Rv were added to the reaction at cycle 3 to generate the complete ermF inactivation cassette. The resulting SOE PCR product was run on a 1% agarose gel; excised from the gel and purified using the Ultraclean DNA purification kit.

2.2.20.2 Electroporation of P. gingivalis Two-hundred ml of P. gingivalis in early exponential growth phase (corresponding to an optical density of 0.5-0.6 measured at 650 nm) were harvested by centrifugation at 8,000 g for 20 min at 4 ˚C. The pelleted bacteria were washed with 200 ml of ice-cold electroporation buffer (10 %

(v/v) glycerol, 1 mM MgCl2) and re-pelleted by centrifugation at 8,000 g for 20 min at 4 ˚C. The cell pellet was suspended in 400 µl ice-cold electroporation buffer. Eighty microliters of cells were aliquoted into Eppendorf tubes containing 200 ng of ermF inactivation cassette. The tubes were incubated on ice for 5 min before being transferred to ice-cold 0.1-cm gap cuvettes and electroporated at 1.8 kV, 200 Ω, and 25 µFaradays. Following electroporation, the cells were suspended in 1 ml of BHI broth supplemented with 0.5 mg/ml cysteine and 5 µg/ml haemin, and incubated overnight anaerobically at 37 ˚C. The cells were centrifuged at 8,000 g for 5 min. Most of the supernatant (~900 µl) was discarded and cells were resuspended in the remaining 100 µl of supernatant and plated onto a HBA plate supplemented with 10 µg/ml of erythromycin for selection. P. gingivalis genomic DNA was purified using a Qiagen DNeasy blood and tissue kit according to the manufacturer’s instructions, and PG0382 deletion was confirmed by DNA sequencing performed by the Centre for Translational Pathology (The University of

Melbourne), with the following primer pairs: 5’ – TGC ATC AGT GTC TTT CGG TG – 3’ and 5 ‘– GTA TCC ATG GAT TCG TCT ATC AT – 3’.

Gingipain proteinase assay Mid exponential phase P. gingivalis were harvested by centrifugation at 8,000 g for 20 min at 4

˚C. The cell pellet was suspended in TC150 buffer (50mM Tris-HCl, pH 7.4, 5 mM CaCl2, and 150 mM NaCl) supplemented with 20 mM cysteine. Kgp- and Rgp-specific activity was assayed using the chromogenic substrates, N-p-tosyl-Gly-Pro-Lys p-nitroanilide (GPKNA), and N-benzoyl-L- arginine p-nitroanilide (L-BAPNA) respectively. GPKNA and L-BAPNA cleavage was monitored at 405 nm for 30 min (at 10 sec intervals) at 37 ˚C using a PerkinElmer 1420 Multilabel Counter VICTOR3™.

2.3 Bioinformatics methods Sequence alignments Multiple sequence alignments were conducted using the ClustalOmega algorithm (with default parameters) on the EMBL-EBI server with Jalview editing software. Sequence alignments were coloured using ClustalX colour scheme. Pairwise sequence alignments were conducted using the EMBOSS NEEDLE algorithm (with default parameters) on the EMBL-EBI server (https://www.ebi.ac.uk/Tools/psa/emboss_needle/).

Structural modelling PG0382 amino acid sequence was subjected to FUGUE analysis to identify structural homologies. FUGUE (http://mizuguchilab.org/fugue/) is an online service that compares query sequences to a database of structural profiles to identify structural homologs. The program calculates a level of confidence of resulting matches by utilising an environment-specific substitution tables and structure-dependent gap penalties to account for the local structural environment of sequence residues (Shi et al., 2001). A list of sequence-structure compatibility scores is generated from potential homologues. A 3D structure of PG0382 was modelled based on structural templates of proteins that were identified in FUGUE using SWISS-MODEL (https://swissmodel.expasy.org/). The predicted structures were visualised and edited using PyMOL.

2.4 Mouse studies Mice Animal studies were approved by the University of Melbourne Ethics Committee for Animal Experimentation (Ethics identification number: 1613985 and 1614048). C57BL/6 and BALB/C

59 mice and (6–12 weeks old) were housed at the Bio21 Institute Biological Research Facility. The mice had access to food and water ad libitum in alternating 12 h periods of light and a dark.

Intragingival injection C57BL/6 mice (6–12 weeks old) were anaesthetised with a mixture of ketamine (0.1 mg/g) and xylazine (0.01 mg/g) using a 26G needle. Recombinant mouse CXCL14 (0.5 µg or 2 µg) in PBS was injected into the gingiva of anaesthetised mice with a 31G needle. Control mice were injected with the same volume of PBS. Mice were killed at 24 or 48 h-post injection by CO2 gassing. The upper maxilla was dissected from the animal and the gingival tissue was stripped. Half of the gingival tissue was frozen on dry-ice and stored at -80 ˚C until used for gene expression analysis.

Mouse gingival RNA extraction RNA from mouse gingival tissue was purified with the Reliaprep RNA Tissue Miniprep System (Promega). Excised mouse gingival tissue was crushed in liquid nitrogen with an ice-cold mortar and pestle. The homogenate was then transferred to 500 µl of LBA lysis buffer containing 1-thioglycerol. An equal volume of RNA dilution buffer was added, and the lysates were mixed by vortexing. Subsequently, 85 μl of isopropanol was added into the cell lysate. The cell lysate was applied to the RNA purification column and centrifuged at 12,000 rpm for 1 min at room temperature. The column was washed with 500 μl of RNA Wash solution at 12,000 rpm for 30 sec at room temperature. Next, 30 μl of DNase I Incubation Mix was added to the column for 15 min at room temperature to degrade contaminating DNA. Subsequently, the column was washed with 200 μl of Column Wash solution, followed by 500 μl of RNA Wash solution in succession at 12,000 rpm for 30 sec. Finally, the RNA purification column was washed with 300 μl of RNA Wash solution at 12,000 rpm for 2 min. The RNA was eluted with 25 μl of nuclease- free water following centrifugation by 12,000 rpm for 1 min. RNA concentration and purity were measured with a NanoDrop Lite Spectrophotometer (Thermo Fisher). The ratio of absorbance at 260 nm and 280 nm was used to assess the RNA purity, and a ratio of >2.0 was considered acceptable.

Intraperitoneal infection P. gingivalis W50 and P. gingivalis ΔPG0382 cultures were grown to mid to late exponential phase (corresponding to an optical density of 0.6-0.8 measured at OD650) and harvested by centrifugation at 8,000 g for 20 min at 4 ˚C and suspended in ice-cold PBS. BALB/c mice (7 weeks old) were injected in the peritoneal cavity with 5×106 or 5×107 bacteria using a 26 G needle. Control mice were injected with the same volume PBS. Mice were killed 6 or 24 h post- injection by CO2 gassing. The intraperitoneal cavity was lavaged by injecting 5 ml ice-cold PBS,

60 gently massaging the mouse cavity, and collecting the fluid with a 21 G needle. The peritoneal cavity fluid was immediately processed for flow cytometric analysis.

Flow cytometric analysis Mouse peritoneal cavity fluid harvested was centrifuged at 300 g for 10 min at 4 ˚C. The cells were suspended in PBS and numerated. This was followed by staining with Fixable Viability Stain 700 (FVS700), used at 1:1000 dilution for 15 min on ice in the dark. The cells were centrifuged at 300 g for 5 min and washed twice with FACS buffer (2.5 % (v/v) FBS and 2 mM EDTA in PBS). The cells were then incubated with anti-mouse CD16/CD32 (1:500) on ice for 15 min. Subsequently, 5×105 cells were aliquoted into respective antibody cocktails (Table 2.5) and incubated on ice for 30 min. Following incubation, the cells were washed twice with FACS buffer, and then fixed with 2 % (w/v) PFA at 4 ˚C for 15 min. The cells were washed thrice with PBS and suspended in FACS buffer for analysis using a BD LSRFortessa X-20 (BD bioscience). A typical forward and side-scatter gate was set to exclude aggregates, and dead cells were discarded on the basis of the FVS700 staining. Data analysis was conducted using the FlowJo software (TreeStar Inc).

2.5 Statistical analysis Data combined from three or more independent experiments are presented as the mean ± standard error of mean (SEM). Where indicated in the figure legends, n is the number of biological replicates. Statistical analyses were performed using GraphPad Prism software version 6.01. Differences between two groups were evaluated using the Student’s t-test. For multiple comparisons, statistical analysis was performed by ANOVA with Dunnett’s or Sidak’s post-hoc test. A p-value ˂0.05 was considered to be statistically significant.

Table 2.1 List of plasmids used in this study.

Plasmid Description Source pFLAG-MAL Expresses FLAG-tagged mouse MAL. Dr. Ashley Mansell pFLAG-MYD88 Expresses FLAG-tagged human MYD88. Dr. Ashley Mansell pEF-V5-PG0382 Expresses V5-tagged P. gingivalis PG0382. This study Expresses V5-tagged P. gingivalis PG0382 pEF-V5-PG0382ΔTD truncated mutant lacking the TIR domain (Asp2 This study to Glu342).

Expresses V5-tagged P. gingivalis PG0382 TIR pEF-V5-PG0382TD Ben Huang domain (Lys341 to Gln490).

Table 2.2 PCR primers used for constructing PG0382 mammalian expression vectors.

Vector PCR primers 1 pEF-V5-PG0382 Fw 5’ – CGA CGC GTG ACT TGG CTG AAG TTT TTG TTA CTG AAG – 3’ Rv 5’ – CGA CGC GTC TAT TGA ACC TTA GAA ATT ATT CTT TC – 3’ pEF-V5-PG0382 ΔTD Fw 5’ – CGA CGC GTG ACT TGG CTG AAG TTT TTG TTA CTG AAG – 3’ Rv 5’ – CGA CGC GTT TCT TTA TAC AAC TTG TTC CAG TCG ATG – 3’

1 Bolded sequences correspond to MluI restriction sites

Table 2.3 PCR primers used to generate PCR products for constructing ermF cassette.

Genome Product Product name Primer 1 PCR primers 2,3 Thermal Cycling conditions position (nt) Description Cycle 1 Denaturation 95 °C 4 min PG0382_IG5 5’-GAA CAT GGC AAG ATT GCG GT-3’ Contains ermF Cycle 2 (4x) Fw sequence for SOE Denaturation 95 °C 30 s 412991 – PCR and PG0382 5’ Annealing 40 °C 1 min PG0382_IG5 413330 intergenic sequences Elongation 72 °C 1 min for homologous Cycle 3 (25x) PG0382_IG5 5’-GCA ATT TCT TTT TTG TCA TAT recombination. Denaturation 95 °C 30 s Rv GCA GTT AAA-3’ Annealing 50 °C 1 min Elongation 72 °C 1 min Cycle 1 Denaturation 95 °C 4 min PG0382_IG3 5’-AAA TTT CAT CCT TCG TAG TTA Contains ermF Cycle 2 (4x) Fw ATT ATA GAA AGG GGG ATT TA-3’ sequence for SOE Denaturation 95 °C 30 s 415107 - PCR and PG0382 3’ Annealing 42 °C 1 min PG0382_IG3 415097 intergenic sequences Elongation 72 °C 1 min for homologous Cycle 3 (25x) PG0382_IG3 5’-GAA GCC AAC CGA TAC CGA AT-3’ recombination. Denaturation 95 °C 30 s Rv Annealing 51 °C 1 min Elongation 72 °C 1 min Cycle 1 ° 5’-TTT AAC TGC ATA TGA CAA AAA Denaturation 95 C 4 min ermF Fw AGA AAT TGC-3’ Cycle 2 (4x) ermF cassette Denaturation 95 °C 30 s containing PG0382 Annealing 40 °C 1 min ermF_PG0382 N/A overlap sequences Elongation 72 °C 1 min 5’-CCT TTC TAT AAT TAA CTA CGA for SOE PCR. Cycle 3 (25x) ermF Rv ° AGG ATG AAA-3’ Denaturation 95 C 30 s Annealing 48 °C 1 min Elongation 72 °C 1 min

1 Fw indicates forward primer, and Rv indicates reverse primer. 2 Bolded sequences correspond to overlapping ermF sequence. 3 Underlined sequences correspond to overlapping PG0382 sequence.

Table 2.4 SOE PCR reaction thermal cycling conditions.

SOE PCR reaction 1 SOE PCR reaction 2 Cycle 1 Cycle 1 Denaturation 95 °C 4 min Denaturation 95 °C 4 min Cycle 2 (4x) Cycle 2 (4x) Denaturation 95 °C 30 s Denaturation 95 °C 30 s Annealing 42.5 °C 1 min Annealing 48.8 °C 1 min Elongation 72 °C 1 min Elongation 72 °C 1 min Cycle 3 Cycle 3 Hold 12 °C 10min Hold 12 °C 10min Cycle 4 (25x) Cycle 4 (25x) Denaturation 95 °C 30 s Denaturation 95 °C 30 s Annealing 48.8 °C 1 min Annealing 51.7 °C 1 min Elongation 72 °C 1 min Elongation 72 °C 1 min

Table 2.5 Flow cytometric analysis antibody cocktails.

Monocyte and granulocyte antibody cocktail Antigen Conjugate Dilution F4/80 PE 1:400 Ly6G FITC 1:500 CD86 PE-Cy7 1:200 Ly6C Per-CP-Cy5.5 1:200 Lymphocyte antibody cocktail Antigen Conjugate Dilution CD45 FITC 1:100 TCR PE-Cy7 1:50 CD4 APC 1:200 CD8 PE 1:50 CD19 Per-CP-Cy5.5 1:50 CD25 BV421 1:100

The regulation of CXCL14 in oral epithelial cells by Porphyromonas gingivalis

3.1 Introduction The barrier function of the oral epithelium is maintained by cycles of cell proliferation and differentiation of basal epithelial cells to replace the cells that are constantly lost at the surface of the epithelium. Mitogens, such as Epidermal Growth Factor (EGF), play important roles in governing the balance between cell proliferation and differentiation, and thus contribute to the maintenance of barrier function. In addition, the oral epithelium also provide host protection by secreting antimicrobial peptides (e.g. -defensins and cathelicidin) and stimulating host inflammation (Dale and Fredericks, 2005).

Importantly, the oral epithelium also serves as an active participant in host defence by initiating inflammation and communicating potential threats to immune cells through PRRs (e.g. TLRs and PARs) (Darveau et al., 2004; Uehara et al., 2005). Once activated, PRRs can stimulate the expression of chemokines to facilitate the host inflammatory response by recruiting a range of immune cells (e.g. neutrophils and macrophages) to the site of infection. Interestingly, EGF receptor (EGFR) signalling appears to play an important role in the differential regulation of chemokine expression by epithelial cells. For instance, EGFR signalling was shown to regulate the expression of IL-8, CCL2 and CXCL10 in epidermal keratinocytes (Mascia et al., 2003). The coordinated response mediated by chemokines through multiple regulatory pathways is essential for maintaining tissue homeostasis and preventing chronic inflammation. However, in the case of chronic periodontitis, the continual stimulation of the host immune response by a dysbiotic microbial community can inflict damaging effects on host tissues (Darveau, 2010). P. gingivalis drives this process by expressing virulence factors (e.g. gingipain proteases) to dysregulate the host immune response and promote microbial dysbiosis (Hajishengallis et al., 2012). For example, P. gingivalis virulence factors can antagonise the host immune response by dysregulating the expression of chemokines. Accordingly, disordered chemokine expression may stimulate a sustained inflammatory response and turn a normally host-protective response into one that causes pathology.

Our laboratory had previously undertaken a preliminary study to identify P. gingivalis- inducible genes in human oral epithelial cells (e.g. OKF6 cells). Specifically, an Open-Array human inflammation panel, which allows the mRNA levels of 586 inflammatory genes to be measured, was used to screen for genes whose expression is regulated by P. gingivalis. The gene encoding the orphan chemokine, CXCL14, was identified as one such gene. Therefore, the aim of this Chapter was to examine the regulation of CXCL14 in oral epithelial cells by P. gingivalis.

3.2 Results P. gingivalis stimulates CXCL14 expression in human oral epithelial cells in a TLR2-independent manner OKF6 cells, which are a non-transformed, telomerase-immortalised human oral epithelial cell line (Dickson et al., 2000), have been used in a number of studies to investigate the response of oral epithelial cells to bacteria (Giacaman et al., 2009; Macpherson et al., 2014). Therefore, OKF6 cells were considered an appropriate in vitro cell system to investigate the inflammatory response of human oral epithelial cells to P. gingivalis.

To validate the identification of CXCL14 as a P. gingivalis-inducible gene, OKF6 cells were challenged with P. gingivalis over a 24 h time-course and CXCL14 mRNA levels were then measured by Real-Time PCR. CXCL14 gene expression was found to be strongly upregulated 24 h-post challenge with P. gingivalis (Fig. 3.1A). The levels of CXCL14 protein in the cell culture supernatants were measured by ELISA to determine whether the upregulation of CXCL14 mRNA resulted in a corresponding increase in CXCL14 protein. Although CXCL14 protein was detected in the cell culture supernatants from cells that had not been challenged with P. gingivalis, CXCL14 protein levels were not increased by P. gingivalis challenge (Fig. 3.1B). Other studies have also reported difficulties in measuring cytokine/chemokine levels in cell culture supernatants due to their proteolysis by P. gingivalis gingipain proteases (Darveau et al. 1998; Stathopoulou et al. 2009; Zhang et al. 1999).

Our laboratory has recently demonstrated that TLR2, IRAK-1 and IRF6 function as a signalling axis to differentially regulate cytokine and chemokine expression in oral epithelial cells (Huynh et al., 2016; Kwa et al., 2014). Therefore, a role for these proteins in regulating P. gingivalis-inducible CXCL14 expression was investigated using a siRNA-mediated gene knockdown approach. The transfection of OKF6 cells with the TLR2 siRNA reduced TLR2 mRNA levels by >80% relative to cells transfected with the non-targeting control siRNA (Fig. 3.1C). The effect of TLR2 knockdown on the induction of CXCL14 expression by P. gingivalis was next evaluated. Notably, knockdown of TLR2 did not inhibit the upregulation of CXCL14 (Fig. 3.1D), suggesting that the regulation of CXCL14 gene expression in response to P. gingivalis was not mediated by the TLR2 signalling pathway. A complementary approach was also taken, whereby OKF6 cells were stimulated with the TLR2 agonist FSL-1. Consistent with the findings from the TLR2 gene silencing experiment, FSL-1 did not stimulate CXCL14 gene expression (Fig. 3.1E). To confirm that OKF6 cells are responsive to stimulation by FSL-1, the expression levels of the chemokine, CCL20, which is known to be a target of TLR2 signalling in human keratinocytes (Niebuhr et al., 2010), were measured. As shown in Figure 3.1F, FSL-1

70 strongly stimulated CCL20 gene expression. Together, these data suggest that P. gingivalis-inducible CXCL14 gene expression is regulated in a TLR2-independent manner.

A B

C D

E F

Figure 3.1 P. gingivalis stimulates CXCL14 gene expression in a TLR2-independent manner. (A-B) OKF6 cells were challenged with P. gingivalis at 100 MOI for the times indicated. CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mocked-challenged cells (n=3). (B) CXCL14 protein levels in the cell culture supernatants were measured by ELISA (n=3). (C-D) OKF6 cells were transfected with a non-targeting (-) or a TLR2 (+) siRNA for 48 h, and subsequently challenged with P. gingivalis at 100 MOI for 24 h. (C) TLR2 mRNA levels were measured by Real-Time PCR, and levels in cells transfected with the non-targeting siRNA control were arbitrarily assigned a value of 100% (n=3). (D) CXCL14 mRNA levels were measured by Real-Time PCR, and shown as a fold change relative to mock-challenged cells (n=3). (E-F) OKF6 cells were stimulated with FSL-1 (100 ng/ml) for the times indicated. (E) CXCL14 and (F) CCL20 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-stimulated cells (n=3). All data are presented as the mean ± SEM (* = p <0.05, *** = p <0.001). 71

Despite the above finding, roles for IRAK-1 and IRF6 in regulating CXCL14 gene expression were also investigated. Transfection of OKF6 cells with the IRAK-1 siRNA, which reduced IRAK-1 gene expression by >90% (Fig. 3.2A), did not inhibit the stimulation of CXCL14 gene expression by P. gingivalis (Fig. 3.2B). Similarly, gene silencing of IRF6 (Fig. 3.2C) did not inhibit P. gingivalis-inducible CXCL14 gene expression (Fig. 3.2D). Collectively, these data suggest that P. gingivalis-inducible CXCL14 gene expression in oral epithelial cells (e.g. OKF6 cells) is regulated in a TLR2-, IRAK-1-, and IRF6-independent manner.

A B

C D

Figure 3.2 P. gingivalis stimulates CXCL14 gene expression in an IRAK-1 and IRF6-independent manner. (A-B) OKF6 cells were transfected with a non-targeting (-) or a IRAK-1 (+) siRNA for 48 h and subsequently challenged with P. gingivalis at 100 MOI for 24 h. (A) IRAK-1 mRNA levels were measured by Real-Time PCR, and levels in cells transfected with the non-targeting siRNA control was arbitrarily assigned a value of 100% (n=3). (B) CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-challenged cells (n=3). (C-D) OKF6 cells were transfected with a non-targeting (-) or an IRF6 (+) siRNA for 48 h and subsequently challenged with P. gingivalis at 100 MOI for 24 h. (C) IRF6 mRNA levels were measured by Real-Time PCR, and levels in cells transfected with the non-targeting siRNA control was arbitrarily assigned a value of 100% (n=3). (D) CXCL14 mRNA levels were measured by Real-Time PCR and shown as a fold change relative to mock-challenged cells (n=3). All data are presented as the mean ± SEM (** = p < 0.01, *** = p < 0.001).

Gingipain proteases mediate the stimulation of CXCL14 expression by P. gingivalis The gingipain proteases are central to the ability of P. gingivalis to antagonise the host immune response (O’ Brien-Simpson et al., 2003). They not only stimulate the expression of cytokines and chemokines in oral epithelial cells, but can also degrade them once secreted (Darveau et al., 1998; Dommisch et al., 2007; Lourbakos et al., 2001), and thereby contribute to the dysregulation of the host immune response. Therefore, a role for the gingipain proteases in the stimulation of CXCL14 gene expression by P. gingivalis was investigated. OKF6 cells were challenged with either wildtype P. gingivalis or P. gingivalis KDP136, an isogenic Kgp/Rgp-deficient mutant (Shi et al., 1999), and CXCL14 gene expression was then measured. In contrast to wildtype P. gingivalis, the gingipain protease-deficient mutant did not stimulate CXCL14 gene expression (Fig. 3.3A). To confirm the importance of the gingipain proteases for P. gingivalis-inducible CXCL14 expression, OKF6 cells were challenged with P. gingivalis that had first been treated with the irreversible serine protease chemical inhibitor, N--Toysl-L-Lysine chloromethyl ketone hydrochloride (TLCK), to inhibit gingipain protease activity. Consistent with the data presented in Figure 3.3A, inhibition of gingipain protease activity with TLCK prevented the stimulation of CXCL14 gene expression by P. gingivalis (Fig. 3.3B). These data therefore suggest that the gingipain proteases are necessary for the stimulation of CXCL14 gene expression in OKF6 cells by P. gingivalis.

In addition to being attached to the cell-surface of P. gingivalis, the gingipain proteases are also found on P. gingivalis outer membrane vesicles (OMVs) (Gui et al., 2015). To determine whether OMV-associated gingipain proteases can likewise stimulate CXCL14 gene expression, OKF6 cells were treated with either cell-free culture supernatants derived from wildtype P. gingivalis or P. gingivalis KDP136. Kgp- and Rgp-specific activity in the cell-free culture supernatants were first determined by measuring the hydrolysis of chromogenic substrates, N-p-Tosyl-Gly-Pro-Lys 4-nitroanilide (GPKNA) and N-Benzoyl-L-arginine 4-nitroanilide hydrochloride (L-BAPNA), respectively. As shown in Figure 3.3C, supernatants derived from wildtype P. gingivalis exhibited Kgp activity, whilst Kgp activity was not detected in supernatants derived from P. gingivalis KDP136. Similarly, Rgp activity was only detected in supernatants derived from wildtype P. gingivalis and not P. gingivalis KDP136 (Fig. 3.3D). The same supernatants were subsequently used to stimulate OKF6 cells. As seen in Figure 3.3E, CXCL14 gene expression was strongly upregulated when stimulated with the wildtype P. gingivalis cell- free culture supernatants. In comparison, the P. gingivalis KDP136 cell-free culture supernatants stimulated a significantly weaker CXCL14 response (Fig 3.3E). In summary,

these data suggest that cell-surface and OMV-associated gingipain proteases are involved in the stimulation of CXCL14 gene expression in OKF6 cells by P. gingivalis.

A B

C D E

Figure 3.3 P. gingivalis gingipain proteases stimulate CXCL14 expression. (A) OKF6 cells were challenged with wildtype P. gingivalis or P. gingivalis KDP136 at 100 MOI for 24 h. CXCL14 mRNA levels were measured by Real-Time PCR and shown as a fold change relative to mock-challenged cells (n=3). (B) OKF6 cells were challenged with P. gingivalis that had been pre-treated with TLCK, and CXCL14 mRNA levels were measured by Real-Time PCR and shown as a fold change relative to mock-challenged cells (n=3). (C-D) Proteolytic activity of (C) Kgp and (D) RgpA/B in cell-free culture supernatant from wildtype P. gingivalis and P. gingivalis KDP136 were measured (n=3). (E) OKF6 cells were stimulated with brain heart infusion broth (BHI) or cell-free culture supernatant from wildtype P. gingivalis and P. gingivalis KDP136 for 24 h. CXCL14 mRNA levels were measured by Real-Time PCR and shown as a fold change relative to BHI-stimulated cells (n=3). All data are presented as the mean ± SEM (** = p < 0.01, *** = p < 0.001).

PAR-3-dependent regulation of P. gingivalis-stimulated CXCL14 gene expression PARs can mediate the recognition of host as well as microbial proteases (Soh et al., 2010). P. gingivalis gingipain proteases have been shown to stimulate cytokine responses in KB cells and human keratinocytes via PAR-1 and PAR-2 (Dommisch et al., 2007; Giacaman et al., 2009; Lourbakos et al., 2001). Given the involvement of the gingipain proteases in the stimulation of CXCL14 gene expression by P. gingivalis, a role for the PARs in regulating CXCL14 response was investigated. The basal gene expression levels of PARs in OKF6 cells were first measured. PAR-2 was found to be most highly expressed, whilst PAR-1 and PAR-3 were expressed at lower levels, and PAR-4 mRNA was not detected (Fig. 3.4A). The effects of P. gingivalis on the expression levels of the PARs were also investigated. P. gingivalis was found to weakly induce the expression of PAR-2 (Fig. 3.4B), whilst PAR-1 and PAR-3 expression remained unchanged (Fig. 3.4C-D).

A B

C D

Figure 3.4 P. gingivalis stimulates PAR-2 gene expression in oral epithelial cells. (A) Basal PAR-1, PAR-2, PAR-3 and PAR-4 mRNA levels in OKF6 cells were measured by Real-Time PCR and are shown as relative to TBP (endogenous control gene) (n=3). (B-D) OKF6 cells were challenged with P. gingivalis at 100 MOI for 24 h. (B) PAR-2, (C) PAR-1, and (D) PAR-3 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-challenged cells (n=3). All data are presented as the mean ± SEM (ND = not detected, *= p < 0.05). 75

The involvement of PAR-1, PAR-2, and PAR-3 in regulating P. gingivalis-inducible CXCL14 expression was subsequently investigated by siRNA-mediated gene silencing. The knockdown of PAR-1, PAR-2, and PAR-3 gene expression in OKF6 cells was confirmed by Real-Time PCR (Fig. 3.5A, C and E). As shown in Figure 3.5B, knockdown of PAR-1 did not affect the stimulation of CXCL14 gene expression by P. gingivalis. Likewise, P. gingivalis-inducible CXCL14 gene expression was not affected by PAR-2 knockdown (Fig. 3.5D). In contrast, P. gingivalis-inducible CXCL14 gene expression was significantly inhibited by knockdown of PAR-3 (Fig. 3.5F). Taken together, these data indicate that P. gingivalis stimulates CXCL14 expression in OKF6 cells in a PAR-3-dependent manner.

A B

C D

E F

Figure 3.5 PAR-3-dependent regulation of P. gingivalis-inducible CXCL14 expression. OKF6 cells were transfected with a non-targeting (-) siRNA, or a (A-B) PAR-1, (C -D) PAR-2, or (E-F) PAR-3 (+) siRNA for 48 h, and subsequently challenged with P. gingivalis at 100 MOI for 24 h. (A) PAR-1, (C) PAR-2, and (E) PAR-3 mRNA levels were measured by Real-Time PCR, and levels in cells transfected with the non-targeting siRNA control were arbitrarily assigned a value of 100%. (B, D, and F) CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-challenged cells (n=3). All data are presented as the mean ± SEM (ns = non-significant, * = p < 0.05, *** = p < 0.001).

EGFR signalling negatively regulates CXCL14 transcription in a MEK-dependent manner Although chemokines are essential for host defence, their expression must be tightly regulated to prevent the development of pathological immune responses. EGFR signalling is associated with epithelial proliferation and differentiation required for maintaining tissue homeostasis. Studies have also demonstrated a role for EGFR in regulating skin inflammation by modulating chemokine expression (Lichtenberger et al., 2013; Mascia et al., 2003; Pastore et al., 2008). EGFR signalling was recently reported to inhibit CXCL14 gene expression in epidermal keratinocytes (Lichtenberger et al., 2013). Therefore, the ability of EGF to regulate CXCL14 gene expression in OKF6 cells was investigated. EGF stimulation was found to strongly suppress CXCL14 gene expression (Fig. 3.6A). Time-course experiments also demonstrated that CXCL14 was rapidly downregulated upon EGF stimulation (Fig. 3.6B). The protein levels of CXCL14 in cell culture supernatants of OKF6 cells were also measured. Consistent with the reduction of CXCL14 mRNA levels in response to EGF stimulation, CXCL14 protein levels were also reduced, although the reduction was not statistically significant (Fig 3.6C). EGF has also been shown to modulate the expression of other chemokines (e.g. CXCL1 and CCL5) (Mascia et al., 2003; Pastore et al., 2005). Therefore, the effects of EGF on the expression of other cytokines/chemokines in OKF6 cells were also investigated. EGF strongly stimulated the expression of the neutrophil chemokine, CXCL1 (Fig. 3.6D). In contrast, EGF suppressed TNF gene expression (Fig. 3.6E). IL-8 and CCL20 gene expression was unaffected by EGF stimulation (Fig. 3.6F-G). Thus, EGF can differentially regulate the expression of specific cytokines and chemokines in OKF6 cells.

A B C

D E

F G

Figure 3.6 EGF differentially regulates cytokine expression. (A) OKF6 cells were stimulated with the indicated concentration of EGF for 24 h. CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-stimulated cells (n=3). (B) OKF6 cells were stimulated with EGF (5 ng/ml) for the times indicated. CXCL14 mRNA levels were then measured by Real-Time PCR, and are shown as a fold change relative to mock-stimulated cells (n=3). (C) OKF6 cells were stimulated with EGF (20 ng/ml) for 24 h, and CXCL14 protein levels in the cell culture supernatants were measured by ELISA. (n=3). (D-G) OKF6 cells were stimulated with the indicated concentration of EGF for 24 h. (D) CXCL1, (E) TNF, (F) IL-8, and (G) CCL20 mRNA levels were measured by Real-Time PCR, and shown as a fold change relative to mock-stimulated cells (n=3). All data are presented as the mean ± SEM (* = < 0.05, ** = p < 0.01, *** = p < 0.001).

Ligand-induced activation of EGFR induces a series of signalling cascades mediated by mitogen-activated protein kinases (MAPKs), including ERK1/2 and p38 MAPK (Avraham and Yarden, 2011). The ability of EGF to stimulate ERK1/2 and p38 MAPK activation in OKF6 cells were first investigated. Specifically, phospho-specific antibodies were used to assess EGF-mediated activation of ERK1/2 and p38 MAPK. As shown in Figure 3.7A-B, ERK1/2 and p38 MAPK were rapidly activated in response to EGF stimulation. Subsequently, the involvement of ERK1/2 and p38 MAPK in the regulation of CXCL14 by EGFR signalling was investigated with pharmacological inhibitors. The blockade of ERK1/2 signalling with the MEK inhibitor, U0126 (Duncia et al., 1998), prevented the suppression of CXCL14 expression by EGF (Fig. 3.7C). By contrast, the p38 MAPK inhibitor, SB203580 (Cuenda et al., 1995), did not prevent the suppression of CXCL14 (Fig. 3.7C). This suggested that EGF suppresses CXCL14 expression by activating the MEK-ERK1/2 pathway. The EGF-MEK-ERK1/2-dependent downregulation of CXCL14 expression could potentially be attributable to transcriptional repression or decreased mRNA transcript stability. The mode of regulation was investigated by inhibiting gene transcription with the RNA polymerase inhibitor, Actinomycin D. Blocking transcription inhibited the upregulation of CXCL14 caused by the MEK inhibitor (Fig. 3.7D). This suggests that the suppression of CXCL14 gene expression by EGF-induced MEK-ERK1/2 signalling is a transcription-dependent event. The rate of CXCL14 mRNA decay in the presence and absence of EGF was also investigated. CXCL14 mRNA stability was found to be unaffected by EGF-MEK-ERK1/2 signalling (Fig. 3.7E). Taken together, these data suggest that EGF signals through the MEK-ERK pathway to inhibit the transcription of CXCL14.

A B EGF (min) 0 5 15 30 EGF (min) 0 5 15 30 40 40 p-ERK1/2 p-p38

40 ERK2 p38 40

C D E

Figure 3.7 EGF suppresses CXCL14 transcription via MEK signalling. (A-B) OKF6 cells were stimulated with EGF (20 ng/ml) for the indicated times. The cell lysates were then subjected to Western blotting with (A) anti-phospho-ERK1/2 and anti-ERK2 antibodies, and (B) anti-phospho-p38 MAP kinase and anti-p38 MAP kinase antibodies. The positions of molecular mass standards (in kDa) are as indicated. The data are representative of two independent experiments. (C) OKF6 cells were stimulated with EGF (20 ng/ml) for 24 h and treated with the MEK inhibitor U0126 (10 µM) or the p38 inhibitor SB203580 (5 µM) for 8 h. CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to cells stimulated with EGF (n=3). (D) OKF6 cells were stimulated with EGF (20 ng/ml) for 24 h and treated with the MEK inhibitor U0126 (10 µM) and Actinomycin D (1 µg/ml) for 8 h. CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to cells stimulated with EGF (n=3). (E) OKF6 cells were treated with Actinomycin D (1 µg/ml) for 1 h, and stimulated with EGF (20 ng/ml) for the indicated times. CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-stimulated cells (n=3). All data are presented as the mean ± SEM (ns = non-significant, ** = p < 0.01).

P. gingivalis gingipain proteases antagonise the negative regulation of CXCL14 gene expression by EGF P. gingivalis is known to antagonise immune signalling networks as a means of subverting host defence (Hajishengallis, 2009, 2014), and has been shown to dampen EGFR signalling in epidermal fibroblasts by inactivating EGF (Pyrc et al., 2013). Thus, there was a possibility that EGF may modulate the stimulation of CXCL14 gene expression by P. gingivalis. To investigate this possibility, OKF6 cells were cultured in the presence or absence of EGF, and subsequently challenged with P. gingivalis. EGF was found to significantly reduce, by around 50%, the stimulation of CXCL14 gene expression by P. gingivalis (Fig. 3.8A). As expected, the P. gingivalis gingipain-deficient mutant (P. gingivalis KDP136) did not stimulate CXCL14 gene expression, and nor did it relieve EGF-mediated suppression of CXCL14 transcription (Fig. 3.8A). For comparison, the effect of EGF on the stimulation of CXCL1 gene expression by P. gingivalis was also examined. Although P. gingivalis and EGF both stimulated CXCL1 gene expression, they did not exert an additive effect (Fig. 3.8B). Notably, the P. gingivalis gingipain-deficient mutant and EGF appeared to exert additive effects on the stimulation of CXCL1 expression (Fig. 3.8B). Taken together, these data suggest that P. gingivalis can antagonise the EGF-mediated negative and positive regulation of CXCL14 and CXCL1 expression, respectively.

The gingipain proteases expressed by P. gingivalis have been demonstrated to be capable of degrading host immunomodulatory factors (e.g. C3 complement factor and IL-8) (Zhang et al., 1999; Stathopoulou et al. 2009; Potempa et al., 2009). Therefore, the ability of the gingipain proteases to degrade EGF was tested. Specifically, EGF was incubated with P. gingivalis for up to 3 h, and proteolysis was then assessed by SDS-PAGE. For comparison, the ratio of P. gingivalis to EGF in Figure 3.8A-B was 2×106 P. gingivalis per ng EGF. The incubation of 500 ng EGF with 1×106 P. gingivalis (i.e. 2×103 P. gingivalis per ng EGF) resulted in significant EGF degradation within 60 min (Fig. 3.8C). Complete EGF degradation was observed within 30 min when incubated with 1×108 P. gingivalis (Fig. 3.8C). Conversely, EGF was not degraded when incubated with 1×106 P. gingivalis KDP136 (Fig. 3.8C); significant degradation was only apparent when EGF was incubated with 1×108 P. gingivalis KDP136 for 3 h (Fig. 3.8C), which was most likely attributable to the activity of other P. gingivalis proteases (e.g. PrtT and Tpr) (Otogoto and Kuramitsu, 1993; Park and McBride, 1993). The ability of purified Kgp to degrade EGF was also investigated. Consistently, EGF was rapidly degraded by Kgp (Fig. 3.8D). Collectively, these data suggest that P. gingivalis gingipain protease-mediated degradation of EGF may dysregulate host expression of CXCL14.

6 A C 1x10 P. gingivalis P. gingivalis KDP136

Time (h) 0 0.5 1 2 3 0 0.5 1 2 3

EGF 3.5 7 1x10 P. gingivalis P. gingivalis KDP136 Time (h) 0 0.5 1 2 3 0 0.5 1 2 3

EGF 3.5 8 1x10 P. gingivalis P. gingivalis KDP136 B Time (h) 0 0.5 1 2 3 0 0.5 1 2 3

EGF 3.5

D Kgp Time (h) 0 0.5 1 2 3 10 EGF 3.5 Kgp 50

Figure 3.8 P. gingivalis antagonises the regulation of CXCL14 by EGF. (A-B) OKF6 cells were stimulated with EGF (20 ng/ml) for 24 h, and then challenged with P. gingivalis or P. gingivalis KDP136 at 100 MOI for 24 h. (A) CXCL14 and (B) CXCL1 mRNA levels then measured by Real-Time PCR, and are shown as a fold change relative to mock-challenged and unstimulated cells (n=3). (C) EGF (500 ng) was incubated with 1×106, 1×107, or 1×108 P. gingivalis or P. gingivalis KDP136 for the indicated times, and then subjected to SDS- PAGE and silver staining. The positions of molecular mass standards (in kDa) are as indicated. The data are representative of three independent experiments. (D) EGF was incubated with purified Kgp for the indicated times, and aliquots of the incubation mixtures were then subjected to SDS-PAGE and silver staining. The positions of molecular mass standards (in kDa) are as indicated. The data are representative of two independent experiments. All data are presented as the mean ± SEM (** = p < 0.01, * = p <0.05).

3.3 Discussion Chemokines are essential immunomodulatory regulators of host defence. For instance, they are important for directing the migration of both innate and adaptive immune cells to sites of infection. In chronic periodontitis, P. gingivalis antagonises and dysregulates the host immune response to create a favourable niche to sustain its growth. Oral epithelial cells lining the surface of the oral mucosa express PRRs, which are essential for recognising and mounting an immune response against oral pathogens. TLR2 is a critical mediator of the inflammatory response to P. gingivalis, whilst other PRRs, such as PAR-1 and PAR-2, can also stimulate inflammation when proteolytically activated by P. gingivalis gingipain proteases (Burns et al., 2010; Dommisch et al., 2007; Giacaman et al., 2009). However, P. gingivalis can hijack and manipulate the host immune response to cause its dysregulation (Hajishengallis, 2014; Hajishengallis et al., 2009). In this Chapter, the expression of the orphan chemokine, CXCL14, in human oral epithelial cells (e.g. OKF6 cells) was identified to be strongly upregulated in response to P. gingivalis. The regulation (and function) of the orphan chemokine CXCL14 has yet to be fully elucidated, therefore the regulation of CXCL14 in oral epithelial cells was investigated.

P. gingivalis gingipain proteases (e.g. Kgp and RgpA/B) are critical virulence factors because of their ability to antagonise and dysregulate host immunity (Imamura, 2003). The results presented in this Chapter showed that the gingipain proteases play a central role in the P. gingivalis-inducible stimulation of CXCL14 expression in OKF6 cells. P. gingivalis has also been demonstrated to induce CXCL14 expression in primary gingival epithelial cells (Chung and Dale, 2008), therefore the stimulation of CXCL14 by P. gingivalis is not limited to OKF6 cells. Epithelial cells typically express cytokines relatively rapidly in response to bacterial challenge, and thus the somewhat delayed induction of CXCL14 expression when OKF6 cells were challenged with P. gingivalis suggests that additional regulatory mechanisms, such as autocrine factors, may be required for the optimal stimulation of CXCL14 expression. Further studies will therefore be required to determine whether other factors induced by P. gingivalis might also contribute, in an autocrine manner, to the regulation of CXCL14 expression.

CXCL14 protein levels in the cell culture supernatant were not significantly increased by P. gingivalis. This was not entirely surprising, because other studies have encountered difficulty in measuring cytokines and chemokines in cell culture supernatants due to their proteolysis by P. gingivalis proteases (Darveau et al. 1998; Zhang et al. 1999; Stathopoulou et al. 2009). In addition to degrading host cytokines and chemokines, the gingipain proteases are also required for the efficient processing of P. gingivalis fimbriae (Nakayama et al., 1996; Kadowaki et al., 1998). Fimbriae not only facilitate the adherence and invasion

84 of oral epithelial cells by P. gingivalis (Weinberg et al., 1997; Yilmaz et al. 2002), they also stimulate the expression of pro-inflammatory genes (Hajishengallis et al., 2009). However, the fact that treating wildtype P. gingivalis with the protease inhibitor TLCK also prevented the stimulation of CXCL14 largely excludes the possibility that impaired fimbriae processing at the cell surface might have been responsible for the lack of CXCL14 stimulation by the isogenic P. gingivalis gingipain protease-deficient mutant, P. gingivalis KDP136. Nonetheless, additional studies will be needed to definitively exclude a role for fimbriae in the stimulation of CXCL14 expression by P. gingivalis, including by mediating epithelial cell adherence and invasion.

In addition to being attached to the outer membrane of P. gingivalis, the gingipain proteases are also associated with OMVs, which are released into the surrounding environment (O’ Brien-Simpson et al., 2003; Potempa et al., 1995). P. gingivalis cell-free culture supernatants containing gingipain proteases have been shown to stimulate chemokine expression (e.g. CCL20) in oral epithelial cells (Dommisch et al., 2007). CXCL14 gene expression in OKF6 cells was similarly upregulated when treated with cell-free culture supernatants derived from wildtype P. gingivalis. Cell-free culture supernatants derived from the P. gingivalis gingipain protease-deficient mutant also weakly induced CXCL14 expression. These findings suggest that, although the stimulation of CXCL14 gene expression in oral epithelial cells in response to P. gingivalis is predominantly mediated by the gingipain proteases, there are also other factors present in the cell-free culture supernatants that can also stimulate CXCL14 gene expression. Further studies will be required to identify and determine the roles of other P. gingivalis-secreted factors in stimulating CXCL14 gene expression in oral epithelial cells.

P. gingivalis-stimulated CXCL14 gene expression was found to be regulated in a PAR-3-dependent manner. Previous studies using P. gingivalis cell-free culture supernatants and purified gingipain proteases have demonstrated roles for PAR-1 and PAR-2 in the induction of inflammatory gene expression (e.g. IL-6 and CCL20) in gingival epithelial cells (Chung et al., 2004; Dommisch et al., 2007; Lourbakos et al., 2001). This is the first study to provide evidence for PAR-3 in regulating chemokine expression in oral epithelial cells in response to P. gingivalis. The function of PAR-3 is poorly characterised. PAR-3 has been shown to act as a co-factor, by dimerising and potentiating PAR-1 activation in endothelial cells (Mclaughlin et al., 2007). Interestingly, the tethered ligand produced from canonical PAR-3 cleavage at Lys38 by thrombin has been shown to also directly activate PAR-1 and PAR-2 (Hansen et al., 2004). Although there is still little evidence to suggest that PAR-3 can signal autonomously, it has recently been reported that thrombin cleavage of endogenous PAR-3 regulated IL-8 gene expression in human

85 lung epithelial cells and astrocytoma cells (Ostrowska and Reiser, 2008). PAR-3 has also been shown to be activated, by non-canonical cleavage at Arg41, in endothelial cells by the coagulation enzyme, activated protease C, to promote endothelial barrier protection (Burnier and Mosnier, 2013). The consequence of differential cleavage of canonical Lys38 and non-canonical Arg41 cleavage in inflammation has yet to be explored, and thus would be an avenue for further investigation. Given that PAR-3 can potentially be activated by both host- and pathogen-produced proteases, the complexity of PAR-3 activation and signalling may therefore depend on the milieu of proteases present. Additionally, PRRs have been shown to act in synergy to regulate cytokine and chemokine expression. For example, the stimulation of THP-1 monocytes with different combinations of PRR agonists, including PAR activating peptides, TLR agonists (e.g. FSL-1 and PolyI:C), NOD agonists (e.g. FK156 and MDP), resulted in additive IL-8 response (Uehara et al., 2008). Therefore, the possibility of PARs cooperating with alternative PRRs may also add to the complexity of PAR signalling.

EGF is an important regulator of epithelial cell proliferation and differentiation, and hence a critical mediator of epithelial tissue homeostasis (Parkar et al., 2001). EGF has been reported to act as an important inflammatory mediator, by regulating cytokine and chemokine expression (Lichtenberger et al., 2013; Mascia et al., 2003; Pastore et al., 2008). In this study, EGF was found to differentially regulate cytokine/chemokine responses differentially in oral epithelial cells (e.g. OKF6 cells). Significantly, EGF strongly inhibited CXCL14 transcription in a MEK-ERK1/2-dependent manner. However, given that Actinomycin D acts as a global transcriptional inhibitor, further studies will be required to determine whether other signalling modulators (e.g. protease or kinase/phosphatase) might also contribute to the regulation of CXCL14 expression by EGF. TNF gene expression was similarly suppressed, whilst CXCL1 gene expression was upregulated with EGF stimulation. Consistently, EGF has also been reported to regulate CXCL14, TNF, and CXCL1 in a similar manner in epidermal keratinocytes (Lichtenberger et al., 2013; Mascia et al., 2003). Although EGF is constitutively present in saliva, gingival tissues, and gingival crevicular fluid, the role of EGF in periodontitis is unclear (Chang et al., 1996; Mogi et al., 1999). EGF has been shown to be important for the proliferation of human periodontal ligament fibroblasts and human gingival fibroblasts (Matsuda et al., 1992). Additionally, during periodontitis suggest that EGFR signalling is likely important for the wound healing response in periodontitis (Chang et al., 1996). Critically, the loss of EGFR signalling in the epidermis has been demonstrated to increase cytokine and chemokine levels and immune cell infiltration (e.g. macrophages and neutrophils), resulting in skin inflammation (Lichtenberger et al., 2013; Pastore et al., 2005). Thus, EGFR signalling in the oral

86 epithelium may also be important for coordinating the recruitment of immune cells during the transition between the inflammatory and the wound healing response.

The findings presented in this Chapter revealed that P. gingivalis can negate EGF-mediated regulation of CXCL14 (and CXCL1) through gingipain-mediated degradation of EGF. Interestingly, in addition to EGF, other growth factors, including keratinocyte growth factor (KGF) and insulin-like growth factor 1 (IGF-1), which likewise activate MEK-ERK1/2, are also upregulated in the oral mucosa during inflammation and wound healing (Li et al., 2005; Werner and Katz, 2004). Therefore, P. gingivalis gingipain protease-mediated degradation of various host factors that can activate the MEK-ERK1/2 pathway may potentially contribute to the optimal stimulation of CXCL14 expression. P. gingivalis can also antagonise EGF signalling through other mechanisms. For instance, P. gingivalis expresses a peptidylarginine deiminase (PAD) that inhibits EGF-induced EGFR signalling by catalysing the citrullination of the C-terminal arginine residue in EGF (Pyrc et al., 2013). Furthermore, P. gingivalis LPS has been shown to attenuate EGFR-regulated signalling, including ERK1/2 activation (Elkaim et al., 2017; Quinchia-Rios et al., 2008). Accordingly, P. gingivalis gingipain proteases likely stimulate CXCL14 through PAR-3-dependent signalling, and by antagonising signalling growth factors (e.g. EGF) that activate the MEK-ERK1/2 pathway.

This Chapter has defined a regulatory pathway for P. gingivalis-stimulated CXCL14 gene expression in oral epithelial cells. Specifically, the upregulation of CXCL14 gene expression by P. gingivalis in OKF6 cells was shown to be dependent on the host protease-activated receptor, PAR-3. Concomitantly, P. gingivalis can potentiate CXCL14 gene expression by degrading EGF to relieve its transcriptional suppression of CXCL14 (Fig. 3.9). Given that P. gingivalis dysregulates the host immune response to promote oral dysbiosis and inflammation, host immunomodulatory factors can participate in the progression of chronic periodontitis. Therefore, further studies will be required to determine whether CXCL14 has a host-protective or destructive role in chronic periodontitis.

P. gingivalis Kgp/Rgp

EGF PAR-3

MEK-ERK1/2

CXCL14

Figure 3.9 A proposed model for the regulation of CXCL14 gene expression in oral epithelial cells by P. gingivalis. The outer membrane-associated gingipain proteinases of P. gingivalis (Kgp/Rgp) stimulate the expression of CXCL14 in a PAR3-dependent manner. Maximal CXCL14 expression requires the gingipain-mediated degradation of EGF, which relieves the transcriptional repression of the CXCL14 gene by the ERK1/2 pathway.

The work presented in this Chapter has given rise to the following publication:

Function of CXCL14

4.1 Introduction Chemokines have pleiotropic immunomodulatory functions in the host immune response to infection. For example, they play important roles in the regulation of immune surveillance and inflammation by directing the migration of immune cells to sites of tissue injury and/or infection. CXCL14, which is an orphan member of the CXC family of chemokines, was initially identified in breast and kidney tissues, and was later found to also be expressed in other epithelia, including the oral epithelium (Hromas et al., 1999; Meuter and Moser, 2008). The conservation of the amino acid sequence of CXCL14 between vertebrates suggests that it likely possesses indispensable function(s). However, our understanding of the physiological function of CXCL14 is still relatively limited due to its orphan state.

CXCL14 has been reported to possess chemotactic activity towards several immune cell populations. For example, in vitro studies have demonstrated the ability of CXCL14 to induce the migration of natural killer cells, dendritic cells, monocytes and neutrophils (Cao et al., 2000; Kurth et al., 2001; Shellenberger et al., 2004; Shurin et al., 2005; Starnes et al., 2006). However, the normal immune phenotype of CXCL14-deficient mice suggests that compensatory mechanisms for CXCL14-regulated chemotaxis likely exist (Meuter et al., 2007). In addition to chemotactic activity for immune cells, CXCL14 has also been reported to regulate angiogenesis by inhibiting the migration of endothelial cells (Shellenberger et al., 2004). Thus, further studies will be required to define the chemotactic activity of CXCL14.

In addition to regulating immune cell chemotaxis, a number of chemokines, including CCL20 and CCL28, have been shown to possess direct antibacterial activity (Hieshima et al., 2003; Yang et al., 2003). The antibacterial activity of the chemokines is likely to be attributed to the formation of a positively-charged surface patch, similar to antimicrobial peptides (e.g. -defensins). CXCL14 has recently been demonstrated to exhibit bactericidal activity; for instance, CXCL14 was shown to kill Streptococcus pyogenes and Pseudomonas pneumoniae by causing cell membrane depolarisation (Dai et al., 2015; Frick et al., 2011). Notably, the bactericidal activity has been mapped to a peptide encompassing Ser1-Arg13 at the N-terminus of CXCL14 (Dai et al., 2015). Importantly, it has been proposed that the constitutive expression of CXCL14 in the epidermis may be an immediate defence mechanism to provide protection against bacterial pathogens upon cutaneous injury (Maerki et al., 2009).

The results presented in Chapter 3 suggest that P. gingivalis may dysregulate CXCL14 gene expression in oral epithelial cells by activating PAR-3-dependent signalling whilst

91 impairing the EGF-mediated transcriptional repression of CXCL14. The function of CXCL14 in the inflammatory response to infection is still unclear. Therefore, this Chapter will examine the potential regulatory role of CXCL14 in inflammation as well as its ability to exert bactericidal activity against oral bacteria.

4.2 Results Effects of CXCL14 on inflammatory gene expression in macrophages CXCL14 is constitutively expressed in the oral epithelium (Hromas et al., 1999; Meuter and Moser, 2008), and the results presented in Chapter 3 demonstrate that its expression by oral epithelial cells is further upregulated in response to P. gingivalis. As briefly described above, CXCL14 has been shown to regulate the migration of immune cells, including monocytes (Kurth et al., 2001). Therefore, the ability of oral epithelial cell-derived CXCL14 to potentially act in a paracrine manner to regulate inflammatory gene expression in macrophages was investigated. Specifically, the mouse macrophage-like cell line, RAW 264.7, was stimulated over a 24 h time-course with recombinant murine CXCL14, and changes in the mRNA expression levels of selected cytokines and chemokines were then measured. Macrophages can produce chemokines that stimulate the recruitment of monocytes to facilitate the clearance of infection (Murray and Wynn, 2012). Therefore, the effect of CXCL14 on CCL2 (aka monocyte chemotactic protein-1 (MCP-1)) gene expression was determined. As shown in Figure 4.1A, CCL2 expression was not modulated by CXCL14 stimulation. In addition to the recruitment of monocytes, macrophages can reinforce the host immune response by producing CCL5 (aka regulated on activation, normal T cell expressed and secreted (RANTES)), which is chemotactic for an array of immune cells, including dendritic cells and T lymphocytes (Marques et al., 2013). However, CXCL14 stimulation did not affect the gene expression of CCL5 (Fig. 4.1B). In addition to chemokines, macrophages also produce pro-inflammatory and anti-inflammatory cytokines to regulate the host inflammatory response (Murray and Wynn, 2012). Therefore, the ability of CXCL14 to regulate the expression of the pro-inflammatory cytokine, TNF, and anti-inflammatory cytokine, IL-10, was also investigated. TNF and IL- 10 gene expression were not affected by CXCL14 stimulation (Fig. 4.1C-D). Collectively, these findings suggest that CXCL14 does not regulate the expression of key inflammatory genes in macrophages (e.g. RAW 264.7 cells). However, given the limited number of genes examined, further studies will be required to establish whether CXCL14 may regulate the inflammatory properties of macrophages.

A B

C D

Figure 4.1 CXCL14 stimulation of mouse macrophage RAW 264.7 cells. RAW 264.7 cells were stimulated with CXCL14 (100 ng/ml) for the indicated times. (A) CCL2, (B) CCL5, (C) TNF, and (D) IL-10 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-stimulated cells (n=3). All data are presented as the mean ± SEM. Effects of CXCL14 on inflammatory gene expression in oral epithelial cells The ability of CXCL14 to act in an autocrine manner to regulate inflammatory gene expression in oral epithelial cells was also investigated. Specifically, OKF6 cells were stimulated over a 24 h time-course with recombinant human CXCL14, and changes in the mRNA expression levels of selected cytokines and chemokines were then measured. Neutrophils are instrumental to the host immune response to infection, and constitute a large proportion of the inflammatory infiltrate in the junctional epithelium in periodontitis (Scott and Krauss, 2013). In addition to mediating direct antimicrobial responses, neutrophils can also act in conjunction with Th17 lymphocytes to sustain the inflammatory response (Hajishengallis, 2014; Pelletier et al., 2010). Accordingly, the ability of CXCL14 to regulate the expression of the neutrophil chemoattractant, IL-8, and Th17 chemoattractant, CCL20, were examined. As shown in Figure 4.2A, IL-8 gene expression was not significantly affected by CXCL14 stimulation. Similarly, CCL20 gene expression was also unaffected by CXCL14 stimulation (Fig. 4.2B). The family of colony stimulating factors: CSF-1 (M-CSF), CSF-2 (GM-CSF) and CSF-3 (G-CSF), are potent 93 haematopoietic growth factors and important for the development and functions of macrophages and neutrophils (Hamilton, 2008; Metcalf, 2008), and as described in Chapter 1, macrophages and neutrophils are important in the pathogenesis of chronic periodontitis. Therefore, the effects of CXCL14 stimulation on these factors were examined. CSF-1 gene expression was unchanged with CXCL14 stimulation (Fig. 4.2C), and whilst CSF-2 expression appeared to be weakly upregulated, the increase was not statistically significant at either the 2 h or 8 h time-point (Fig. 4.2D). Likewise, CSF-3 gene expression was not significantly regulated by CXCL14 (Fig. 4.2E). CXCL14 gene expression was also measured to determine whether CXCL14 may regulate its own expression, potentially as part of a feedback mechanism; however, its expression was unchanged with CXCL14 stimulation (Fig. 4.2F). Taken together, these results suggest that CXCL14 does not regulate the expression of genes involved in the control of immune cells in oral epithelial cells (e.g. OKF6 cells), though this conclusion is based on examination of a limited number of genes.

A B C

D E F

Figure 4.2 CXCL14 stimulation of OKF6 cells. OKF6 cells were stimulated with CXCL14 (100 ng/ml) for the indicated times. (A) IL-8, (B) CCL20, (C) CSF-1, (D) CSF-2, (E) CSF-3, and (F) CXCL14 mRNA levels were measured by Real-Time PCR, and are shown as a fold change relative to mock-stimulated cells (n≥3). All data are presented as the mean ± SEM.

Effects of CXCL14 on inflammatory gene expression in the mouse gingiva In vitro studies are limited in that they cannot entirely reflect what occurs in vivo. Thus, a pilot study was undertaken to investigate the ability of CXCL14 to regulate inflammatory gene expression in vivo. Briefly, recombinant murine CXCL14 (either 0.5 µg or 2 µg) was injected into the gingiva of BALB/c mice. Mice were killed either 24 h or 48 h post- injection, and gingival tissues were harvested for analysis of inflammatory gene expression. The effect of CXCL14 administration on the expression of the neutrophil chemoattractant CXCL1 was first assessed. There was no difference in CXCL1 mRNA levels 24 h or 48 h post-intragingival injection of either 0.5 μg or 2.0 μg CXCL14, when compared to time-matched, PBS-injected control mice (Fig. 4.3A). The effects of CXCL14 on the mRNA expression levels of the pro-inflammatory cytokines, TNF and IL-6, were also investigated. TNF mRNA levels in mice injected with either 0.5 μg or 2.0 μg CXCL14 were comparable to PBS-injected, control mice at 24 or 48 h-post injection (Fig. 4.3B). Similarly, intragingival injection of CXCL14 did not affect IL-6 mRNA levels (Fig. 4.3C). Consistent with previous findings (Meuter and Moser, 2008), CXCL14 mRNA was found to be highly expressed in the gingiva of mice (Fig. 4.3D); however, they were not further modulated in response to intragingival injection of CXCL14 (Fig. 4.3D). Because the mRNA levels for the genes measured were highly variable between mice within the same treatment group, no conclusive results were obtained from this pilot study. In addition, the time-points at which inflammatory gene expression were measured may also have not been optimal.

A B

C D

Figure 4.3 Effects of CXCL14 on inflammatory gene expression in the mouse gingiva. Murine CXCL14 (0.5 µg or 2 µg) was injected into the gingiva of BALB/c mice for 24 and 48 h. (A) CXCL1, (B) TNF, (C) IL-6, and (D) CXCL14 mRNA levels in the gingiva were measured by Real-Time PCR and are shown as relative to HPRT (endogenous control gene) (n=3). All data are presented as the mean ± SEM.

Effect of CXCL14 on oral epithelial cell migration Wound healing is a highly dynamic process, involving reepithelisation and accompanied by angiogenesis. In addition to regulating immune cell chemotaxis, CXCL14 has been reported to antagonise IL-8-mediated endothelial cell migration and thereby inhibit angiogenesis (Shellenberger et al., 2004). CXCL14 gene expression has also been found to be dysregulated in various cancers (e.g. pancreatic and breast cancer) and associated with enhanced cancer cell invasion (Pelicano et al., 2009; Wente et al., 2008). Therefore, a role for CXCL14 in regulating oral epithelial cell migration was investigated with the well- established “scratch wound” assay. The assay is a relatively simple method that allows assessment of cell migration based on the rate of gap closure (Liang et al., 2007). Briefly, OKF6 cells were cultured to confluence, and a “scratch” (wound) was then created in the cell monolayer with a sterile pipette tip. The cells were washed with warm culture medium to remove detached cells, and then cultured in medium (without EGF and BPE) in the presence and absence of added CXCL14 for 6 h. Photographic images were taken at 2 h intervals to enable assessment of gap (wound) closure. The images taken at the 2 h time- point showed the uniform migration of OKF6 cells at the leading edge of the scratch, and by 6 h the wound area was visibly reduced (Fig. 4.4A). The rate of gap closure was quantified by measuring the area of the gap at each time point and expressing the value as a percentage relative to the gap at time = 0 h. Gap closure increased by approximately 10% within 2 h, and 60% after 6 h (Fig. 4.5B). However, CXCL14 did not appear to affect the rate of gap closure (Fig. 4.4B). Therefore, these results suggest that CXCL14 does not regulate oral epithelial cell (e.g. OKF6 cells) migration.

A PBS CXCL14

0 h

2 h

4 h

6 h

Figure 4.4 Effect of CXCL14 on OKF6 cell migration. (A) OKF6 cells were subjected to a scratch-wound assay, and stimulated with PBS or CXCL14 (100 ng/ml). The data presented is representative of three independent experiments. Scale bar: 100 µm. (B) Gap closure was quantified by measuring the area of the gap at each time point, and expressed as % gap closure relative to 0 h. The data is presented as the mean ± SEM.

Bactericidal activity of CXCL14 against oral bacteria In addition to being inflammatory mediators, some chemokines (e.g. CCL20) have also been demonstrated to exert direct bactericidal activity. CXCL14 was recently reported to possess bactericidal activity against bacteria, including Streptococcus pneumoniae, which is associated with respiratory tract infection (Dai et al., 2015; Frick et al., 2011; Yang et al., 2003). Therefore, the bactericidal activity of CXCL14 against oral bacteria was investigated. Because CXCL14 has previously been shown to kill E. coli (Dai et al., 2015; Maerki et al., 2009), the bactericidal activity of recombinant CXCL14 against E. coli was used as a positive control in this study. Consistent with those previous studies, E. coli was found to be highly susceptible to killing by CXCL14. Approximately 50% killing was achieved at the lowest CXCL14 concentration tested (e.g. 0.25 µM), and 100% killing achieved at ≥1 µM CXCL14 (Fig. 4.5A). The bactericidal activity of CXCL14 against oral bacteria was next investigated. Notably, P. gingivalis appeared to be resistant to killing by CXCL14, even when treated with 2 μM CXCL14 (Fig. 4.5B). Contrastingly, Streptococcus gordonii, a Gram-positive bacterium and early coloniser of the oral biofilm, was highly susceptible to killing by CXCL14 (Fig. 4.5C). For example, >95% killing of S. gordonii was achieved at 1 μM CXCL14. Streptococcus sp. OT058, which is frequently associated with periodontal health, was likewise highly susceptible to CXCL14-mediated killing (Fig. 4.5D). These data demonstrate significant differences in the ability of CXCL14 to kill oral bacteria. Moreover, they show that P. gingivalis is resistant to killing by CXCL14.

A B

C D

Figure 4.5 Bactericidal activity of CXCL14. (A) Escherichia coli, (B) P. gingivalis, (C) Streptococcus gordonii, and (D) Streptococcus sp. OT058 were incubated with the indicated concentration of recombinant CXCL14 for 1 h, and killing then measured by CFU assay (n=3). All data are presented as the mean ± SEM (* = p < 0.05, ** = p < 0.01, *** = p < 0.001).

100

Gingipain protease-dependent degradation of CXCL14 by P. gingivalis The mature human CXCL14 protein contains fourteen lysine and seven arginine residues (Fig. 4.6A), thus potentially making CXCL14 susceptible to degradation by the P. gingivalis gingipain proteases, Kgp and RgpA/B. This raised the possibility that P. gingivalis may resist CXCL14-mediated killing by proteolytically degrading CXCL14. Therefore, the ability of the gingipain proteases to degrade CXCL14 was investigated. Briefly, CXCL14 was incubated with P. gingivalis for up to 1 h, and proteolysis of CXCL14 was then assessed by SDS-PAGE. The incubation of 500 ng CXCL14 with 1×104 P. gingivalis, which was equivalent to the ratio of CXCL14 to P. gingivalis in the antimicrobial assays in Figure 4.5, resulted in significant proteolysis of CXCL14 (Fig. 4.6B). CXCL14 was completely degraded when incubated with greater numbers of P. gingivalis (e.g. 1×106 P. gingivalis) (Fig. 4.6B). In contrast, when CXCL14 was incubated with the isogenic P. gingivalis gingipain protease-deficient mutant, P. gingivalis KDP136, CXCL14 degradation was only evident when incubated with 1×108 bacteria (Fig 4.6B). The ability of purified Kgp to degrade CXCL14 was also tested. As shown in Figure 4.6C, CXCL14 was rapidly degraded by Kgp. These findings indicate that the gingipain proteases can directly degrade CXCL14.

A 10 20 30 40 50 60 70 SKCKCSRKGPKIRYSDVKKLEMKPKYPHCEEKMVIITTKSVSRYRGQEHCLHPKLQSTKRFIKWYNAWNEKRRVYEE

B P. gingivalis 4 6 8 1x10 1x10 1x10 C Kgp Time (h) 0 0.5 1 0 0.5 1 0 0.5 1 Time (h) 0 0.5 1 2 3 10 10 CXCL14

CXCL14 P. gingivalis KDP136 3.5 4 6 8 1x10 1x10 1x10 Time (h) 0 0.5 1 0 0.5 1 0 0.5 1 50 10 Kgp CXCL14

Figure 4.6 Gingipain protease-mediated degradation of CXCL14. (A) Amino acid sequence of mature human CXCL14. (B) CXCL14 (500 ng) was incubated with 1×104, 1×106, or 1×108 P. gingivalis or P. gingivalis KDP136 for the time indicated, and then subjected to SDS-PAGE and silver staining. (C) CXCL14 was incubated with purified Kgp for the indicated times, and then subjected to SDS-PAGE and silver staining. The positions of molecular mass standards (in kDa) are as indicated. The data in (B) and (C) are representative of two independent experiments.

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Identification of CXCL14 peptides resulting from Kgp digestion A recent report revealed that the first thirteen amino acids of the mature CXCL14 protein (i.e. Ser1‒Arg13) largely mediate its bactericidal activity, whilst Tyr14-Lys54 was shown to stimulate chemotaxis of THP-1 monocytes (Dai et al., 2015). Thus, aliquots of the digestion reactions shown in Fig. 4.6C were subjected to analysis by mass spectrometry to identify the CXCL14-derived peptides generated by Kgp digestion. Several peptides from the N-terminal half of CXCL14 were identified; however, none contained Ser1-Arg13 or Tyr14-Lys54 (Table 4.1). This was not particularly surprising given that the amino acid sequence spanning Ser1‒Arg13 contains four lysine residues, and Tyr14-Lys54 contains six lysine residues (Fig. 4.6A). Several relatively short peptides from the C-terminal half of CXCL14 were also identified (Table 4.1). Taken together, these data indicate that the gingipain proteases of P. gingivalis can degrade CXCL14, and thereby likely inhibit its bactericidal activity.

Table 4.1 CXCL14 peptides identified by MS from Kgp digestion.

Peptides Identified Ion Score 12 19 IRYSDVKK 10 19 25 KLEMKPK 34 20 32 57 LEMKPKYPHCEEK 26 32 YPHCEEK 17 33 39 MVIITTK 53 64 71 WYNAWNEK 36 64 77 WYNAWNEKRRVYEE 9

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4.3 Discussion The host immune system relies on the tightly coordinated actions of immunomodulatory factors to clear infection by pathogens. Chemokines are instrumental in coordinating the immune response as they regulate the recruitment of both innate and adaptive immune cells to sites of infection. Although the inflammatory response is typically host-protective, P. gingivalis can manipulate and subvert the host immune response for its own benefit. The dysregulation of cytokine and chemokine expression by P. gingivalis can lead to the excessive recruitment of immune cells with suboptimal functions, and thus cause chronic inflammation that mediates host tissue breakdown to increase nutrient availability (e.g. haem). The data presented in Chapter 3 demonstrated that P. gingivalis can potentially dysregulate CXCL14 gene expression by stimulating its expression via PAR-3, while concurrently inhibiting EGF-mediated transcriptional repression of CXCL14. This Chapter explored some of the potential immunomodulatory functions of CXCL14. A gingipain protease-mediated mechanism that may enable P. gingivalis to further dysregulate CXCL14 function was also identified.

The amino acid sequence of CXCL14 is highly conserved in vertebrates, and its constitutive expression in various epithelia suggests that CXCL14 may have important inflammatory and homeostatic functions. The participation of CXCL14 in modulating the inflammatory response was examined in this study. It was found that CXCL14 did not regulate the expression of the inflammatory genes examined in either human oral epithelial cells (e.g. OKF6 cells) or mouse macrophages (e.g. RAW 264.7 cells). However, the potential to identify CXCL14-regulated inflammatory genes was limited, as only a small number of genes were investigated in this study. A more comprehensive, and unbiased approach, such as RNA-seq, would likely provide greater insight into the potential role of CXCL14 as a regulator of inflammatory genes in oral epithelial cells and macrophages. However, given that the receptor for CXCL14 is still unknown, there is also the possibility that neither of these cell types express the receptor for CXCL14.

Recent studies suggest that CXCL14 can interact with CXCR4, the receptor for CXCL12 (aka stromal cell-derived factor-1 (SDF-1)) (Collins et al., 2017; Hara and Tanegashima, 2014; Tanegashima et al., 2013). However, CXCL14 does not stimulate CXCR4 receptor signalling, instead it functions as an allosteric modulator to prime and sensitise CXCL12-mediated CXCR4 signalling. This regulatory function for CXCL14 was demonstrated in natural killer cells, T lymphocytes and B lymphocytes (Collins et al. 2017). Interestingly, CXCR4 has been shown to participate in signalling crosstalk with TLR2 and TLR4 to dampen the host inflammatory response (Hajishengallis et al., 2008; Kishore et al., 2005). For example, P. gingivalis fimbriae can stimulate the co-association of CXCR4 and TLR2 in lipid rafts in

103 macrophages to promote signalling crosstalk. Importantly, this inhibits TLR2-mediated NF-B activation and nitric oxide production, which is required for the intracellular killing of P. gingivalis (Hajishengallis et al., 2008). In a similar vein, CXCL12-mediated activation of CXCR4 signalling has been shown to inhibit TLR4 activation at low LPS concentrations, potentially preventing inappropriate inflammatory responses to the normal microbiota (Kishore et al., 2005). Accordingly, the stimulation of CXCL14 expression by P. gingivalis might potentiate the aforementioned signalling crosstalk by enhancing the potency of CXCR4 ligands. It is therefore tempting to speculate that the dysregulation of CXCL14 expression by P. gingivalis might potentiate CXCR4 crosstalk and contribute to microbial dysbiosis by not only suppressing macrophage antimicrobial activity, but also increasing the activation threshold of TLR4 and thereby alter the host inflammatory response to the oral biofilm. Thus, the contribution of CXCL14 to CXCR4/TLR signalling crosstalk warrants further investigation.

A mouse intragingival injection model was established to enable in vivo investigation of CXCL14 function. Consistent with a previous study (Meuter and Moser, 2008), CXCL14 mRNA was found to be highly expressed in the gingival tissues of mice. However, due to the variability in gene expression between mice, it is unclear whether CXCL14 may potentially play a role in regulating inflammatory gene expression in the gingiva. Furthermore, the time-points selected may not have been optimal to identify changes in the expression of CXCL14-target genes. The mouse model undertaken here was a pilot study, with the aim of gaining data to justify a follow-up study involving larger numbers of mice. Although in vivo mouse models allow biological studies to be conducted in a more physiologically relevant environment, responses can be highly variable between mice. Relatively large numbers of mice are therefore often required to provide experiments with sufficient statistical power.

Nevertheless, the ability of recombinant CXCL14 to promote leukocyte recruitment when injected into the footpads of BALB/c nude mice (Sleeman et al., 2000) suggests that CXCL14 is capable of regulating inflammation in vivo. In the study by Sleeman et al., CXCL14 induced a similar inflammatory response when injected into C3H/HeJ mice, indicating that the response was specific to CXCL14, rather than a consequence of potential endotoxin (LPS) contamination (Sleeman et al., 2000). Furthermore, CXCL14 has been found to be strongly upregulated in inflamed joints in autoimmune arthritis (Lindberg et al., 2006). The transgenic expression of CXCL14 in mice exacerbates collagen-induced arthritis, and was found to be associated with an increased T lymphocyte response (Chen, Guo, et al., 2012). Moreover, in vitro studies have demonstrated that CXCL14 possesses pleiotropic chemotactic activity towards monocytes, B lymphocytes,

104 neutrophils, immature dendritic cells and natural killer cells (Dai et al., 2015; Shellenberger et al., 2004; Sleeman et al., 2000; Starnes et al., 2006). CXCL14-deficient mice appear to have a normal immune phenotype, although the mice weigh less than their wildtype littermate counterparts (Meuter et al., 2007). Therefore, the possibility of functional redundancy that compensates for the chemotactic activity of CXCL14 cannot be excluded. Intriguingly, CXCL14 knockout mice are born at a lower than expected Mendelian frequency, suggesting a potential role for CXCL14 in embryogenesis (Meuter et al., 2007).

CXC chemokines that lack the ELR (glutamic acid-leucine-arginine) motif (e.g. CXCL9 and CXCL10) have been shown to be potent angiostatic factors (Strieter et al., 1995, 2005). As an ELR-negative chemokine, CXCL14 has been demonstrated to inhibit angiogenesis by interacting with angiogenic ligands (e.g. IL-8 and basic fibroblast growth factor (bFGF)) to inhibit endothelial cell migration (Shellenberger et al., 2004). The ability of CXCL14 to direct oral epithelial cell migration was assessed in this study by the “scratch assay”. The findings obtained suggest that CXCL14 does not regulate the rate of oral epithelial cell migration (e.g. OKF6 cells). However, CXCL14 has been shown to regulate the migration of other cell types, such as osteosarcoma cells and mammary epithelial cells (Lu et al., 2015; Pelicano et al., 2009). The heterogenous expression of CXCL14 in various cancers (e.g. breast and pancreatic cancer) suggests that CXCL14 may be involved in mediating cancer progression (Pelicano et al., 2009; Wente et al., 2008). It is uncertain whether CXCL14 might be acting as a tumour suppressor or oncogene, because CXCL14 gene expression has been reported to be increased in some cancers (e.g. pancreatic and prostate cancer) and decreased in others (e.g. lung cancer) (Augsten et al., 2009; Tessema et al., 2010; Wente et al., 2008). Various studies have provided evidence for the involvement of CXCL14 in promoting cancer metastasis (Augsten et al., 2009; Pelicano et al., 2009), whilst others have shown that the expression of CXCL14 suppressed tumour growth (Ozawa et al., 2006; Tessema et al., 2010). Therefore, CXCL14 tumour-suppressing and tumour-promoting activities are possibly dependent on the tumour type. Nevertheless, the functional characteristics of CXCL14 in cancer studies could potentially provide inferences about the role of CXCL14 in the context of infection, inflammation, and wound healing.

CXCL14 has recently been shown to possess bactericidal activity against some skin and respiratory pathogens, including Pseudomonas aeruginosa and Staphylococcus aureus (Dai et al., 2015). Here, CXCL14 was shown to kill oral Streptococcus species, namely S. gordonii and S. sp. OT058. S. gordonii is a primary coloniser of the tooth-accreted oral biofilm and promotes P. gingivalis colonisation (Park et al., 2005). Accordingly, S. gordonii has been described as being an accessory pathogen in chronic periodontitis (Lamont et al., 1993;

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Lamont and Hajishengallis, 2015). Conversely, the abundance of S. sp. OT058 in the biofilm has been reported to be associated with periodontal health (Hong et al., 2015). CXCL14 has also been demonstrated to kill Streptococcus mitis, one of the most abundant Streptococcus species in the biofilm (Dai et al., 2015). Significantly, P. gingivalis was shown to be resistant to CXCL14-mediated killing. This is potentially attributable to the ability of the gingipain proteases produced by P. gingivalis to proteolytically degrade CXCL14. The results presented in this Chapter suggest that CXCL14 may also be susceptible to degradation by other P. gingivalis proteases (e.g. PrtT and Tpr). In addition to proteolytic degradation of antimicrobial peptides, unique modifications of P. gingivalis LPS have also been proposed to protect P. gingivalis from cationic antimicrobial peptides (e.g. LL-37 and polymyxin B) (Bachrach et al., 2008; Coats et al., 2009). Specifically, phosphorylation of the lipid A moiety of LPS is proposed to contribute resistance to antimicrobial peptides by imparting P. gingivalis with a negative surface-charge. Therefore, the P. gingivalis outer membrane LPS composition may also affect the ability of CXCL14 to kill P. gingivalis.

Human CXCL14 is translated as an 111-amino acid pro-protein, and is cleaved C-terminal of Gly-34 to yield the mature 77-amino acid CXCL14 protein (Cao et al., 2000). The antimicrobial activity of the mature CXCL14 protein has been demonstrated to lie within a short sequence at the N-terminus (i.e. Ser1-Arg13), while Tyr14-Lys54 has been shown to be sufficient for CXCL14 to promote chemotaxis of human monocytes (e.g. THP-1 cells) (Dai et al., 2015). Peptides arising from in vitro Kgp digestion of CXCL14 were analysed by mass spectrometry to determine if they contain Ser1-Arg13 or Tyr14-Lys54. Although this analysis identified several peptides from the N-terminal half of CXCL14, Ser1-Arg13 and Tyr14-Lys54 peptides were not detected. This suggests that the gingipain protease- mediated degradation of CXCL14 likely eliminates CXCL14 bactericidal and chemotactic activity, and hence protects P. gingivalis from CXCL14-mediated host defence. A similar protease-mediated protective mechanism has been proposed to explain the resistance of Finegoldia magna, an opportunistic pathogen, to CXCL14 (Frick et al., 2011). Importantly, the degradation of CXCL14 by P. gingivalis might also potentially protect otherwise susceptible bacteria, including accessory pathogens (e.g. S. gordonii), from killing by CXCL14, and thereby play a role in promoting dysbiosis. In addition, loss of CXCL14- mediated chemotactic activity might also contribute to the proliferation of a dysbiotic biofilm. Additional studies will be required to determine whether the CXCL14 peptides generated from gingipain protease digestion have bactericidal activity against other bacteria, as this could potentially also contribute to the development of dysbiosis.

This study has explored several potential functions of CXCL14 relevant to the oral mucosa. Due to the orphan nature of CXCL14, it is unclear whether CXCL14 can regulate the

106 inflammatory response of oral epithelial cells or macrophages. The findings from this study also suggest that CXCL14 does not regulate the migration of oral epithelial cells. However, this study did demonstrate that CXCL14 can exert differential bactericidal activity against oral bacteria. Significantly, the dysregulation of CXCL14 by P. gingivalis may potentially destabilise the proportions of bacterial species in the biofilm and thereby promote biofilm dysbiosis. Thus, although the expression of CXCL14 by oral epithelial cells is a host-initiated response, its role in host defence might need to be reconsidered in the context of infection by pathogens that rely on dysbiosis to fulfil their nutritional requirements.

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The work presented in this Chapter has given rise to the following publication:

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Identification and characterisation of P. gingivalis TIR domain-containing proteins

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5.1 Introduction The dysregulation of the host immune response by P. gingivalis can induce biofilm dysbiosis and chronic inflammation (Darveau, 2010; Hajishengallis, 2014). Pattern- recognition receptors, such as TLRs, are important for recognising and mounting immune responses against potential pathogens. The TIR domain of TLRs is an essential signalling module in innate immunity. As described in Chapter 1, homotypic interactions between the TIR domain of TLRs provides a platform for the assembly of adaptor proteins, which is necessary for the activation of downstream signal transduction. Importantly, TLR signalling can be hijacked by P. gingivalis as a means of immune subversion (Hajishengallis, 2014). Other bacterial species have been shown to express TIR domain- containing proteins (Tcps) as a means of immune subversion. Newman et al. were the first to show that TlpA from S. enterica was not only capable of suppressing TLR4-mediated NF-B activation in vitro, but was also required for S. enterica virulence in vivo (Newman et al., 2006). This led to the subsequent identification of additional bacterial Tcps (e.g. TcpC from E. coli CFT037 and TcpB from B. melitensis), and the proposal of the “subversion hypothesis”, whereby pathogens adopt molecular mimicry to inhibit the host immune response (Cirl and Miethke, 2010; Radhakrishnan et al., 2009; Sengupta et al., 2010). Although P. gingivalis can interfere with TLR signalling through multiple virulence factors (refer to Section 1.7 for specific details), the existence of P. gingivalis Tcp(s) has not been reported. Therefore, this Chapter will characterise putative P. gingivalis Tcps, using a combination of bioinformatics and biochemical approaches.

5.2 Results Identification of putative P. gingivalis TIR domain-containing proteins A search for TIR domains in the Interpro database was conducted to identify putative P. gingivalis Tcps. The search revealed that there are over 9,000 bacterial proteins annotated with TIR domain signatures, and P. gingivalis is annotated to have twelve Tcps in nine different strains (Table 5.1). A phylogenetic tree was subsequently constructed, using the BLOSUM62 score matrix (Henikoff and Henikoff, 1992), to infer the evolutionary relationships between the Tcps. P. gingivalis W83 and W50 share highly related, if not nearly identical genomes, and thus are denoted as P. gingivalis W83/W50 (Chen et al., 2004). Phylogenetic analysis suggested that the P. gingivalis Tcps can be divided into two groups, which, for simplicity, are denoted as Group A and Group B (Fig. 5.1). The Group A proteins Q7MX37 (PG0382) from P. gingivalis W83/W50 and U2JPW4 (HMPREF1989_02328) from P. gingivalis F0566 differ by only two amino acids (Met169 and Gly231 in Q7MX37, and Arg169 and Glu231 in U2JPW4), and thus are the same

110 protein. The three unique Tcps in Group A are <500 amino acids in length, and share between 13 and 20% overall amino acid sequence identity (Table 5.2). The putative TIR domains of Group A proteins share amino acid sequence identities between 15 to 32% (Table 5.3.). Tcps from Group B comprise proteins >800 amino acids in length, and share >60% overall amino acid sequence identity (Table 5.2). Notably, the putative TIR domains in Tcps within Group B have amino acid sequence identities >70% (Table 5.3). Furthermore, the TIR domains in W1R842 and F5X7L6, and T2N8U1 and A0A0K2J754, are identical (Table 5.3).

Table 5.1 Annotated TIR domain-containing proteins of P. gingivalis strains.

P. gingivalis Accession Gene Protein strain number length ATCC 33277 B2RLS0 PGN1796 1125 W83/W50 Q7MTS7 PG1864 1266 W83/W50 Q7MX37 PG0382 490 AJW4 A0A0K2J754 PGJ00016810 1109 F0566 U2JFL6 HMPREF1989_01737 875 F0566 U2JPW4 HMPREF1989_02328 490 F0566 U2JRV0 HMPREF1989_00095 401 F0570 A0A0E2LT79 HMPREF1555_00114 881 JCVI SC001 T2N8U1 A343_0215 814 JCVI SC001 T2NDV9 A343_1154 440 SJD2 W1R842 SJDPG2_06385 1145 TDC60 F5X7L6 PGTD600128 1384

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Group A

Group B

Figure 5.1 Phylogenetic tree of P. gingivalis TIR domain-containing proteins.

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Table 5.2 Amino acid sequence identity (%) between P. gingivalis TIR domain-containing proteins.

Strain W83/W50 F5066 F0566 SC001 ATCC33277 F0566 SJD2 TDC60 F0570 W83 SC001 SC001

Strain Protein Q7MX37 U2JRV0 U2JPW4 T2NDV9 B2RLS0 U2JFL6 W1R842 F5X7L6 A0A0E2LT79 Q7MTS7 T2N8U1 A0A0K2J754 W83/W50 Q7MX37 - 12.8 99.6 20.1 10.1 12.0 9.4 8.6 12.6 9.8 14.1 9.5 F0566 U2JRV0 12.8 13.0 13.9 5.9 7.3 5.7 4.9 7.9 5.9 7.9 5.9 F0566 U2JPW4 99.6 13.0 - 20.1 10.0 11.9 9.3 8.8 12.5 9.8 14.6 9.4 SC001 T2NDV9 20.1 13.9 20.1 - 2.7 3.3 2.5 2.1 4.0 2.6 3.7 2.9 ATCC33277 B2RLS0 10.1 5.9 10.0 2.7 - 71.5 81.5 62.4 67.9 71.5 79.9 77.9 F0566 U2JFL6 12.0 7.3 11.9 3.3 71.5 - 61.4 51.3 80.5 57.5 79.7 66.2 SJD2 W1R842 9.4 5.7 9.3 2.5 81.5 61.4 - 68.1 67.5 70.9 59.5 76.4 TDC60 F5X7L6 8.6 4.9 8.8 2.1 62.4 51.3 68.1 - 53.3 71.5 52.5 64.4 F0570 A0A0E2LT79 12.6 7.9 12.5 4.0 67.9 80.5 67.5 53.3 - 62.0 84.4 72.2 W83 Q7MTS7 9.8 5.9 9.8 2.6 71.5 57.5 70.9 71.5 62.0 - 59.6 76.1 SC001 T2N8U1 14.1 7.9 14.6 3.7 79.9 79.7 59.5 52.5 84.4 59.6 - 69.3 AJW4 A0A0K2J754 9.5 5.9 9.4 2.9 77.9 66.2 76.4 64.4 72.2 76.1 69.3 -

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Table 5.3 Amino acid sequence identity (%) between P. gingivalis TIR domain-containing proteins TIR domains.

Strain W83/W50 F5066 F0566 SC001 ATCC33277 F0566 SJD2 TDC60 F0570 W83 SC001 SC001

Strain Protein Q7MX37 U2JRV0 U2JPW4 T2NDV9 B2RLS0 U2JFL6 W1R842 F5X7L6 A0A0E2LT79 Q7MTS7 T2N8U1 A0A0K2J754 W83/W50 Q7MX37 - 15.2 100.0 32.1 18.6 18.6 22.7 22.7 17.4 17.4 17.4 17.4 F5066 U2JRV0 15.2 - 15.2 19.5 22.0 22.0 19.5 19.5 19.5 19.5 19.5 19.5 F0566 U2JPW4 100.0 15.2 - 32.1 18.6 18.6 22.7 22.7 17.4 17.4 17.4 17.4 SC001 T2NDV9 32.1 19.5 32.1 - 23.5 23.5 23.7 23.7 24.5 24.5 25.5 25.5 ATCC33277 B2RLS0 18.6 22.0 18.6 23.5 - 98.7 74.7 74.7 98.7 87.5 81.6 81.6 F0566 U2JFL6 18.6 22.0 18.6 23.5 98.7 - 75.8 75.8 88.2 87.5 73.7 82.9 SJD2 W1R842 22.7 19.5 22.7 23.7 74.7 75.8 - 100.0 73.7 73.7 73.7 73.7 TDC60 F5X7L6 22.7 19.5 22.7 23.7 74.7 75.8 100.0 - 73.7 73.7 73.7 73.7 F0570 A0A0E2LT79 17.4 19.5 17.4 24.5 98.7 88.2 73.7 73.7 - 98.0 94.7 94.7 W83 Q7MTS7 17.4 19.5 17.4 24.5 87.5 87.5 73.7 73.7 98.0 - 92.8 92.8 SC001 T2N8U1 17.4 19.5 17.4 25.5 81.6 73.7 73.7 73.7 94.7 92.8 - 100.0 AJW4 A0A0K2J754 17.4 19.5 17.4 25.5 81.6 82.9 73.7 73.7 94.7 92.8 100.0 -

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A NCBI Conserved Domain Search (www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) was performed to establish whether the proteins contain other annotated domains. A nucleoside- triphosphatase (NTPase) domain, which can catalyse the hydrolysis of nucleoside triphosphates to nucleotides (Vetter and Wittinghofer, 1999), was predicted to be present in several Tcps, including U2JPW4/Q7MX37 (PG0382) (Fig. 5.2). A leucine-rich repeat (LRR) domain, which is involved in mediating protein-protein interactions (Kobe and Kajava, 2001), was predicted to be present in all Group B Tcps, except T2N8U1. Interestingly, T2NDV9 was predicted to contain a SIR2 domain, which are involved in transcriptional regulation (North and Verdin, 2004). Therefore, in addition to a TIR domain, putative P. gingivalis Tcps may possess other functional domains. Based on its predicted structural domains, Q7MX37 (referred to as, PG0382, hereafter) from P. gingivalis W83/W50 was selected for further investigation.

Q7MX37/ U2JPW4 Domain: TIR T2NDV9 NTPase U2JRV0 SIR2

LRR

B2RLS0

U2JFL6

W1R842

F5X7L6

A0A0E2LT79

A0A0K2J754

Q7MTS7

T2N8U1

Figure 5.2 Predicted protein domains of annotated P. gingivalis TIR domain-containing proteins.

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PG0382 is predicted to contain a coiled-coil motif Some bacterial Tcps have been identified to contain coiled-coil motifs comprised of heptad sequence repeats, which may facilitate dimer formation and thereby potentially enhance inhibition of TLR signalling (Low et al., 2007; Rana et al., 2011; Fekonja et al. 2012). Therefore, the presence of putative coiled-coil motifs in PG0382 was investigated, using the COILS prediction server (https://embnet.vital-it.ch/software/COILS_form.html) (Lupas et al., 1991). The COILS prediction server compares input sequences with a database of known two-stranded parallel coiled-coils to derive a similarity score, and then generates a probability score that reflects the likelihood of the input protein sequence having a coiled-coil motif (Lupas et al., 1991). Based on the COILS output, PG0382 was predicted to form coiled-coil motifs, in a 14-residue window, between: Gln200-Lys250, Gly350-Glu400, and Thr450-Gln490 (Fig. 5.3). PG0382 is also predicted to form coiled-coil motifs at the same positions when calculated by applying a 21-residue window, albeit with lower probability. Therefore, PG0382 likely contains coiled-coil motifs within the N-terminal half of the protein.

0.8

0.6

0.4

coil forming probability forming coil

- 0.2

Coiled 0

0 50 100 150 200 250 300 350 400 450 500

Residue number

Figure 5.3 COILS analysis output for PG0382

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Comparison of PG0382 with mammalian TLRs and adaptor proteins PG0382 is comprised of 490 amino acids and predicted to contain an NTPase domain between Val8 and Leu52, and a TIR domain between Lys341 and Gln490 (Fig. 5.4). Pairwise sequence alignments were conducted with EMBOSS Needle (www.ebi.ac.uk/Tools/psa/emboss_needle/) to determine the amino acid sequence identity between the PG0382 TIR domain and TLR TIR domains. The accession numbers of the mammalian TLRs are presented in Appendix Table A1. The alignment indicated that the PG0382 TIR domain shares highest sequence identity with TLR1 (17.8%), and lowest identity with TLR4 (3.8%) (Table 5.4). In comparison, TLR TIR domains have sequence identities between 87% (TLR1 and TLR6) and 6% (TLR1 and TLR9). TLR1 and TLR6 are highly homologous, most likely because they heterodimerise with TLR2. However, most TLR TIR domains have sequence identities around 20-30%.

10 20 30 40 50 MDLAEVFVTE GFPHLTYVEP LNYYEILIDV KSKKKPVIIE GQTGTGKTST 60 70 80 90 100 ILKILSDIKE EIHFEYLSAR NIDETTKINQ LINSNFEEGG NFVIDDFHRL 110 120 130 140 150 VDHLKLRLSN IAKLAADNVA NPKYPKLVII GINQTGRELL KLSPDIAKRF 160 170 180 190 200 GVHKIQPATE ENVKSIIEMG EKLLNIKFLR HKPIYSESKG DYWLTQHICQ 210 220 230 240 250 TICTQNGVIN TQEETKQIKL NIKEARRKII GRLEYIYNDI VKEFCRGKRF 260 270 280 290 300

RPSNDAYIKF LESVSKMDDF PIDLNELIGN VDDHHRIAIS SIKGHRLDVL

310 320 330 340 350 IKEKIDLRNN FYYSKDTKLF NIEDPALQYY IKHIDWNKLY KECGFKKNNG 360 370 380 390 400 SYKYDIAISF AGEKRELAEE IADQLQRSDY EVFYDRLYED NYLGMSLSDE 410 420 430 440 450 FERIFTSESK FVVCLLDKNH KKKIWPTFER DCFLEKVQTN EVIPIFLDDT 460 470 480 490 KFPGIPNDIA CIRYEEKQED KKRSERVQRE IIERIISKVQ

Figure 5.4 Amino acid sequence of PG0382. The putative NTPase domain is shown in orange text, and the TIR domain is shown in purple text.

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Table 5.4 Amino acid sequence identity (%) between PG0382 TIR domain and TLR TIR domains.

PG0382 TLR1 TLR2 TLR3 TLR4 TLR5 TLR6 TLR7 TLR8 TLR9

TLR1 17.8 - 51.0 27.7 38.9 25.3 86.9 29.6 33.1 28.9 TLR2 14.0 51.0 - 27.0 40.5 28.2 49.7 38.1 39.1 28.9 TLR3 17.6 27.7 27.0 - 25.2 27.3 27.0 29.9 32.5 28.0 TLR4 3.8 38.9 40.5 25.2 - 28.7 35.1 25.9 28.2 30.9 TLR5 12.6 25.3 28.2 27.3 28.7 - 22.7 24.3 28.3 25.8 TLR6 17.0 86.9 49.7 27.0 35.1 22.7 - 30.5 32.5 27.6 TLR7 14.0 29.6 38.1 29.9 25.9 24.3 30.5 - 58.8 42.4 TLR8 8.9 33.1 39.1 32.5 28.2 28.3 32.5 58.8 - 43.3 TLR9 6.0 28.9 28.9 28.0 30.9 25.8 27.6 42.4 43.3 -

The TIR domain contains conserved sequence motifs in box 1, box 2 and box 3 (Watters et al., 2007). Therefore, a multiple sequence alignment analysis was conducted using ClustalOmega to identify sequence similarities in the conserved TIR domain box 1, box 2 and box 3 sequence motifs of PG0382 and human TLRs (Fig. 5.5). PG0382 contains multiple amino acid residues corresponding to the TIR domain box 1 consensus sequence: F/YDAFISY, including Tyr354, Asp355, Ile358, and Ser359. The sequence alignment also identified differences between PG0382 and TLRs. For example, all TLRs contain the consensus alanine residue, which is replaced by an isoleucine (Ile356) in PG0382. In addition, the phenylalanine residue is replaced by an alanine (Ala357) in PG0382. At the same position, TLRs either contain the conserved phenylalanine or is replaced by tyrosine. Like TLR5 and TLR9, the final consensus tyrosine residue is replaced by phenylalanine (Phe360) in PG0382. The BB loop in box 2 is characterised by an RDxxPG motif, where x denotes a hydrophobic amino acid. Although PG0382 does not contain the consensus arginine residue, it does contain the consensus aspartic acid residue (Asp390) and glycine residue (Gly394). The consensus proline residue, which is critical for TLR4 signalling (Poltorak et al., 1998), and present in all TLRs (except TLR3), is replaced by a leucine (Leu393) in PG0382. Box 3 is defined by an FW motif, and like TLR3 and TLR5, box 3 is absent in PG0382.

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box 1 box 2 F/YDAFISY RDxxPG

box 3 FW

Figure 5.5 Multiple sequence alignment of PG0382 TIR domain with TLR TIR domains. The consensus sequence for box 1, box 2, and box 3 are as indicated. The x in box 2 denotes any hydrophobic amino acid.

Pairwise amino acid sequence alignments were also performed to compare the TIR domain of PG0382 with those of mammalian TLR adaptor proteins (e.g. MYD88, MAL, TRAM, TRIF, and SARM). The accession numbers of the mammalian TLRs are presented in Appendix Table A1. The sequence alignments revealed that the PG0382 TIR domain shares highest sequence identity with MAL (15.2%), and lowest identity with TRIF (9.7%) (Table 5.5). The TIR domains of TLR adaptor proteins have sequence identities between 25.5% (MYD88 and MAL) and 7.8% (MYD88 and TRIF). A multiple sequence alignment was also conducted to compare the TIR domain with adaptor protein TIR domains (Fig. 5.6). PG0382 box 1 shares some conserved amino acid residues with the TLR adaptor proteins. Like MAL and MYD88, PG0382 contains the conserved aspartate residue (Asp355). The amino acid residues following Asp355 in PG0382 are less well conserved with the adaptor proteins; however, the substituted amino acids share similar chemical properties. For instance, PG0382 Iso356, MAL Val88 and MYD88 Ala163 are aliphatic, hydrophobic amino acids. PG0382 and TLR adaptor proteins (except MAL), contain the conserved isoleucine residue in the box 1 sequence motif. The box 2 sequence motif of

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PG0382 was more similar to MAL and MYD88, than to TRAM, TRIF and SARM. Like MAL and MYD88, PG0382 contains the consensus aspartate (Asp390) and glycine (Gly394) residue. By contrast, the box 2 sequence motif in TRAM, TRIF and SARM are less well conserved. TRAM and TRIF only share the conserved glycine (Gly118 and Gly435, respectively) with PG0382. SARM lacks both the proline and glycine residues important for mediating TIR-TIR interactions. Like PG0382, the box 3 sequence motif in TLR adaptor proteins is also poorly defined. In conclusion, the PG0382 TIR domain contains a number of amino acid residues that are identical to the TIR domain consensus sequence in box 1 and box 2. In addition, the TIR domain of PG0382 was also found to be more similar to the TIR domain of TLR adaptor proteins than TLRs.

Table 5.5 Amino acid sequence identity (%) between PG0382 TIR domain and TIR domains of TLR protein adaptors.

PG0382 MYD88 MAL TRIF TRAM SARM

MYD88 10.9 - 25.5 7.8 12.6 14.0 MAL 15.2 25.5 - 12.8 22.9 12.7 TRIF 9.7 7.8 12.8 - 20.9 10.4 TRAM 15.0 12.6 22.9 20.9 - 9.2 SARM 14.8 14.0 12.7 10.4 9.2 -

box 1 box 2 F/YDAFISY RDxxPG

Figure 5.6 Multiple sequence alignment of PG0382 TIR domain with TLR adaptor proteins TIR domains. The consensus sequence for box 1 and box 2 are as indicated. The x in box 2 denotes any hydrophobic amino acid.

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Comparison of PG0382 with bacterial Tcps The TIR domain of PG0382 was also compared with the TIR domain of previously characterised bacterial Tcps (Table 5.6). The accession numbers of the bacterial Tcps are presented in Appendix Table A2. Pairwise sequence alignment revealed that PG0382 TIR domain shares highest sequence identity with the TIR domain of P. dentrificans PdTIR (25.3%), and lowest sequence identity with E. coli CFT037 TcpC (15.1%). The bacterial Tcps have sequence identities between 51.9% (PdTIR and TcpB) and 12.4 % (TcpC and YpTdp). Multiple sequence alignment of bacterial Tcp TIR domains showed that box 1 of bacterial Tcps were relatively similar between one another, and contained amino acid residues equivalent to the consensus sequence (Fig. 5.7). The bacterial Tcps all shared the conserved isoleucine (except TirS, which is substituted with leucine) and serine residues in box 1. The bacterial Tcps also have a histidine residue in box 1, while PG0382 contains a phenylalanine (Phe360) at that position. The conserved box 2 sequence motif of bacterial Tcps differs from the mammalian TIR domain consensus sequence. Like PG0382, the conserved proline residue in box 2 is absent in TcpB, TcpC and TirS. The conserved glycine residue in PG0382 is also present in the bacterial Tcps, except YpTdp. Box 3 is not present in the bacterial Tcps. Taken together, these analyses indicate that the PG0382 TIR domain has features similar to the TIR domain of other bacterial Tcps.

Table 5.6 Amino acid sequence identity (%) between PG0382 TIR domain and bacterial Tcp TIR domains.

PG0382 TcpB TcpC PdTIR YpTdp TirS

TcpB 19.2 - 51.1 51.9 17.3 29.3 TcpC 15.1 51.1 - 39.5 12.4 29.5 PdTIR 25.3 51.9 39.5 - 12.8 28.2 YpTdp 18.0 17.3 12.4 12.8 - 14.6 TirS 17.5 29.3 29.5 28.2 14.6 -

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box 1 box 2 F/YDAFISY RDxxPG

Figure 5.7 Multiple sequence alignment of PG0382 TIR domain with TIR domains of known bacterial TIR domain-containing proteins. The consensus sequence for box 1 and box 2 are as indicated. The x in box 2 denotes any hydrophobic amino acid.

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Structural prediction of PG0382 Mammalian TIR domains are structurally defined by a flavodoxin-fold comprised of five central -sheets and five -helices (Xu et al., 2000). In silico analysis was performed to determine whether the putative PG0382 TIR domain might adopt a similar structural conformation. FUGUE analysis (http://mizuguchilab.org/fugue/prfsearch.html), which aligns sequence- structure pairs to generate a list of structural homologs (Shi et al., 2001), was performed to first identify potential PG0382 structural homologs for tertiary structure modelling. This analysis revealed that PG0382 shares greatest structural homology with the TIR domain of PdTIR (Table 5.7). The TIR domains of human TLR1, TLR2, and MAL were also identified to be PG0382 structural homologs (Table 5.7). Interestingly, TIR domains in plants (e.g. Linum usitatissimum and Arabidopsis thaliana) were also identified as potential structural homologs of the PG0382 TIR domain (Table 5.7).

Table 5.7 Identification of PG0382 TIR domain structural homologs using FUGUE.

Protein PDB code Species Z-score

PdTIR 3H16 Paracoccus dentrificans 11.81

TLR1 TIR domain 1FYV Homo sapiens 9.85

MAL variant TIR domain 3UB4 Homo sapiens 8.43

L6 TIR domain 3OZI Linum usitatissimum 7.41

TLR2 variant TIR domain 1FYX Homo sapiens 7.07

RRS1 TIR domain 4C6S Arabidopsis thaliana 5.21

Nuclear receptor coactivator 5 1V95 Homo sapiens 4.81 binding domain

IL-17R SEFIR domain 3VBC Mus musculus 4.74

Homology modelling with SWISS-MODEL (https://swissmodel.expasy.org/) was performed next to produce a structural model based on protein sequence and model protein structures (Schwede et al., 2003). Given that PdTIR (PDB 3H16) was identified as a potential structural homolog for the TIR domain of PG0382 (Table 5.7), it was used as the template structure for modelling (Fig. 5.8A). As shown in Figure 5.8C, the model for PG0382 was predicted to consist of four -helices surrounding three -sheets. PG0382 was also modelled using the TIR domain of TLR1 (PDB 1FYV) as the template, for comparison (Fig. 5.8B). The resulting PG0382 homology

123 model generated suggested that the PG0382 TIR domain may form a similar / conformation, as when PdTIR was used as the structural template (Fig. 5.8D). The PdTIR TIR domain contains -helices that are more compact when compared to TLR1 TIR domain, but the overall structure of the PdTIR and TLR1 TIR domains are similar (Fig. 5.8A-B). In summary, the homology models suggest that the annotated TIR domain of PG0382 may adopt a TIR domain-like conformation.

PdTIR TIR domain TLR1 TIR domain

A B

PG0382 (PdTIR template) PG0382 (TLR1 TIR domain template)

C D

Figure 5.8 PG0382 structural prediction. Three dimensional models of the TIR domains of (A) PdTIR, (B) PG0382 modelled using PdTIR as a template, (C) TLR1, and (D) PG0382 modelled using TLR1 as a template. Box 1, box 2 and box 3 are as indicated by the yellow, green and pink colouring, respectively.

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Expression of PG0382 in HEK293T cells To better understand the biochemical properties of PG0382, mammalian expression plasmids encoding PG0382 and a mutant thereof lacking the TIR-like domain (i.e. PG0382ΔTD) were constructed (Fig. 5.9A). An expression plasmid encoding the TIR-like domain of PG0382 (i.e. PG0382TD) was created by another student (Ben Huang) in the laboratory. Briefly, the coding sequences for PG0382 and PG0382ΔTD were amplified by PCR, using P. gingivalis W50 genomic DNA as the template. The PCR products generated were subsequently cloned into the mammalian expression vector, pEF-V5, such that the expressed proteins contain an N-terminal V5 epitope tag. Following transformation into E. coli DH5α bacteria, positive transformants were identified by restriction endonuclease digestion (Fig. 5.9B-C). The absence of mutations in the cloned cDNA sequences was confirmed by DNA sequencing (performed at The Centre for Translational Pathology, The University of Melbourne).

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342 A 8 52 490

PG0382 NTPase N-terminus (coiled-coil motif) TIR domain

8 52 342

PG0382ΔTD NTPase N-terminus (coiled-coil motif)

342 490

PG0382TD TIR domain

V5 -

B - C

pEF

1 kb 1 Bioline DNA ladder

1 kb 1 NEB DNA ladder 3 Clone

Clone 1 Clone

Clone 4 Clone

Clone3 4 Clone

Clone2

Clone2 Clone5 Clone1 Size (kb) Size (kb)

10 8.0 3.0 4.0 2.0 3.0 1.0 2.0

1.5 0.2

Figure 5.9 Construction of pEF-V5-PG0382 and pEF-V5-PG0382ΔTD expression vectors. (A) Schematic representation of PG0382, PG0382ΔTD and PG0382TD. (B-C) Restriction endonuclease (HindIII) analyses of (B) pEF-V5-PG0382 and (C) pEF-V5-PG0382ΔTD. the bolded clones contain plasmid inserts with the correct orientation. The positions of DNA standards are as indicated.

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The expression of the proteins (PG0382, PG0382ΔTD, and PG0382TD) was tested by transient transfection of HEK293T cells, and subsequently subjecting lysates of the cells to Western blot analysis with an anti-V5 antibody. Other studies have demonstrated a propensity for bacterial Tcps to form dimers (Rana et al., 2011; Fekonja et al., 2012). Coiled-coil domains can drive oligomerisation of proteins (Low et al., 2007), and thus can affect protein solubility. PG0382 is predicted to contain a coiled-coil motif in the N-terminal half of the protein, and therefore both the detergent-soluble and detergent-insoluble fractions of the cells were analysed. Briefly, the cells were lysed with buffer containing 1% NP-40 (non-ionic detergent) and centrifuged to pellet insoluble material. The supernatant (representing the detergent-soluble fraction) was retained, whilst the pelleted material was solubilised with LDS sample buffer (representing the detergent-insoluble fraction). As shown in Figure 5.10, PG0382 and PG0382ΔTD were detected in both the detergent-soluble and detergent-insoluble fractions. Higher molecular weight bands were also detected for PG0382ΔTD (Fig. 5.10). Contrastingly, PG0382TD was found exclusively in the detergent-soluble fraction (Fig. 5.10), suggesting that the N-terminus of PG0382 may strongly influence its solubility under these experimental conditions. Full-length PG0382 was expressed at lower levels in comparison to PG0382ΔTD and PG0382TD (Fig. 5.10).

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Detergent-soluble Detergent-insoluble

Anti-V5 30

Anti-HSP90 80

Figure 5.10 Ectopic expression of V5-PG0382, V5-PG0382ΔTD, and V5-PG0382TD in HEK293T cells. HEK293T cells were transiently transfected with vectors expressing the indicated proteins, and lysed 24 h post-transfection. The detergent-soluble and detergent- insoluble fractions were subjected to Western blotting with an anti-V5 and anti-HSP90 (loading control) antibodies. The positions of molecular mass standards (in kDa) are as indicated. The data presented are representative of three independent experiments.

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Given the differences in their detergent-solubilities, PG0382, PG0382ΔTD, and PG0382TD were also investigated by immunofluorescence confocal microscopy; this also enabled examination of their subcellular localisation. PG0382 and PG0382ΔTD were detected in the cytoplasm, and appeared to form localised punctate bodies (Fig. 5.11). By contrast, PG0382TD exhibited dispersed cytoplasmic staining (Fig. 5.11). Collectively, the findings presented in Fig. 5.10 and Fig. 5.11 suggest that the structural elements in the N-terminal half of PG0382 may be important in dictating the solubility and subcellular localisation of PG0382.

Merged Anti-V5 DAPI

V5-PG0382

V5-PG0382ΔTD

V5-PG0382TD

Figure 5.11 Subcellular localisation of ectopically expressed PG0382, PG0382ΔTD, and PG0382TD in HEK293T cells. HEK293T cells were transiently transfected with vectors expressing the indicated proteins. The cells were fixed 24 h-post transfection, and stained with an anti-V5 antibody (red staining); nuclei were counterstained with DAPI (blue staining). The data presented are representative of three independent experiments. Scale bar = 10 µm.

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PG0382 expression causes the loss of MAL and MYD88 in HEK293T cells Bacterial Tcps have previously been shown to inhibit TLR signalling by interacting with MAL and MYD88 (Sengupta et al., 2010). Therefore, the effects of PG0382 on MAL and MYD88 were investigated. FLAG-tagged murine MAL and FLAG-tagged human MYD88 were transiently co-expressed with PG0382, PG0382ΔTD, or PG0382TD, in HEK293T cells. MAL was exclusively detected in the detergent-soluble fraction (Fig. 5.12A). However, MAL was not detected, in either the detergent-soluble or detergent-insoluble fraction, when co-expressed with PG0382 (Fig. 5.12A-B). MAL was not similarly affected when co-expressed with either PG0382ΔTD or PG0382TD (Fig. 5.12A-B). The effect of PG0382 on MYD88 was also investigated. Notably, MYD88 was exclusively detected in the detergent-insoluble fraction (Fig. 5.12C). MYD88 levels were greatly reduced when co-expressed with PG0382, whilst PG0382ΔTD and PG0382TD did not affect MYD88 levels (Fig. 5.12C-D). The mRNA expression levels of MAL and MYD88 were also measured by Real-time PCR to determine whether PG0382 might have affected the transcription of the MAL and MYD88 expression plasmids. Because murine MAL was used in this study, the level of overexpressed MAL was not directly comparable with endogenous MAL (Fig. 5.13A). MYD88 mRNA levels were increased by ~100-fold when transfected with the MYD88 expression plasmid (Fig. 5.13B). As shown in Figure 5.13A-B, co-expression with PG0382 did not cause a reduction in the mRNA levels of ectopically expressed MAL or MYD88. These data suggest that PG0382 causes the loss of MAL and MYD88 protein when co-expressed in HEK293T cells.

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A Detergent-Soluble Detergent-Insoluble

FLAG-MAL + + + + - + + + + - V5-PG0382 - + - - - - + - - - V5-PG0382 ΔTD - - + - - - - + - - V5-PG0382 TD - - - + - - - - + - 30 Anti-FLAG B 50

Anti-V5 30

Anti-HSP90 80

C Detergent-Soluble Detergent-Insoluble

FLAG-MYD88 + + + + - + + + + - V5-PG0382 - + - - - - + - - - V5-PG0382 ΔTD - - + - - - - + - - V5-PG0382 TD - - - + - - - - + -

Anti-FLAG 30 D

Anti-V5 30

Anti-HSP90 80

Figure 5.12 Effects of PG0382 on MAL and MYD88 expression. HEK293T cells were transiently transfected vectors expressing the indicated proteins, and were lysed 24 h-post transfection. (A and C) The detergent-soluble and detergent-insoluble fractions were subjected to Western blotting. The positions of molecular mass standards (in kDa) are as indicated. The data presented are representative of three independent experiments. (B and D). The protein levels of MAL from (A) and MYD88 from (C) were quantified by densitometric analysis. (B) MAL and (D) MYD88 transfected alone were given an arbitrary value of 100%. All data are presented as the mean ± SEM (*=p<0.05, ***=p<0.001).

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A B

N D

Figure 5.13 Effects of PG0382 on MAL and MYD88 mRNA levels in HEK293T cells. HEK293T cells were transiently transfected with the vectors expressing the indicated proteins for 24 h. (A) MAL and (B) MYD88 mRNA levels were measured by Real-Time PCR, and are shown as relative to HPRT (endogenous control gene) (n=3). All data are presented as the mean ± SEM (ND=not detected, *=p<0.05).

Given the findings above, the effects of PG0382 on MAL and MYD88 were further investigated in titration experiments, where increasing amounts of the PG0382 expression plasmid were co-transfected with the MAL or MYD88 expression plasmids. MAL protein was reduced by approximately 90%, even at the lowest PG0382 plasmid concentration tested (Fig. 5.14A-B). MYD88 was less susceptible to the effects of PG0382, in comparison to MAL. MYD88 protein expression was reduced by approximately 50% when co-transfected with the lowest amount of PG0382 plasmid, and was reduced by approximately 90% at the highest amount of PG0382 plasmid transfected (Fig. 5.14C-D). Taken together, these findings suggest that the MAL is more susceptible to the effects of PG0382 than MYD88.

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A MAL (0.1 µg) + + + + - B PG0382 (µg) 0 0.1 0.4 1 0

MAL 30

PG0382 50

HSP90 80

C MYD88 (0.1 µg) + + + + - D PG0382 (µg) 0 0.1 0.4 1 0

MYD88 30

PG0382 50

HSP90 80

Figure 5.14 Concentration-dependent effects of PG0382 on MAL and MYD88 protein expression. HEK293T cells were transiently transfected with vectors expressing the indicated proteins, and were lysed 24 h-post transfection. (A and C) The detergent-soluble and detergent- insoluble fractions were subjected to Western blotting. The detergent-soluble fraction was probed for MAL and HSP90, whilst the detergent-insoluble fraction was probed for MYD88 and PG0382. The positions of molecular mass standards (in kDa) are as indicated. The data presented are representative of three independent experiments. (B and D) The protein levels of MAL from (A) and MYD88 from (C) were quantified by densitometric analysis. (B) MAL and (D) MYD88 transfected alone were given an arbitrary value of 100%. The data are presented as the mean ± SEM (***=p<0.001).

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Lack of complex formation between PG0382TD and MAL The ability of PG0382 to form a stable complex with MAL was next investigated. Its ability to likewise interact with MYD88 was not investigated because denaturing conditions were required to recover MYD88 from cell lysates. Additionally, because PG0382 causes the loss of MAL protein, PG0382TD was used to determine whether the TIR domain of PG0382 can interact with MAL. Briefly, FLAG-tagged murine MAL was co-expressed with V5-PG0382 in HEK293T cells, and lysates of the cells were subsequently subjected to immunoprecipitation with an anti-FLAG antibody. As shown in Figure 5.15, FLAG-MAL was successfully immunoprecipitated from the detergent-soluble cell lysates. However, V5-PG0382TD was not detected in the anti-FLAG immunoprecipitates. PG0382TD was detected in the detergent-soluble cell lysates (Fig. 5.15), indicating that the protein had been successfully expressed. This suggests that the absence of complex formation between the PG0382TD and MAL is unlikely to be due to poor expression of PG0382TD. Therefore, it was concluded that the TIR domain of PG0382 does not form a stable complex with MAL.

FLAG-IP V5-PG0382TD - - + FLAG-MAL - + + 20 Anti-V5

30 Anti-FLAG

Input 20 Anti-V5

Figure 5.15 Analysis of PG0382TD and MAL interaction by co-immunoprecipitation. HEK293T cells were transiently transfected with vectors expressing the indicated proteins, and lysed 24 h-post transfection. FLAG-MAL was immunoprecipitated from the cell lysates with anti-FLAG antibodies, followed by Western blotting with anti-V5 and anti-FLAG antibodies. The lysates (input) were subjected to Western blotting with an anti-V5 antibody. The positions of molecular mass standards (in kDa) are as indicated. The data is representative of three independent experiments.

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Subcellular localisation of PG0382TD, MAL and MYD88 Complex formation by proteins might not be detectable by immunoprecipitation assay if the proteins form weak or transient interactions. Therefore, immunofluorescence confocal microscopy was also performed to investigate whether PG0382TD might co-localise or affects the subcellular localisation of MAL and/or MYD88. When expressed in HEK293T cells, MAL appeared to be largely localised towards the cell periphery (Fig. 5.16A). When co-expressed with PG0382TD, MAL remained localised around the cell periphery, and did not co-localise with PG0382TD (Fig. 5.16A). In contrast to MAL, MYD88 was observed to form distinct foci throughout the cytoplasm (Fig. 5.16B), potentially consistent with MYD88 forming detergent- resistant homo-oligomers. The subcellular localisation of MYD88 was not affected by the co-expression of PG0382TD, and nor did MYD88 co-localise with PG0382TD (Fig. 5.15B). Taken together, these data suggest that PG0382 does not co-localise with MAL or MYD88, or affect their subcellular localisation.

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A Merged Anti-FLAG Anti-V5

FLAG-MAL

FLAG-MAL and V5-PG0382TD

Merged Anti-FLAG Anti-V5 B

FLAG-MYD88

FLAG-MYD88 and V5-PG0382TD

Figure 5.16 Co-localisation of PG0382TD with MAL and MYD88 by immunofluorescence. HEK293T cells were transiently transfected with vectors expressing the indicated proteins. The cells were fixed 24h-post transfection, and were stained with an anti-FLAG antibody (green staining), and an anti-V5 antibody (red staining); nuclei were counterstained with DAPI (blue staining). The data presented are representative of three independent experiments. Scale bar = 10 µm.

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5.3 Discussion TLRs are integral to the host innate immune system where they play important roles in the recognition of pathogens and initiating the ensuing immune response. Upon recognition of their cognate ligand, TLRs trigger a series of intracellular signalling cascades that culminate in the stimulation of host inflammation. Notably, the intracellular TIR domains of TLRs are critical for facilitating downstream signalling because they recruit TIR domain-containing signalling adaptor proteins, such as MAL and MYD88, into the receptor complex. Given the critical roles of TLRs, pathogens have evolved mechanisms to suppress TLR signalling. Bacterial Tcps have been proposed to be a subversion mechanism to block the interaction of TLRs with downstream adaptor proteins. Indeed, in vivo studies have demonstrated important roles for Tcps in bacterial virulence and dissemination (Cirl et al., 2008; Newman et al., 2006; Radhakrishnan et al., 2009). Tcp(s) have yet to be defined in P. gingivalis. Thus, potential P. gingivalis Tcps were identified and characterised in this Chapter.

The InterPro database integrates amino acid “signatures” representing protein domains from multiple source databases, such as Pfam and Prosite, into a single consortium (Hunter et al., 2009). The database can therefore provide a global overview of identified and annotated protein domains across various species. This approach led to the identification of eleven putative Tcps, across nine different strains, in P. gingivalis. The putative Tcps were classified into two distinct groups, which were denoted, Group A and Group B. Proteins in Group A had low overall amino acid sequence conservation (5-20%), and shared approximately 20% sequence identity between TIR domains. For comparison, mammalian TIR domains typically share around 20-30% sequence conservation (Xu et al., 2000). Therefore, it was not surprising that the TIR domains of Group A Tcps exhibited relatively low sequence conservation. The low degree of sequence conservation in TIR domains of TLRs and adaptor proteins is proposed to provide the domains with sufficient structural diversity to enable specific TIR-TIR interactions (Xu et al., 2000). In contrast to the Tcps in Group A, Group B Tcps shared a higher level of amino acid sequence conservation (60-80%), and the TIR domains have sequence identities greater than 70%. In addition, NCBI Conserved Domain searches revealed that Group B Tcps also have similar domain arrangements, suggesting that they may have similar functions. Cluster analysis suggests that the distribution of TIR domains in bacteria occurred through horizontal gene transfer (Zhang et al., 2011), however the relationship between the two groups of P. gingivalis Tcps is unclear. Further studies will be required to determine whether the proteins from the two groups are involved in similar biological functions.

Despite having low overall amino acid sequence conservation, TIR domains contain conserved sequence motifs, namely box 1, box 2 and box 3, which have been shown to be important for

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TLR signalling. Multiple sequence alignment analysis revealed that PG0382 shares conserved amino acid residues with TLRs, TLR adaptor proteins, and bacterial Tcps in box 1. Box 2 of PG0382 was found to be more similar to TLR adaptor proteins (e.g. MAL and MYD88) and bacterial Tcps than TLRs. PG0382 lacks box 3, which is also absent in TLR adaptor proteins and bacterial Tcps. The role of box 2 is best understood and has been shown to be important for mediating heterotypic TIR-TIR interaction (Poltorak et al., 1998; Slack et al., 2000). PG0382 lacks the invariant proline residue found in box 2. Mutation of the proline residue rendered TLR4 hyporesponsive to LPS stimulation (Poltorak et al., 1998). However, the mutation did not prevent TLR4 from interacting with MAL or MYD88 (Dunne et al., 2003). This contrasts with the effect of mutating the corresponding proline residue in TLR2, which inhibited MYD88 binding (Xu et al., 2000). This implies that the conserved proline residue may be important for surface properties (e.g. surface charge) required for some TIR-TIR interactions, but not others. PG0382 contains the consensus box 2 glycine residue conserved in TLRs, adaptor proteins and bacterial Tcps. Mutation of the conserved glycine residue (Gly158 to Ala) in TcpB impaired the inhibition of TLR2-mediated NF-B activation (Alaidarous et al., 2014; Radhakrishnan et al., 2009). Similarly, the conserved glycine was also found to be important for SARM-mediated immune suppression (Carlsson et al., 2016). Similar mutagenesis studies will be required to determine the importance of the conserved box 2 residues (e.g. Gly394) for PG0382 function. Nonetheless, PG0382 contains TIR domain sequence features similar to TLR adaptor proteins and previously characterised bacterial Tcps.

Structurally, TIR domains are comprised of five central -sheets and five -helices, described as a flavadoxin fold (Xu et al., 2000). Structural homology modelling suggested that the annotated TIR domain in PG0382 might adopt a similar flavadoxin fold, with central -sheets surrounded by -helices. The prediction of coiled-coil motifs within the PG0382 TIR domain (Gly350-Glu400, and Thr450-Gln490) is consistent with the ability of PG0382 TIR domain to form -helices, because coiled-coil motifs, like -helices, form secondary helical structures. In addition, box 1 and box 2 in PG0382 were predicted to adopt similar secondary conformations compared to resolved TIR domain crystal structures. However, the generation of a reliable protein structure model with known structural templates typically requires >30% amino acid sequence identity (Sánchez and Šali, 1997). The relatively low sequence conservation between the TIR domain of PG0382 and structural homologs identified (e.g. PdTIR TIR domain and TLR1 TIR domain) thus posed as a restriction on generating an accurate structural prediction of PG0382. Moreover, the models are limited by the fact that they are built around existing structures and hence are restricted by computational algorithms. Therefore, laboratory-based

138 experimental approaches (e.g. circular dichroism and x-ray crystallography) with purified PG0382 will likely be required to provide further insight into the structure of PG0382.

The COILS prediction analysis suggested that PG0382 may contain a coiled-coil motif within the N-terminal half (Gln200-Lys250) of the protein. Other bacterial Tcps that have been studied for their ability to suppress TLR-mediated immune responses, including TcpC, TcpB, and YpTdp, also possess coiled-coil motifs (Alaidarous et al., 2014; Cirl et al., 2008; Rana et al., 2011). The primary role of a coiled-coil domain is to mediate protein oligomerisation (Burkhard et al., 2001). For example, purified full-length PdTIR, as well as its isolated N-terminal coiled-coil domain, have been shown to exist in monomers and oligomers (Low et al., 2007). Therefore, it is tempting to speculate that the predicted coiled-coil motif in PG0382 may likewise promote its oligomerisation. Notably, the detection of PG0382 and PG0382ΔTD in the detergent-insoluble fraction of transfected HEK293T cells, and their subcellular distribution as punctate bodies, is potentially consistent with PG0382 forming dimers/oligomers. TirS from S. aureus has also been shown to be highly insoluble (Patot et al., 2017). Interestingly, the addition of an artificial coiled-coil motif to the MYD88 TIR domain was shown to inhibit TLR4 activation in vitro (Fekonja et al., 2012). The addition of a coiled-coil motif was proposed to not only promote MYD88 TIR domain dimerisation, and thereby provide a larger interface to interact with dimerised TLRs, but also to sterically hinder the recruitment of downstream TLR adaptor proteins (Fekonja et al., 2012). Therefore, bacterial Tcps may have adopted coiled-coil motifs to potentiate the inhibition of TLR signalling. Further biochemical studies, such as size exclusion chromatography and sedimentation equilibrium analysis, will be required to gain direct insight into the ability of PG0382 to form dimers/oligomers.

Other studies have demonstrated the ability of bacterial Tcps to interact with MAL and MYD88 as a mechanism to inhibit NF-B signalling (Alaidarous et al., 2014; Askarian et al., 2014; Cirl et al., 2008; Newman et al., 2006; Sengupta et al., 2010). The results presented in this study revealed that co-expression with PG0382 in HEK293T cells significantly reduced the protein expression of MAL and MYD88. This effect likely occurred at the protein level, because PG0382 did not similarly affect MAL or MYD88 mRNA expression levels. The interaction of TcpB with MAL was shown to promote MAL ubiquitination, leading to its rapid proteasomal degradation (Sengupta et al., 2010). The mechanism whereby PG0382 causes MAL and MYD88 protein levels to be markedly reduced is unclear at this time. Co-immunoprecipitation and immunofluorescence confocal microscopy experiments suggested that the PG0382 TIR domain does not interact with MAL or MYD88. Chemical cross-linking reagents (e.g. disuccinimidyl suberate) can be used in co-immunoprecipitation assays to “capture” weak/transient interactions, and thus could be used to further investigate the ability of PG0382 to bind MAL or

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MYD88. However, it is important to appreciate that cross-linking reagents can potentially result in the “capturing” of artefactual protein complexes that do not form under native conditions. It is also possible that other regions/domains of PG0382 might be required for its interaction with MAL and MYD88. Therefore, further studies will be required to elucidate the effects of PG0382 on MAL and MYD88.

In summary, the bioinformatics analyses performed in this Chapter identified eleven putative P. gingivalis Tcps, which can be divided into two groups with distinctive characteristics. Further bioinformatics analysis of P. gingivalis PG0382 revealed that its putative TIR domain contains sequence features similar to TLR adaptor proteins and bacterial Tcps. In addition, homology modelling suggests that PG0382 may adopt a TIR-like structure. Further characterisation by transient expression in HEK293T cells and analysis by Western blotting and immunofluorescence confocal microscopy showed that the N-terminus of PG0382 exhibits properties consistent with it promoting oligomerisation. Significantly, the co-expression of PG0382 reduced MAL (and MYD88) protein levels, but the PG0382 TIR domain does not appear to directly interact with MAL. Therefore, further studies will be required to better understand the effects of PG0382 on MAL and MYD88.

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Investigation of PG0382 and host inflammation

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6.1 Introduction The oral epithelium expresses TLRs (e.g. TLR2) that enable the detection of potential pathogens and stimulation of host inflammation. The activation of TLR signalling in oral epithelial cells leads to the production of inflammatory mediators, including cytokines and chemokines, which promote the recruitment of immune cells. Neutrophils and inflammatory monocytes are typically the first immune cells that respond to infection by pathogens. Collectively, neutrophils and inflammatory monocytes mount a robust antimicrobial response. They also activate and direct adaptive immune cells (e.g. lymphocytes) to generate an appropriately tailored immune response. Together, the two arms of the immune system collaborate to efficiently eliminate potential pathogens.

Bacterial TIR domain-containing proteins (Tcps) have been proposed to subvert the host immune response by suppressing TLR signalling. Several in vitro studies have demonstrated the ability of bacterial Tcps to inhibit the activation of the transcription factor NF-B (Cirl et al., 2008; Newman et al., 2006; Radhakrishnan et al., 2009). Importantly, blockade of NF-B activity potentially translates to reduced inflammatory cytokine production. Consistently, macrophages mounted enhanced cytokine responses when infected with a TcpC-deficient E. coli CFT037 mutant, in comparison to infection with wildtype E. coli CFT037(Cirl et al., 2008). Moreover, in vivo studies have demonstrated a role for bacterial Tcps in virulence. For example, mice infected with a B. melitensis TcpB-deficient mutant had prolonged survival, when compared to mice infected with wildtype B. melitensis (Salcedo et al., 2013). TcpC was also shown to be important for the survival of E. coli CFT037 in mice, and induction of kidney pathology in a mouse model of urinary tract infection (Cirl et al., 2008). In this Chapter, in vitro and in vivo systems were used to investigate a role for the putative P. gingivalis Tcp, PG0382, in modulating the host inflammatory response.

6.2 Results Generation of an isogenic P. gingivalis PG0382-deficient mutant To study the function of PG0382, an isogenic P. gingivalis PG0382-deficient mutant (ΔPG0382) was created (Fig. 6.1). Purified genomic DNA from P. gingivalis W50 was used as the template to generate PCR products from the 5’ and 3’ intergenic regions of the PG0382 gene (Fig. 6.2A). Similarly, PCR was also used to amplify the erythromycin-resistance gene (ermF) from the pAL30 plasmid (Dashper et al., 2009). The PCR reactions were conducted with primers containing complementary sequence overhangs to subsequently enable a single, linear PCR product to be generated by splice-overlap extension PCR (SOE PCR) (Fig. 6.1). The resulting SOE PCR product was comprised of the ermF gene, flanked by 5’ and 3’ intergenic regions of the

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PG0382 gene (Fig. 6.2B). The SOE PCR product was then introduced into P. gingivalis W50 by electroporation. Successful integration of the SOE PCR product into the P. gingivalis genome occurs through homologous recombination with the 5’ and 3’ intergenic regions of the PG0382 gene. Transformed P. gingivalis were selected on blood agar plates supplemented with erythromycin. Genomic DNA from positive transformants was then purified, and PCR primers specific for the 5’ and 3’ intergenic regions of the PG0382 gene were used to amplify the corresponding region of the P. gingivalis genome for DNA sequencing to confirm that the PG0382 gene had been replaced with the ermF gene (Fig. 6.2C).

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P. gingivalis W50 genomic DNA pAL30

5’ intergenic 3’ intergenic region PG0382 region ermF

PG0382_IG5 PG0382_IG3 ermF_PG0382 PCR products

Splice Overlap Extension PCR

5’ intergenic 3’ intergenic region ermF region

Electroporation

5’ intergenic 3’ intergenic region ermF region

Homologous recombination PG0382

Selection

5’ intergenic 3’ intergenic region ermF region

Figure 6.1 Strategy for the generation of an isogenic P. gingivalis PG0382-deficient mutant.

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A B

PG0382_IG3

PG0382_IG5

ermF_PG0382

1 kb ladder 1ladder kb product SOE PG0382 Size (bp) ladder bp 100 Size (bp) 1000 4000 2000 500 300 1000 500

250/253

Clone 1 Clone

Clone 2 Clone

Clone 3 Clone 4 Clone 1 kb ladder 1ladder kb Size (bp) Wildtype

4000 2000

1000

500 250/253

Figure 6.2 Gel electrophoresis analysis of PCR products. (A-B) Gel electrophoresis analysis of the indicated PCR products. (C) Genomic DNA from positive P. gingivalis transformants were purified and used as a template for PCR amplification. The PCR products were then analysed by gel electrophoresis. Bolded clones contain ermF integrated into the positive P. gingivalis transformants. The positions of DNA standards are as indicated.

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Phenotypic characterisation of P. gingivalis ΔPG0382 P. gingivalis ΔPG0382 growth properties were first characterised to ensure that the deletion of PG0382 does not affect P. gingivalis phenotype. The mean generation time of P. gingivalis ΔPG0382 was determined to assess whether the absence of PG0382 might affect the growth of the bacterium. The mean generation time for P. gingivalis ΔPG0382 was 3.3±0.1 h, which was comparable to the mean generation time of 3.0±0.2 h for wildtype P. gingivalis (Fig. 6.3). The latter finding is consistent with published studies, which also established that wildtype P. gingivalis has a mean generation time of approximately 3 h (Aruni et al., 2011; Fletcher et al., 1995). Therefore, the deletion of the PG0382 gene does not affect the growth rate of P. gingivalis, at least under the in vitro conditions used.

P. gingivalis Mean generation time (h) Wildtype 3.0±0.2 ΔPG0382 3.3±0.1

Figure 6.3 Growth rates of wildtype P. gingivalis and P. gingivalis ΔPG0382. The mean generation time was calculated based on the rate of change of optical density (OD650) (n=3). The data are presented as the mean ± SEM. P. gingivalis forms black-pigmented colonies when cultured on horse blood agar plates due to the degradation of haemoglobin by the gingipain proteases (Smalley et al., 1998; Zambon et al., 1981). The gingipain proteases are also major components of the electron-dense surface layer (EDSL) of the outer membrane of P. gingivalis (Chen et al., 2011). The effects of deleting the PG0382 gene on colony pigmentation and the EDSL were therefore examined. Like wildtype P. gingivalis (Fig. 6.4A), P. gingivalis ΔPG0382 also grew as black-pigmented colonies when cultured on horse blood agar plates (Fig. 6.4B). Cryo-electron microscopy was performed to determine if the PG0382 gene is required for EDSL formation. Like wildtype P. gingivalis (Fig. 6.4C), P. gingivalis ΔPG0382 had a distinct EDSL associated with the outer membrane

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(Fig. 6.4D). Collectively, these results suggest that the absence of PG0382 does not affect these defining characteristics of P. gingivalis.

A Wildtype P. gingivalis B P. gingivalis ΔPG0382

C Wildtype P. gingivalis D P. gingivalis ΔPG0382 EDSL OM IM

EDSL OM IM

Figure 6.4 Phenotypic characterisation of P. gingivalis ΔPG0382. Images of (A) wildtype P. gingivalis and (B) P. gingivalis ΔPG0382 cultured on 10% defibrinated horse blood agar plates for 7 days at 37°C in anaerobic conditions. (C-D) EDSL of (C) wildtype P. gingivalis and (D) P. gingivalis ΔPG0382 were examined by cryo-EM. EDSL = Electron surface dense layer, OM = outer membrane, and IM = inner membrane. Scale bar = 200 nm.

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Gingipain protease activity of P. gingivalis ΔPG0382 The findings above suggested that deletion of the PG0382 gene does not affect the expression and processing of the gingipain proteases. Nonetheless, this was also directly assessed by measuring the Kgp and RgpA/B proteolytic activity of P. gingivalis ΔPG0382. P. gingivalis ΔPG0382 was cultured to exponential growth phase, and both whole-cells and cell-free culture supernatants were then harvested to assay Kgp and RgpA/B activity. The whole-cell Kgp activity of P. gingivalis ΔPG0382 was comparable to wildtype P. gingivalis (Fig. 6.5A). Similarly, the cell-free culture supernatants derived from P. gingivalis ΔPG0382 and wildtype P. gingivalis exhibited comparable Kgp activity (Fig. 6.5B). Deletion of the PG0382 gene also did not affect whole-cell RgpA/B activity (Fig. 6.5C) or RgpA/B activity in cell-free supernatants (Fig. 6.5D). Therefore, it can be concluded that deletion of the PG0382 gene does not affect the cell-surface attachment or secretion of Kgp or RgpA/B.

A B

C D

Figure 6.5 P. gingivalis gingipain protease activity. (A-B) Kgp and (C-D) RgpA/B proteolytic activity in (A and C) whole-cells and (B and D) cell-free culture supernatants from wildtype P. gingivalis and P. gingivalis ΔPG0382 were measured (n=3). All data are presented as the mean ± SEM.

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Stimulation of inflammatory gene responses in oral epithelial cells by P. gingivalis ΔPG0382 The bioinformatics and functional biochemical analyses in Chapter 5 suggest that PG0382 contains TIR-like properties comparable to bacterial Tcps that have been shown to suppress TLR-mediated inflammatory cytokine responses. Therefore, a role for PG0382 in modulating inflammatory cytokine responses in oral epithelial cells was investigated. OKF6 cells were challenged with either wildtype P. gingivalis or P. gingivalis ΔPG0382, and changes in the mRNA expression levels of inflammatory cytokines were then assessed. Fimbriae from P. gingivalis have previously been shown to stimulate IL-8 expression in a TLR2-dependent manner in human gingival epithelial cells (Asai et al., 2001; Eskan et al. 2007). Thus, the stimulation of IL-8 expression in OKF6 cells by P. gingivalis ΔPG0382 was investigated. As shown in Figure 6.6A, no difference was found between P. gingivalis ΔPG0382 and wildtype P. gingivalis in the stimulation of IL-8 gene expression. Recent work by our laboratory has demonstrated that TLR2 signalling in OKF6 cells also regulates the expression of IL-36G in response to P. gingivalis (Huynh et al., 2016). Therefore, the ability of P. gingivalis ΔPG0382 to stimulate IL-36G expression was investigated. The stimulation of IL-36G gene expression by P. gingivalis ΔPG0382 was comparable to that by wildtype P. gingivalis (Fig 6.6B). Tumour necrosis factor alpha-induced protein 3 (TNFAIP3) expression is also induced via TLR signalling, and functions to negatively regulate NF-B signalling (Boone et al., 2004). Both P. gingivalis ΔPG0382 and wildtype P. gingivalis were found to stimulate comparable TNFAIP3 responses (Fig. 6.6C). Taken together, these data suggest that PG0382 does not suppress the stimulation of inflammatory gene expression in oral epithelial cells (e.g. OKF6 cells) by P. gingivalis, although only a small number of genes were examined.

A B C

Figure 6.6 Effects of P. gingivalis ΔPG0382 on inflammatory responses of oral epithelial cells. OKF6 cells were challenged with wildtype P. gingivalis or P. gingivalis ΔPG0382 at 100 MOI for 24 h. (A) IL-8, (B) IL-36G, and (C) TNFAIP3 mRNA levels were then measured by Real- Time PCR, and are shown as a fold change relative to mock-challenged cells (n=3). All data are presented as the mean ± SEM (* = p <0.05).

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Stimulation of inflammatory gene responses in macrophages by P. gingivalis ΔPG0382 Macrophages are critical mediators of host inflammation, and TLR2 and TLR4 have been shown to be important regulators of the inflammatory responses of macrophages to P. gingivalis (Holden et al., 2014; Papadopoulos et al., 2013). Therefore, the effect of PG0382 deletion on P. gingivalis-inducible macrophage inflammatory responses was investigated with the murine macrophage cell line, RAW 264.7 (Raschke et al., 1978). TNF and IL-6 levels closely correlate with disease progression in chronic periodontitis, and are induced by P. gingivalis (Graves, 2008; Graves and Cochran, 2003), therefore the stimulation of TNF and IL-6 by P. gingivalis were examined. Wildtype P. gingivalis stimulated strong TNF gene expression (>10-fold) in RAW 264.7 cells (Fig. 6.7A). P. gingivalis ΔPG0382 stimulated similar upregulation of TNF gene expression, and accordingly, there was no difference in the stimulation of TNF expression between wildtype P. gingivalis and P. gingivalis ΔPG0382 (Fig. 6.7A). As shown in Figure 6.7B, wildtype P. gingivalis stimulated robust IL-6 gene expression (>150-fold) in RAW 264.7 cells, albeit more slowly than TNF (Fig. 6.7A). P. gingivalis ΔPG0382 also stimulated strong IL-6 gene expression (Fig. 6.7B). Although the IL-6 responses stimulated by wildtype P. gingivalis and P. gingivalis ΔPG0382 were not statistically different, the trend for all experiments was the same, whereby P. gingivalis ΔPG0382 stimulated weaker IL-6 responses. The expression of the monocyte/macrophage chemokine, CCL2, was also found to be strongly induced by both wildtype P. gingivalis and P. gingivalis ΔPG0382, with no statistically significant differences between the responses (Fig. 6.7C). However, the trend for all experiments revealed a weaker CCL2 response when the cells were challenged with P. gingivalis ΔPG0382 (Fig. 6.7C). The anti-inflammatory cytokine, IL-10, plays an important role in limiting inflammatory responses (Couper et al., 2008). IL-10 gene expression was induced (5 to 10-fold) by both wildtype P. gingivalis and P. gingivalis ΔPG0382 (Fig. 6.7D). Notably, the IL-10 response 24 h-post challenge was weaker with P. gingivalis ΔPG0382 (Fig. 6.7D). Overall, these data suggest that PG0382 may affect the inflammatory responses elicited in macrophages by P. gingivalis.

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A B

C D

Figure 6.7 Effects of P. gingivalis ΔPG0382 on inflammatory cytokine responses of macrophages. RAW 264.7 cells were challenged with wildtype P. gingivalis or P. gingivalis ΔPG0382 at 100 MOI for up to 24 h. (A) TNF, (B) IL-6, (C) CCL2, and (D) IL-10 mRNA levels were then measured by Real-Time PCR, and are shown as a fold change relative to time- matched and mock-challenged cells (n≥3). All data are presented as the mean ± SEM (* = p<0.05, ** = p<0.01, *** = p<0.001). Innate immune response to P. gingivalis ΔPG0382 in mice Given the limitations of in vitro systems for studying immunological responses, an in vivo approach was also undertaken to investigate the potential role of PG0382 in modulating the host immune response to P. gingivalis. Due to time limitations towards the end of my PhD candidature, a pilot study was undertaken to determine whether PG0382 may affect the recruitment of host immune cells in response to P. gingivalis. Briefly, wildtype P. gingivalis and P. gingivalis ΔPG0382 were injected into the peritoneal cavity of BALB/c mice, and intraperitoneal fluid was then harvested 6 or 24 h-post infection. The cells recovered in the peritoneal lavage fluid were stained with fluorochrome-conjugated antibodies, and immune cell populations of interest (e.g. macrophages, inflammatory monocytes, and neutrophils) were then quantified by fluorescence-activated cell sorting (FACS) analysis. A representative FACS-gating strategy is shown in Figure 6.8. Debris, cell doublets and aggregates were excluded from the analyses. Fixable viability dye 700 (FVS700) is a cell-impermeable dye that reacts with free

151 amines in the cytoplasm of dead cells (Perfetto et al., 2006), and hence was used to exclude dead cells from the analysis.

Cells Single cells Live cells

H -

FSC SSC FSC

FSC-A FSC-A FVS700

Activated

macrophages

+ + (F4/80 CD86 ) 80

Inflammatory / F4 F4/80 monocytes lo + (F4/80 Ly6C )

Ly6C CD86

Neutrophils + lo

(Ly6G Ly6C )

Ly6G

Ly6C

Figure 6.8 FACs Gating strategy for identifying innate immune cells. Single cells were analysed by excluding debris (top left panel) and cell aggregates (top centre panel). Live/dead discrimination was determined using Fixable Viability Dye 700 (top right panel). Cells were stained with fluorochrome conjugated mouse antibodies to identify F4/80+ CD86+ activated macrophages and F4/80lo Ly6C+ inflammatory monocytes. Ly6G+ Ly6Clo neutrophils were identified by excluding macrophages and monocytes from the analyses.

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Tissue-resident macrophages are critical immune sentinels; therefore, the proportion of macrophages that were activated in response to intraperitoneal infection by P. gingivalis was measured. F4/80 is a unique, mouse macrophage cell-surface marker (Austyn and Gordon, 1981) and routinely used for the identification of macrophage populations by FACS analysis. Activated macrophages can be identified based on increased expression levels of CD86, a costimulatory molecule involved in T lymphocyte activation (Mosser and Edwards, 2008). Thus, cells that were double-positive for F4/80 and CD86 (i.e. F4/80+, CD86+) were defined as activated macrophages. The analysis showed that the numbers of activated macrophages from mice infected with 5×106 wildtype P. gingivalis or P. gingivalis ΔPG0382 were similar to sham-infected mice, at either 6 or 24 h post-infection (Fig. 6.9A-B). The infection of mice with a larger inoculum of P. gingivalis (e.g. 5×107) did not appear to affect the numbers of activated macrophages (Fig. 6.9A-B).

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P. gingivalis P. gingivalis P. gingivalis ΔPG0382 P. gingivalis ΔPG0382 A 6 Sham 5×10 5×107 5×106 5×107

6 h

F4/80

F4/80 F4/80 F4/80

CD86 CD86 CD86 CD86 CD86

24 h

F4/80

F4/80 F4/80 F4/80

CD86 CD86 CD86 CD86 CD86 B Figure 6.9 Activation of macrophages in response to P. gingivalis. BALB/c mice were infected intraperitoneally with wildtype P. gingivalis or P. gingivalis ΔPG0382. Mouse peritoneal fluid was harvested 24 h or 48 h-post injection and subjected to flow cytometric analysis. (A) F4/80+ CD86+ FACS plots from individual mice. NB: Due to the variation in response between mice, the FACs plots presented do not necessarily represent the “average” response. (B) Quantification of F4/80+ CD86+ activated macrophages. Each data point represents an individual mouse (n=5).

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In addition to tissue-resident macrophages, inflammatory monocytes are recruited to sites of infection to reinforce the host immune response. Inflammatory monocytes express low levels of F4/80 but high levels of Ly6C (i.e. F4/80lo, Ly6C+) (Geissmann et al., 2003, 2008) and were identified on that basis. For most mice, infection with wildtype P. gingivalis or P. gingivalis ΔPG0382 stimulated an increase in the numbers of inflammatory monocytes by 6 h-post infection (Fig. 6.10A-B). However, the increases in inflammatory monocytes were not statistically significant due to the variation in responses between mice, specifically, the responses by mice infected with 5×106 wildtype P. gingivalis. (The increases are statistically significant if the two mice that did not respond to infection with 5×106 wildtype P. gingivalis are excluded from the analysis). Infection with 5×107 wildtype P. gingivalis or P. gingivalis ΔPG0382 did not stimulate a further increase in the numbers of inflammatory monocytes (Fig. 6.10B), suggesting that infection with 5×106 P. gingivalis induces a maximal response in this model. Notably, the numbers of inflammatory monocytes recruited in response to P. gingivalis was not affected by deletion of the PG0382 gene (Fig. 6.10B).

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P. gingivalis P. gingivalis P. gingivalis ΔPG0382 P. gingivalis ΔPG0382 A Sham 5×106 5×107 5×106 5×107

6 h

F4/80

F4/80 F4/80 F4/80

Ly6C Ly6C Ly6C Ly6C Ly6C

24 h

F4/80

F4/80 F4/80 F4/80

Ly6C Ly6C Ly6C Ly6C Ly6C B

Figure 6.10 Recruitment of inflammatory monocytes in response to P. gingivalis. BALB/c mice were infected intraperitoneally with wildtype P. gingivalis or P. gingivalis ΔPG0382. Mouse peritoneal fluid was harvested 24 h or

48 h-post injection and subjected to flow cytometric analysis. (A) F4/80lo Ly6C+ FACS plots from individual mice. NB: Due to the variation in response between mice, the FACs plots presented do not necessarily represent the “average” response. (B) Quantification of F4/80lo Ly6C+ inflammatory monocytes. Each data point represents an individual mouse (n=5).

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Neutrophils are typically the major responders during the early stages of infection, and mediate bacterial clearance by producing reactive oxygen species and releasing granules containing antimicrobial proteins (e.g. -defensins) (Faurschou and Borregaard, 2003; Rice et al., 1987). Thus, the recruitment of neutrophils in response to P. gingivalis was also examined. Neutrophils express high levels of Ly6G and low levels of Ly6C, and do not express F4/80 (i.e. Ly6G+, Ly6Clo, F4/80-) (Lee et al., 2013). Neutrophil numbers were found to have significantly increased 6 h-post infection with either wildtype P. gingivalis or P. gingivalis ΔPG0382 (Fig. 6.11A-B). As for inflammatory monocytes, infection with 5×107 wildtype P. gingivalis or P. gingivalis ΔPG0382 did not stimulate a further increase in neutrophil numbers (Fig. 6.11). In contrast to inflammatory monocytes, however, the neutrophil response was greatly reduced by 24 h-post infection (Fig. 6.11A-B). Collectively, the results presented in Figure 6.10 and Figure 6.11 suggest that expression of PG0382 by P. gingivalis does not affect the recruitment of inflammatory monocytes and neutrophils.

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P. gingivalis P. gingivalis P. gingivalis ΔPG0382 P. gingivalis ΔPG0382 A 6 7 6 7 Sham 5×10 5×10 5×10 5×10

6 h Ly6G

Ly6G

Ly6G Ly6G

Ly6C Ly6C Ly6C Ly6C Ly6C

24 h

Ly6G

Ly6G Ly6G Ly6G

Ly6C Ly6C Ly6C Ly6C Ly6C

B Figure 6.11 Recruitment of neutrophils in response to P. gingivalis. BALB/c mice were infected intraperitoneally with wildtype P. gingivalis or P. gingivalis ΔPG0382. Mouse peritoneal fluid was harvested 24 h or 48 h-post injection and subjected to flow cytometric analysis. (A) Ly6G+ Ly6Clo FACS plot from individual mice. NB: Due to the variation in response between mice, the FACs plots presented do not necessarily represent the “average” response. (B) Quantification of Ly6G+ Ly6Clo neutrophils. Each data point represents an individual mouse (n=5).

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6.3 Discussion The TIR domain plays a central role in TLR signalling by acting as a modular, interactive domain to enable the recruitment of specific downstream signalling adaptor proteins (e.g. MYD88 and MAL). Signal transduction initiated by TLRs results in the expression of cytokines and chemokines, which stimulate the activation and recruitment of immune cells to combat infection. P. gingivalis can subvert the host immune response by suppressing TLR activation. For instance, P. gingivalis produces heterogeneous forms of LPS with different acylation patterns that inhibit or weakly activate TLR4 signalling (Dixon and Darveau, 2005; Reife et al., 2006). By contrast, bacterial Tcps are proposed to interact with TLR adaptor proteins to attenuate signalling (Patterson and Werling, 2013; Rana et al., 2013). The bioinformatics analyses and functional biochemical experiments performed in Chapter 5 suggested that P. gingivalis PG0382 possesses characteristics of a bacterial Tcp. Therefore, this Chapter examined a potential immunomodulatory role for PG0382.

An isogenic P. gingivalis PG0382-deficient mutant was generated in this study to determine whether PG0382 can modulate the host immune response by subverting TLR signalling. The phenotype of P. gingivalis ΔPG0382 were first examined. The mean generation time of P. gingivalis ΔPG0382 (approximately 3 h) was comparable to wildtype P. gingivalis, suggesting that the PG0382 gene is not important for P. gingivalis growth. Like wildtype P. gingivalis, the P. gingivalis ΔPG0382 mutant grew as black-pigmented colonies when cultured on horse-blood agar. P. gingivalis expresses extracellular proteases (e.g. Kgp and RgpA/B gingipain proteases) that degrade haemoglobin. The subsequent accumulation of µ-oxo bishaem-containing pigments accounts for the black pigmentation of P. gingivalis colonies (Zambon, Reynolds and Slots, 1981; Smalley et al., 1998). In addition to normal pigmentation, cryo-EM revealed that P. gingivalis ΔPG0382 also exhibits a normal EDSL. Although the exact protein components of the EDSL are still unclear, the reduced EDSL of the P. gingivalis gingipain protease-deficient mutant, P. gingivalis W50ABK, suggests that Kgp and RgpA/B are required for the formation of the EDSL (Chen et al., 2011; Gorasia et al., 2015). Accordingly, the absence of black pigmentation and EDSL can indicate impaired gingipain protease activity, or disrupted transportation and/or attachment of gingipain proteases to the outer cell membrane of P. gingivalis. The importance of PG0382 for gingipain protease activity was also directly tested, and P. gingivalis ΔPG0382 was found to retain wildtype levels of Kgp and RgpA/B activity. The gingipain proteases play a central role in the dysregulation of the host immune response by P. gingivalis. Consequently, a different host immune response to P. gingivalis ΔPG0382 would not be attributable to altered gingipain protease activity.

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Studies have shown that P. gingivalis can regulate TLR-mediated cytokine responses in different cell types, including oral epithelial cells (Hajishengallis et al., 2008; Lu et al., 2009; Maekawa et al., 2014). Therefore, the stimulation of inflammatory gene expression in oral epithelial cells (e.g. OKF6 cells) by P. gingivalis ΔPG0382 was investigated. The findings presented above suggest that PG0382 does not function to repress the inflammatory response (e.g. TNF and IL-36G expression) of oral epithelial cells towards P. gingivalis. Mammalian TIR domains are intracellular signalling modules present in TLRs and adaptor proteins (e.g. MYD88 and MAL). As such, bacterial Tcps need to be translocated into the host cell cytoplasm to exert inhibitory activity on TLR signalling. In the case of B. melitensis, TcpB was shown to be delivered into the cytoplasm of RAW 264.7 cells, possibly via a type IV secretion system (Salcedo et al., 2013). TcpC has been shown to be secreted by E. coli CFT037 and then taken up by macrophages (e.g. mouse bone marrow-derived macrophages) (Cirl et al., 2008). While S. aureus TirS was detected in cell culture medium, its mechanism of secretion is not known (Askarian et al., 2014). The mechanisms of secretion and host cell entry of other bacterial Tcps, including TlpA from S. enterica and YpTdp from Y. pestis, are yet to be elucidated. P. gingivalis has a type IX secretion system, which is involved in the extracellular secretion of proteins with a C-terminal domain (CTD) (Nakayama, 2015; Seers et al., 2006; Slakeski et al., 2011). This system is unlikely to mediate the secretion of PG0382 because it does not have a CTD. Although the precise mechanism for the delivery of the P. gingivalis SerB phosphatase into host cells is not known, SerB has been shown to be secreted as well as produced intracellularly following the invasion of gingival keratinocytes by P. gingivalis (Takeuchi et al., 2013). The efficient invasion of gingival epithelial cells by P. gingivalis is dependent on its fimbriae (Yilmaz et al., 2002, 2003). P. gingivalis W50 is an afimbriated strain and appears to exhibit reduced invasion efficiency, relative to other P. gingivalis strains (Duncan et al., 1993; Watanabe et al., 1992). Consequently, the ability of PG0382 to inhibit TLR signalling in oral epithelial cells might largely depend on the secretion and subsequent uptake of PG0382. Additional studies, including single-cell analysis, will therefore be required to determine whether PG0382 is secreted by P. gingivalis, and if so, whether it is delivered into the cytoplasm of oral epithelial cells where it can interact with TLR adaptor proteins.

P. gingivalis has also been shown to modulate TLR-mediated cytokine responses in macrophages (Hajishengallis et al., 2007; Wang et al., 2007, 2010). Thus, the effects of PG0382 gene deletion on the cytokine responses of RAW 264.7 cells to P. gingivalis were determined. Although the responses elicited by wildtype P. gingivalis and P. gingivalis ΔPG0382 were not statistically different, P. gingivalis ΔPG0382 appeared to consistently stimulate weaker IL-6, CCL2 and IL-10 responses. These findings go against the hypothesised role of PG0382 functioning as an inhibitor of TLR-mediated inflammatory 160 responses. Indeed, they contrast with findings from experiments where RAW 264.7 cells challenged with a TcpC-deficient E. coli CFT037 mutant mounted stronger inflammatory cytokine responses (e.g. TNF and IL-6) when compared to cells challenged with wildtype E. coli CFT037 (Cirl et al., 2008). Moreover, the inhibitory effect of TcpC was shown to be MYD88-dependent, because TcpC suppressed TNF secretion in wildtype mouse bone marrow-derived macrophages but not in MYD88-deficient macrophages (Cirl et al., 2008). Mouse bone marrow-derived dendritic cells have also been found to exhibit increased TNF secretion when challenged with a TcpB-deficient B. melitensis mutant (Salcedo et al., 2013). Interestingly, TcpB has also been shown in vitro to cause restructuring of the endoplasmic reticulum in macrophages to trigger an unfolded protein response (Smith et al., 2013). The stimulation of the unfolded protein response is proposed to support B. melitensis replication by mobilising amino acid transport to support lipid biogenesis. In addition, mice infected with a TcpB-deficient B. melitensis mutant had reduced bacterial burden and colonisation of the liver and spleen, compared to mice infected with wildtype B. melitensis (Radhakrishnan et al., 2009). While PG0382 may have a direct immunomodulatory role, the possibility that PG0382 may affect the inflammatory response indirectly, for instance through the modulation of cell-surface expression of proteins and/or lipids that stimulate the host inflammatory response to P. gingivalis, cannot be excluded. Accordingly, additional functional experiments will be required to understand how PG0382 might modulate the inflammatory response of macrophages to P. gingivalis, or whether PG0382 may possess other functions that extend beyond modulating cytokine responses.

The potential immunomodulatory function of PG0382 was further investigated in a mouse model of peritoneal infection. P. gingivalis infection stimulated an increase in the numbers of inflammatory monocytes (albeit not statistically significant due to variation between mice) and neutrophils. Interestingly, the numbers of activated macrophages did not appear to increase following P. gingivalis infection. Intraperitoneal injection of heat-killed P. gingivalis has previously been reported to cause a decline in the numbers of activated macrophages in the first 24 h, followed by an increase in macrophage numbers that peaked 5 days-post infection (Lam et al., 2014). The reduction in macrophage numbers has been coined the “macrophage disappearance reaction”, which is attributable to the adhesion of macrophages to the peritoneal lining following their activation by inflammatory stimuli (Barth et al., 1995). Notably, peritoneal infection with wildtype P. gingivalis and P. gingivalis ΔPG0382 induced comparable increase in the numbers of inflammatory monocytes and neutrophils. The effects of other bacterial Tcps on the recruitment of immune cells in vivo have yet to be directly investigated. However, analysis of skin lesions resulting from cutaneous infection of mice with a S. aureus TirS-deficient mutant revealed increased levels of myeloperoxidase activity and inflammatory gene 161 expression (e.g. IL-1, IL-6 and CXCL1), relative to mice infected with wildtype S. aureus (Patot et al., 2017). This suggests that TirS may play a role in suppressing immune cell (e.g. neutrophil) infiltration and/or activity. In addition, results from an in vitro study with a B. melitensis TcpB-deficient mutant suggest that TcpB can interfere with dendritic cell maturation by inhibiting TLR2 signalling (Salcedo et al., 2008). Although P. gingivalis ΔPG0382 stimulated comparable recruitment of inflammatory monocytes and neutrophils as wildtype P. gingivalis, there is a possibility that P. gingivalis ΔPG0382 may compromise their activity (e.g. phagocytosis and nitric oxide production). Therefore, it would be worthwhile investigating whether there are any differences in the kinetics of immune cell recruitment and activity between the two groups of cells isolated from mice infected with wildtype P. gingivalis or P. gingivalis ΔPG0382 in a comprehensive model with an extended time-course.

In conclusion, the results presented in this Chapter indicate that deletion of the PG0382 gene does not affect the growth rate, EDSL or gingipain protease activity of P. gingivalis. Furthermore, the in vitro assays performed suggest that PG0382 does not modulate the inflammatory response of oral epithelial cells towards P. gingivalis. Intriguingly, PG0382 may in fact enhance cytokine responses in macrophages; however, further studies will be required to determine whether this is a direct or indirect effect. It is also unclear at this stage whether PG0382 is delivered into host cells, and how the delivery process may impact on the ability of PG0382 to modulate host inflammatory responses. Peritoneal infection of mice with P. gingivalis stimulated the rapid recruitment of neutrophils and inflammatory monocytes. The same response was seen when mice were infected with the P. gingivalis PG0382-deficient mutant. Thus, further in vivo studies, including the mouse model of P. gingivalis-induced periodontitis, might provide insight into whether PG0382 plays a role in modulating the host immune response to P. gingivalis, and hence contributes to the progression of chronic periodontitis.

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General Discussion

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7.1 Summary This thesis explored two facets of the interaction between the host and P. gingivalis, including their effects on the host immune response. Specifically, the regulation of the orphan chemokine, CXCL14, in oral epithelial cells by P. gingivalis illustrates how the interplay between host- and pathogen-derived factors can potentially modulate the host immune response. CXCL14 was shown to exhibit bactericidal activity against health- associated Streptococcus species, whilst P. gingivalis was largely resistant to killing by CXCL14. This thesis also identified and investigated a potential P. gingivalis TIR domain-containing protein (Tcp), namely PG0382. P. gingivalis PG0382 shares similar sequence and structural characteristics with TLR adaptor proteins and bacterial Tcps. Moreover, PG0382 was demonstrated to exert a negative impact on the TLR adaptor proteins MAL and MYD88. A PG0382-deficient P. gingivalis mutant (P. gingivalis ΔPG0382) was created to investigate how PG0382 might influence the host immune response to P. gingivalis. Interestingly, P. gingivalis ΔPG0382 stimulated a weaker cytokine response in macrophages. Studies in mice revealed that peritoneal infection with P. gingivalis ΔPG0382 stimulated a similar innate immune response as wildtype P. gingivalis. Taken together, this thesis has identified and defined novel interactions between host-derived and pathogen-derived factors in modulating the host immune response. In particular, it has provided a molecular basis for exploring potential roles for CXCL14 and P. gingivalis PG0382 in the development and progression of chronic periodontitis.

7.2 Implications of a dysregulated CXCL14 response for chronic inflammation and microbial dysbiosis The expression of chemokines by oral epithelial cells is important for the recruitment of immune cells to prevent or clear infection. In this study, oral epithelial cells (e.g. OKF6 cells) were shown to upregulate the expression of CXCL14 in response to P. gingivalis. The stimulation of CXCL14 expression was primarily mediated by the gingipain proteases produced by P. gingivalis, and was dependent on the host protease-activated receptor, PAR-3. Significantly, the activation of MEK-ERK1/2 signalling by epidermal growth factor (EGF) was shown to suppress CXCL14 gene transcription. However, the gingipain proteases can antagonise the EGF-mediated suppression of CXCL14 by proteolytically degrading EGF. P. gingivalis can therefore not only directly stimulate CXCL14 expression via PAR-3 but also promotes its expression by antagonising EGF signalling. Accordingly, this is likely to result in a dysregulated CXCL14 response in the context of P. gingivalis infection/colonisation. Furthermore, the ability of the gingipain proteases to degrade CXCL14 would likely result in a suboptimal CXCL14 response, whereby the impaired recruitment of CXCL14 target cells may compromise the immune response to P. gingivalis. Importantly, the gingipain proteases associated with outer-membrane vesicles (OMVs) 164 released by P. gingivalis may also provide a mechanism to dysregulate CXCL14 expression and/or activity at sites distant from the tooth-accreted biofilm. Although CXCL14 is expressed by oral epithelial cells, it may possess context-dependent functions based on its localised expression at specific sites in the oral mucosal epithelium (e.g. junctional epithelium vs. sulcular epithelium), and thus this aspect should be taken into consideration when further establishing a role for CXCL14 in chronic periodontitis.

This is the first study to demonstrate PAR3-dependent stimulation of CXCL14 expression. Other bacterial species, such as the respiratory pathogen Pseudomonas aeruginosa, have been demonstrated to exert agonistic and antagonistic effects on the host immune response by differentially modulating PAR signalling. For example, the P. aeruginosa extracellular protease, LepA, was shown to activate NF-B in bronchial epithelial cells by stimulating PAR-1, PAR-2 and PAR-4 signalling (Kida et al., 2008), whereas the extracellular LasB protease produced by P. aeruginosa can degrade the extracellular domain of PAR-2 to prevent receptor signalling (Dulon et al., 2005). As such, it would be interesting to determine whether proteases produced by other oral bacterial species (e.g. T. denticola) can also regulate CXCL14 via PAR-3. Accordingly, localised expression of CXCL14 within specific sites in the oral mucosa (e.g. junctional epithelium) might be dependent on the protease milieu.

The findings presented in this thesis indicate that CXCL14 is bactericidal against certain oral bacteria (e.g. S. gordonii). However, P. gingivalis is resistant to killing by CXCL14 killing, most likely because the gingipain proteases can degrade CXCL14. CXCL14 is constitutively expressed in various mucosal tissues, including in the oral epithelium, and hence has been proposed to also play a role in homeostatic defence. For instance, CXCL14 may function as a mediator of cutaneous antimicrobial defence to prevent infections that might otherwise arise following minor skin abrasions (Frick et al., 2011). CCL28 is another homeostatic chemokine, and is constitutively expressed by the epithelial cells of the salivary glands (Hieshima et al., 2003). Like CXCL14, CCL28 has been shown to have bactericidal activity against periodontal pathogens, including P. gingivalis and A. actinomycetemcomitans (Watkins et al., 2007). CXCL14, and other homeostatic chemokines, are therefore likely to have important roles in maintaining periodontal health. Importantly, oral commensal species can participate in maintaining periodontal health by acting as antigenic stimulants to facilitate the activity of an effective non-destructive inflammatory barrier against potential pathogens. The stimulation of hBD-2 expression in human oral epithelial cells by F. nucleatum has been proposed to be important for maintaining periodontal tissue homeostasis by inhibiting the overgrowth of the tooth-accreted biofilm (plaque) (Ghosh et al., 2018). In contrast, P. gingivalis appears

165 to stimulate a weaker hBD-2 response and thereby potentially compromise host defence against the biofilm (Ghosh et al., 2018; Lu et al., 2009). The role of oral commensal species in regulating the expression of homeostatic chemokines, including CXCL14, has yet to be investigated. Therefore, the ability of oral commensal species (e.g. S. mitis) and pathogens (e.g. A. actinomycetemcomitans) to regulate CXCL14 may provide further important insight into the role of CXCL14 in maintaining tissue homeostasis.

The potential “double-edged” nature of host immunomodulatory factors (e.g. cytokines and chemokines) can make it difficult to clearly define their role as being host-protective and host-destructive when dysregulated. Although typically protective, the persistence of innate immune cells (e.g. neutrophils) due to unresolved infection can be highly detrimental to the host. CXCL14 exerts pleiotropic chemotactic activity towards various innate immune cells (e.g. neutrophils, macrophages and dendritic cells) (Shellenberger et al. 2004; Shurin et al. 2005; Kurth et al. 2001; Cao et al. 2000). Thus, the dysregulation of CXCL14 by P. gingivalis might contribute to the development of chronic inflammation by promoting dysregulated immune cell recruitment, and thereby promote oral dysbiosis. CXCL14 bactericidal activity may also contribute to microbial dysbiosis. For example, appropriately regulated CXCL14 expression might be important for limiting the growth of susceptible bacterial species in the tooth-accreted biofilm. However, the ability of the P. gingivalis gingipain proteases to proteolytically degrade CXCL14 may enable P. gingivalis to not only protect itself, but also protect closely associated bacterial species (e.g. accessory pathogens) in the biofilm, which might otherwise be susceptible to killing by CXCL14. The Gram-negative oral pathogen, P. intermedia expresses the protease, interpain A, which is homologous to the Streptococcus pyogenes protease, speB. Significantly, speB has been shown to protect S. pyogenes from CXCL14-mediated killing by degrading CXCL14 (Frick et al., 2011). Therefore, proteases produced by other oral bacteria may also be able to compromise CXCL14 activity. Taken together, the dysregulation of CXCL14 may potentially contribute to chronic periodontitis by promoting chronic inflammation and oral dysbiosis.

The findings from this thesis have provided important molecular insights into the regulation and role of CXCL14. However, the regulation and role of CXCL14 also needs to be considered in a broader context in vivo. Experimental mouse model studies could provide pre-clinical evidence with respect to the relevance of CXCL14 in chronic periodontitis. Periodontal inflammation and alveolar bone loss are key indicators of disease progression in chronic periodontitis. Therefore, the potential contribution and role of CXCL14 in the development of chronic periodontitis could be investigated with CXCL14-deficient mice in the P. gingivalis-induced mouse model of periodontitis. In

166 addition to assessing the impact on alveolar bone resorption, analysis of the oral microbiome, for example, by next generation sequencing, might also provide further insight into the role of CXCL14 in promoting host-microbe homeostasis, and specific consequences of its dysregulation by P. gingivalis.

7.3 CXCL14 in tissue regeneration Current treatment for chronic periodontitis involves scaling and root planing to remove plaque (Pihlstrom et al., 2005). This results in at least the temporary resolution of inflammation and attenuation of disease progression (Cobb, 2002). Antibiotics are sometimes employed as an adjuvant to inhibit biofilm growth when scaling and root planing are not sufficient to resolve inflammation and prevent disease progression (Jepsen and Jepsen, 2016; Slots and Rams, 1990). Stem cell transplantation is being explored as a therapeutic avenue to promote the repair of the periodontium in chronic periodontitis (Chen et al., 2012). Notably, CXCL14 was recently suggested to be a trophic factor produced by dental pulp-derived mesenchymal stem cells (Hayashi et al., 2015). In particular, CXCL14 was proposed to facilitate tissue regeneration, in an ectopic mouse model of tooth root transplantation, by promoting endogenous cell migration (Hayashi et al., 2015). However, the target migratory cells in which CXCL14 might act to promote tissue regeneration were not identified. The results from this thesis suggest that CXCL14 does not regulate the migration of oral epithelial cells, or at least OKF6 cells in an in vitro setting. Interestingly, a study by Shellenberger et al. suggests that CXCL14 can inhibit angiogenesis by blocking IL-8-mediated endothelial cell migration (Shellenberger et al., 2004). This may have implications in regeneration therapy, as angiogenesis is crucial for tissue regeneration. Therefore, further studies will be required to definitively establish whether CXCL14 is a tropic factor, and hence how it could potentially be therapeutically exploited to promote tissue regeneration in chronic periodontitis.

7.4 A potential role for PG0382 in immune subversion The family of Toll-like receptors (TLRs) share a common modular intracellular TIR domain that initiates downstream signalling to induce changes in gene expression by forming heterotypic TIR-TIR interactions with TLR adaptor proteins (e.g. MAL, MYD88, TRAM and TRIF). Bacterial Tcps are proposed to facilitate bacterial immune subversion by blocking TLR signalling. The bioinformatic analyses performed in this thesis suggest that several P. gingivalis strains express putative Tcps (e.g. PG0382). Through bioinformatic analysis, PG0382 from P. gingivalis W83/W50 was found to share similar properties to TLR adaptor proteins and bacterial Tcps. Significantly, PG0382 caused MAL and MYD88 levels to be markedly reduced when co-expressed in mammalian cells (e.g. HEK293T cells). The mechanism underlying the ability of PG0382 to reduce MAL and MYD88 levels 167 has yet to be determined. However, studies of other bacterial Tcps might provide further insight. TcpB from B. melitensis was shown to promote the ubiquitination MAL, and thereby target MAL for degradation by the proteasome (Sengupta et al., 2010). P. gingivalis has been shown to suppress neutrophil antimicrobial responses by inducing signalling crosstalk between the C5aR and TLR2 to promote the ubiquitination and proteasomal degradation of MYD88 (Maekawa et al., 2014). In addition to being degraded via the proteasomal pathway, cellular proteins can be degraded via the lysosomal pathway (Ciechanover, 2005). Thus, these two pathways could potentially be exploited by PG0382 to reduce MAL and MYD88 levels. This could be investigated with pharmacological agents that can selectively inhibit the proteasomal and lysosomal degradation pathways (e.g. MG132 and chloroquine, respectively).

Recent studies have also reported roles for bacterial Tcps in modulating inflammasome activation (Jakka et al., 2017; Waldhuber et al., 2016). As described in Chapter 1, inflammasome activation is critical for IL-1 maturation. Canonical inflammasome activation is mediated by NLRP1, NLRP3 or NLRC4 (Latz et al., 2013), whereas non-canonical inflammasome activation is mediated by caspase-4 (Kayagaki et al., 2011). Upon binding intracellular LPS, caspase-4 oligomerises and induces caspase-1-mediated IL-1 maturation. TcpB has been shown to interact with caspase-4, resulting in the ubiquitination and degradation by the proteasome. Moreover, TcpB was shown to attenuate IL-1 secretion by macrophages (e.g. THP-1 cells and RAW 264.7 cells) (Jakka et al., 2017). In contrast, TcpC can inhibit canonical caspase-1 processing of IL-1 by inhibiting NLRP3-mediated inflammasome activation (Waldhuber et al., 2016). Significantly, in a mouse model of urinary tract infection, IL-1 levels in the urine of mice infected with wildtype E. coli CFT073 were lower than those infected with a TcpC-deficient mutant. These findings suggest that bacterial Tcps may also execute immunomodulatory activity through TLR-independent signalling pathways. Therefore, it would be interesting to determine whether PG0382 can interact and modulate the functions of other immune signalling molecules, including specific components of inflammasome pathways.

Intriguingly, instead of heightened inflammatory cytokine responses, RAW 264.7 cells appeared to mount weaker responses when challenged with an isogenic PG0382-deficient P. gingivalis mutant, suggesting that PG0382 may promote a pro-inflammatory response. This contrasts with other bacterial Tcps, which have shown to exert anti-inflammatory effects on the host immune response (Cirl et al., 2008). P. gingivalis ΔPG0382 has normal Kgp and Rgp activity, and therefore the apparent reduced cytokine activity in RAW 264.7 cells challenged with P. gingivalis ΔPG0382 is unlikely to be attributable to impaired gingipain protease activity. In addition to the gingipain proteases, P. gingivalis produces an 168 array of surface-associated virulence factors (e.g. haemagglutinins) (Holt et al., 1999; How et al., 2016), and therefore changes to the expression of surface-associated proteins could potentially affect the host inflammatory response. The ability of PG0382 to modulate, either directly or indirectly, the expression and/or function of P. gingivalis surface-associated proteins has yet to be elucidated. It will therefore be important to analyse the proteomic profiles of different cellular compartments of P. gingivalis ΔPG0382 (e.g. outer membrane and periplasm), for example by applying mass spectrometry-based approaches (Gorasia et al., 2015).

A mouse model of peritoneal infection was used to assess the potential role of PG0382 in modulating the innate immune response to P. gingivalis. However, the absence of PG0382 expression by P. gingivalis did not appear to affect the responses elicited. The manipulation of TLR signalling by P. gingivalis can result in reduced macrophage and neutrophil activity (Maekawa et al., 2014; Wang et al., 2010). To that end, it would be interesting to determine whether PG0382 can affect the antimicrobial functions of the immune cells recruited, for instance, macrophage phagocytic activity and reactive oxygen species production by neutrophils. Although the peritonitis model is useful for investigating immune cell recruitment, it does not represent the natural course of P. gingivalis colonisation/infection and proliferation that occurs in chronic periodontitis. Thus, the P. gingivalis-induced mouse model of periodontitis might provide more relevant insight into the potential role of PG0382 in virulence.

7.5 The TIR domain as a primordial microbial signalling module Many proteins involved in regulating innate immunity (e.g. TLRs and NLRs) have a modular structure comprising different domains. In addition to the TIR domain, other examples include LRR, ATPase/NTPase and NACHT domains (Dunin-Horkawicz et al., 2014; Koonin and Aravind, 2002). These protein domains often have homologs of unknown function in bacteria (Koonin and Aravind, 2002). The bioinformatic searches performed in Chapter 5 revealed that there are eleven putative P. gingivalis Tcps, and some P. gingivalis strains were found to have two or more Tcps. The propagation of P. gingivalis Tcps between strains may have occurred through DNA exchange by natural competence and conjugation (Tribble et al., 2007). Other bacterial Tcps are largely found in genomic regions within phage origins, and thus their exchange between bacteria was likely to have occurred through horizontal gene transfer events (Zhang et al., 2011). Interestingly, cluster analysis suggests that protein domains associated with mammalian innate immune signalling components, including TIR and NACHT domains, are of bacterial origin, and were acquired through mitochondrial endosymbiosis and additional horizontal gene transfer events (Koonin and Aravind, 2002). Therefore, the likely bacterial origin of

169 the TIR domain suggests that not all bacterial Tcps have evolved to mediate the subversion of the host immune system.

The TIR domain has a broad phylogenetic distribution and widespread within the genomes of microorganisms, including many that are not pathogenic. For instance, there is an over-representation of TIR domains within cyanobacteria (e.g. Anabaena variabilis and Nodularia spumigena), which rarely engage in immune subversion (Spear et al., 2009). Although experimental evidence points to an immune subversive role for at least some bacterial Tcps, it is important to acknowledge that in vitro systems may not always accurately reflect the role of bacterial Tcps in immune subversion. For instance, YpTdp from Y. pestis was shown to bind MYD88 and inhibit LPS-stimulated NF-B activation in an in vitro expression system (e.g. HEK293T cells). However, YpTdp was not required for Y. pestis virulence in mice (Spear et al., 2012). Therefore, in vivo experiments are crucial to determine the importance of bacterial Tcps in causing disease.

Most studies exploring bacterial Tcps have largely focused on their potential function as virulence factors. However, bacterial Tcps often contain other functional domains, and therefore it is possible that they have evolved to mediate specific physiological purposes. Specifically, given that the TIR domain is a modular interactive domain, it is possible that bacterial Tcps engage in protein-protein interactions to mediate intrinsic physiological processes. The identification of multiple Tcps in the same strain of P. gingivalis, for instance, PG0382 and PG1864 in P. gingivalis W83/50, and A343_0215 and A343_1154 in P. gingivalis JCVISC001, therefore raises the possibility that the Tcps might form both homotypic and/or heterotypic interactions through their TIR domains. The generation of a complemented P. gingivalis ΔPG0382 strain in which the re-expressed PG0382 protein is tagged with an appropriate epitope (e.g. V5 epitope) would be useful for not only studying the localisation of PG0382 in P. gingivalis but also identifying potential binding partners. Such information would provide greater insight into the likely function(s) of PG0382.

7.6 Bacterial Tcps as therapeutic agents for inflammation In addition to their host-protective roles, TLRs can also contribute to the pathogenesis of infectious and/or inflammatory diseases. The subversion of TLR signalling by Mycobacterium tuberculosis contributes to pathology in tuberculosis (Harding and Boom, 2010), while the ability of P. gingivalis to subvert TLR2 is central to the pathogenesis of chronic periodontitis (Hajishengallis and Lamont, 2014). Manipulating the unique interactions between TLRs and adaptor proteins could therefore potentially be exploited to provide new therapeutic opportunities for treating some diseases (O’Neill et al., 2009). For example, a dominant-negative form of MYD88 was shown in rats to protect the

170 myocardium from tissue damage following ischemia/reperfusion injury by inhibiting NF-B activation (Hua et al., 2005). In addition, chemical compounds (e.g. compound 4a) that mimic the BB loop in the TIR domain of MYD88 have been shown in mice to attenuate IL-1-induced fever (Bartfai et al., 2003). The ability of bacterial Tcps to inhibit TLR signalling might therefore be utilised for therapeutic intervention in TLR-mediated inflammatory diseases. Bacterial Tcps studied thus far for their ability to modulate TLR signalling have been shown to contain coiled-coil motifs (Alaidarous et al., 2014; Cirl et al., 2008; Rana et al., 2011). The addition of an artificial coiled-coil domain to the TIR domain of MYD88 resulted in potent inhibition of TLR signalling (Fekonja, Bencina and Jerala, 2012). Significantly, the TcpC TIR domain was shown to ameliorate inflammation in the mouse model of collagen-induced arthritis (Pasi et al., 2016). The therapeutic effects were attributed to the ability of the TcpC TIR domain to block MYD88 signalling, and thus inhibit pathogenic Th17 cell responses. Therefore, elucidating the mechanism by which bacterial Tcps interact with TLRs and/or TLR adaptor proteins to suppress TLR signalling could potentially be useful for the design of novel therapeutics to inhibit TLR-mediated pathogenic inflammatory responses.

7.7 Conclusion The ecological properties of the oral cavity are complex, dynamic and highly variable between individuals. The interactions between host cells and oral microbiota is important for shaping various microbial communities in the oral cavity. P. gingivalis causes an imbalance in microbial distribution and shifts microbiota homeostasis to dysbiosis, whereby tissue-destructive chronic inflammation ensues. The novel host cell-P. gingivalis interactions identified and explored in this thesis provide further insight into the role of P. gingivalis as a major pathogen in chronic periodontitis. Finally, the findings from this thesis should also be examined in a broader context to provide greater perspective into the potential involvement of CXCL14 and PG0382 in the development of chronic periodontitis.

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Appendix Table A2 Mammalian TIR domains used for protein sequence alignments

Protein Accession number TIR domain TLR1 Q15399 635-779 TLR2 O60603 639-784 TLR3 O15455 754-896 TLR4 O00206 672-818 TLR5 O60602 691-837 TLR6 Q9Y2C9 640-784 TLR7 Q9NTK1 878-1036 TLR8 Q9NR97 878-1025 TLR9 Q9NR96 868-1025 MAL P58753 84-221 MYD88 Q99836 159-296 TRAM Q86XR7 73-232 TRIF Q8IUC6 390-460 SARM Q6SZW1 559-657

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Table A3 Bacterial TIR domain-containing proteins

Species Protein Accession number TIR domain Porphyromonas gingivalis PG0382 I9E6I1 341-490 Brucella melitensis TcpB Q8YF53 120-490 Escherichia coli TcpC G8Z3N0 171-307 Paracoccous dentrificans PdTIR A1AY86 168-258 Yersinia pestis YpTdp Q8CL16 130-285 Staphycoccous aureus TirS ORF020 142-245

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Author/s: Aw, Jiamin

Title: Host-pathogen interactions of porphyromonas gingivalis

Date: 2018

Persistent Link: http://hdl.handle.net/11343/214396

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